the characterization and optimization of an irox-based
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Graduate Studies The Vault: Electronic Theses and Dissertations
2014-09-30
The Characterization and Optimization of an
IrOx-based Glucose Biosensor
Sebastian, Holly, Bri
Sebastian, H. (2014). The Characterization and Optimization of an IrOx-based Glucose Biosensor
(Unpublished doctoral thesis). University of Calgary, Calgary, AB. doi:10.11575/PRISM/28028
http://hdl.handle.net/11023/1868
doctoral thesis
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UNIVERSITY OF CALGARY
The Characterization and Optimization of an IrOx-based Glucose Biosensor
by
Holly Bri Sebastian
A THESIS
SUBMITTED TO THE FACULTY OF GRADUATE STUDIES
IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE
DEGREE OF DOCTORATE OF SCIENCE
DEPARTMENT OF CHEMISTRY
CALGARY, ALBERTA
September, 2014
© Holly, Bri Sebastian 2014
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ABSTRACT
Persons who live with diabetes must continuously monitor their blood glucose levels (BGLs)
throughout the day to ensure their BGLs stay within the normal range. For this reason, glucose
sensors must be extremely reliable, robust, accurate, reproducible and sensitive. However, glucose
sensor designs face obstacles, such as poor accuracy and reproducibility, which is often the result
of O2-dependency and poor biocompatibility. Therefore, the primary focus of this thesis has been
to optimize and characterize a biocompatible Ir oxide (IrOx) nanoparticle (NP)/glucose oxidase
(GOx) based glucose biosensor, with the goal of achieving an accurate and reproducible O2-
independent glucose response.
Two approaches were used for the fabrication of this IrOx-based glucose sensor. In the first,
a sensing ink consisting of three key components, IrOx NPs, GOx and Nafion®, and a
water/ethanol solvent was aliquot-deposited on a Au substrate. In the second method, GOx
molecules were immobilized directly to a Au or carbon substrate by covalently linking GOx to a
diazonium-derived film deposited on the substrate. An aliquot of the ethanolic Ir ink was then
(assumed to be) intercalated into the film. Both films were dried, and the Ir was subsequently
electrochemically oxidized to IrOx.
The first goal of sensor construction using aliquot-deposition was to understand the effect of
each of the ink components on their distribution in the dried films and the interconnectedness
particularly of the IrOx NPs, then establishing the effect of these variables on the glucose response.
The goal of diazonium sensor construction was to optimize the immobilization matrix, and obtain
uniform film morphologies that exhibit sensitive and reproducible O2-independent glucose signals.
IrOx NP films (fabricated from ethanolic inks) exhibited uniform film morphologies, and
rapid electron transfer kinetics, while films fabricated from aqueous or GOx-containing inks were
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aggregated and thus easily dislodged from the Au. The addition of 1 wt. % Nafion® to the ink
enhanced IrOx NP stability and redox kinetics. The GOx regeneration route could be significantly
affected by manipulating the film component ratios. Direct mediation was favored by altering the
GOx:IrOx ratio, as well as the water and Nafion® content in the ink. Film thickness and ink
sonication prior to film deposition also influences the GOx regeneration route, as well as film
reproducibility and sensitivity.
To prevent IrOx aggregation (observed for films fabricated from water or GOx-containing
inks), GOx was immobilized (diazonium chemistry) to the current collector, and the ethanolic Ir
ink was then (assumed to be) intercalated into the film. GOx regeneration took place via the desired
IrOx-mediation, independent of the amount of O2 in solution. By increasing the surface coverage
of the diazonium-derived layer (to a maximal coverage) and the amount of IrOx NPs in the film,
the glucose signal was enhanced, but was not comparable to those obtained using the aliquot
method.
Overall, it is clear that our sensor sensitivity, reproducibility and O2-independence are quite
good relative to other published work, improved here by optimization of the sensing film
morphology and stability Further, the goal of achieving an O2-independent glucose signal was
essentially met by avoiding film aggregation initiated by the non-uniform drying tendency of the
aqueous and ethanolic phases in the ink.
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ACKNOWLEDGEMENTS
I would like to extend my sincerest thanks and deepest appreciation to my supervisor, Dr.
Viola I. Birss, for allowing me the opportunity to belong to such an exceptional research team and
for guiding and supporting me through my project, and some of the most important years of my
life. Her patience, meticulous editing and enthusiasm for electrochemistry was crucial to my
success. Thank you for doing your part in shaping who I am and who I will someday be.
This thesis would not have been possible without the assistance of my supervisory
committee members Drs. David Cramb and Venkataraman Thangadurai. Their stimulating
discussions during our meetings, and my final defense, were very valuable and very much
appreciated. I would also like to thank the other members of my defence committee, Drs. Mark
McDermott and Matthias Amrein for a very pleasant defence experience.
My gratitude also extends to Dr. Hanna Elzanowska (University of Warsaw), who
mentored me at many stages of this project, providing insightful discussions and a wonderful
friendship. Her guidance helped drive this project while I was a summer student, which eventually
led me to completing my doctorate.
Financial assistance is appreciated from the Department of Chemistry at the University of
Calgary, as well as the Natural Sciences and Engineering Research Council of Canada (NSERC).
I would also like to thank the helpful Chemistry support staff, including Mark Toonen (Glass
shop), Ed Cairns and his summer student Duncan (Electronics shop), and finally Jan Crawford and
Bonnie King in the graduate student office, for their tireless efforts, as without them the completion
of my project would not have been possible. I would also like to express my appreciation to Dr.
Tobias Fürstenhaupt for his insightful suggestions and explanations during my time at the
Microscopy & Imaging Facility.
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I wish to thank both the Birss and the Thangadurai group who I had the honor of sharing
both an office and laboratory space with. I am indebted to them for their support, friendship, and
insightful discussions. I would like to especially mention those who played a close role in my
research, helping prepare me for countless presentations and helpful suggestions for future steps.
Thank you to Dr. Scott Paulson, Corie Horwood, Anusha Abhayawardhana, Dr. Ehab El-Sawy,
Dr. Hany El-Sayed, Dr. Enam Alsrayheen, Dr. Dustin Banham, Dr. Samar Gharaibeh, Dr. Jason
Young, Robert Mayall, Samuel Aquino, Farisa Forouzandeh, Xiaoan Li, and Natsumi Yokura.
These exemplary individuals went out of their way to give experimental advice, help analyze data,
prepare for oral presentations or edit and critique certain aspects of my thesis.
A very special thanks to all my friends whom I cherish, Ken, Steph, Theresa, Keenan, Lisa,
Nikki, Mel, Kat and Pam for being there for me during my years of schooling. A countless number of
thanks to my family (Mom and Dad, Logan and Keifer, Margaux and Terry, Grandpa and Grandma
Porth, Grandpa and Grandma Sebastian, Grandma Butler, and Barb and Ray) for loving and supporting
me through all of these years, and being there for me when I needed them the most (and even when I
didn’t). To my Mom and Dad, in particular; you always told me that my smarts were 99% genetic and
I wouldn’t have been nearly as smart (or cheeky) if you two were not my parents. As such, I must pay
tribute to and acknowledge how incredibly thankful I am that you two raised me to be who I am today.
P.S. I am sure that sometimes Viola wishes you had put a little bit more effort into teaching me
grammar (this would likely have cut down on her editing time). This is only speculation, of course!
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TABLE OF CONTENTS
ABSTRACT………………………………………………………………………………….......ii
ACKNOWLEDGEMENTS………………………………………………………………….…iv
TABLE OF CONTENTS…………………………………………………………………….…...vi
LIST OF TABLES……………………………………………………………………………….xi
LIST OF FIGURES AND SCHEMES……….....………………………………………….……xiii
LIST OF ABBREVIATIONS…………………………………………………………………...xx
LIST OF SYMBOLS…………………………………………………………………….…….xxiii
CHAPTER ONE: INTRODUCTION………………………………………………...………..….1
1.0 Project Background……………………………………………...………………………….....1
1.1 Research objectives……………………….…………………………………………………...3
1.2 Organization of thesis……………………………….……………………………………...….3
CHAPTER TWO: RELEVENT BACKGROUND……………………………………………..…6
2.0 Introduction to glucose biosensors…………………………………………………….….…...6
2.1 Development of glucose sensors for the management and monitoring of diabetes……………6
2.1.1 Types of biosensors………………………………….………………………...……..6
2.1.2 Glucose sensors in clinical trials and currently on the market……………………...8
2.1.3 Existing challenges of glucose biosensors…………………………………….…....11
2.2 Glucose biosensor components……………………………………………………………....12
2.2.1 Glucose sensing enzymes…………………………………………………….….….12
2.2.1.1 Enzyme immobilization techniques…………………………………….…14
2.2.1.2 Diazonium salt chemistry………………………………………………...16
2.2.1.3 Thiol-derived Self Assembled Monolayers……………………..…………17
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2.2.2 Enzyme immobilization matrices…………………………………………………...18
2.3 Four generations of amperometric glucose biosensors……..…………………………….…..20
2.4 Iridium oxide/Nafion®/GOx-based glucose biosensor components……...…………………..23
2.4.1 Ir and Ir oxide nanoparticles……………………………………………...…….…23
2.4.2 The role of Nafion® in sensing films…………………….……………………….….27
2.4.3 Glucose Oxidase…………………………………………………..…………….….29
2.4.4 Early glucose sensing work in the Birss group…………………………………….30
2.5 Enzyme kinetics………………………………………………………………………...……31
2.6 Techniques used in this work…………………………………………………..…………….35
2.6.1 Quartz crystal microbalance (QCMB) technique………………….………………35
2.6.2 Cyclic voltammetry and amperometry…………………………..…………………36
2.6.3 Scanning and transmission electron microscopy………….……….………………37
2.6.4 X-Ray photoelectron spectroscopy………………………………….……………...38
2.6.5 X-Ray diffraction…………………………………………..……………………….40
CHAPTER THREE: EXPERIMENTAL METHODS…………………………………………...41
3.1 Preparation of Ir sols and sensing inks…………………………………...…………………..41
3.1.1 Preparation of ethanolic Ir sol-based inks……………………………………..……..…41
3.1.2 Preparation of Ir/Nafion®/GOx sensing film via aliquot deposition…………..…..41
3.1.2.1 Preparation of GOx solution………...…………………………………...41
3.1.2.2 Preparation of Nafion® solution…………………...……………………..41
3.1.2.3 Aliquot deposition of sensing ink on Au substrate…………….……….…42
3.1.3 Sensing film fabrication using diazonium salt chemistry……………….………….42
3.1.3.1 Diazonium salt grafting and GOx immobilization………………………..42
3.1.3.2 Infiltration of IrOx/Nafion® into immobilized GOx layer…………..….…43
3.2 Electrochemical characterization…………………………………………………………….44
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3.2.1 Working electrodes (WEs)………………………………………………………….44
3.2.1.1 Au working electrodes………………………..…………………………..44
3.2.1.2 Carbon working electrodes……………………………………...……….45
3.2.1.3 Electrochemical cleaning of working electrodes……………………...…45
3.2.1.4 Real Ir and Au surface area determination……………………...……….46
3.2.2 Counter Electrodes (CE)……………………………………………..…………….47
3.2.3 Reference Electrodes (RE)…………………………………………………………47
3.2.4 Electrochemical methods…………..………………………………………………48
3.2.4.1 Electrochemical cell…………………...…………………………………48
3.2.4.2 Solutions………………………………………………………………….50
3.2.4.3 Environment……………….……………………………………………..50
3.2.4.4 Electrochemical instrumentation and software…………………………..51
3.2.5 Glucose addition experiments……………………………………………….……..51
3.3 Other techniques……………………………………………………………………………...51
3.3.1 Transmission electron microscopy (TEM)………………………..………………..51
3.3.2 Scanning electron microscopy (SEM).………………………………………..……52
3.3.3 Quartz Crystal Microbalance (QCMB) technique…………………………………52
3.3.4 X-Ray photoelectron spectroscopy (XPS)……………………………...…………..54
3.3.5 X-Ray Diffraction (XRD)…………………………………………………………...54
3.4 Error analysis……………………..…………………………………………………………..54
CHAPTER FOUR: UNDERSTANDING IRIDIUM-BASED INK
DRYING AND ELECTROCHEMICAL PROPERTIES…………………………………….….57
4.1 Introduction…………………………………………………………………………………..57
4.2 Characterization of IrOx nanoparticles synthesized from 100% ethanolic sols……….….….59
4.3 Effect of % water in Ir ink…………………………………………………….………….…..65
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4.3.1 Effect of water content of Ir inks on ink and film properties after drying……….…65
4.3.2 Effect of water content Ir ink on subsequent IrOx film properties………………….68
4.4 Effect of Nafion® in Ir inks…………………………………………………………….…….72
4.4.1 Effect of 1 wt. % Nafion® in Ir ink on Ir film morphology………….………………72
4.4.2 Effect of Nafion® content in ink on subsequent IrOx electrochemistry………….....73
4.5 Effect of glucose, H2O2 and gluconolactone in solution on the IrOx electrochemistry……...77
4.5.1 Effect of glucose in the cell solution on IrOx redox kinetics……………………….77
4.5.2 Effect of H2O2 in the cell solution on IrOx redox kinetics…………………………..78
4.5.3 Effect of gluconolactone on IrOx kinetics…………………………………….….…80
4.6 Summary………………….………………………………………………………….………81
CHAPTER FIVE: EFFECT OF SENSOR PREPARATION ON
DUAL-MEDIATION SENSING OF GLUCOSE…………………..…………………………...83
5.1 Introduction………………………………………………..…………………………………83
5.2 Dual-mediation glucose response of IrOx/Nafion®/GOx films…………………………...…85
5.2.1 Typical glucose signal from an IrOx/Nafion®/GOx
sensing film in aerated and deaerated solutions..………………………………………..85
5.2.2 Effect of potential used for glucose sensing on glucose signal………….…………87
5.3 Physical and Electrochemical Characterization of Ir/GOx films………………...…………..89
5.4 Effect of % water in GOx-containing inks…………………………………..……………….94
5.4.1 Effect of % water content on GOx activity…………………………………………94
5.4.2 Effect of % water on IrOx and GOx–containing films………….……….…………98
5.4.3 Effect of % water in Ir oxide/GOx ink on glucose detection……………….………100
5.5 Effect of GOx:Ir mass ratio on glucose sensing………………………………………...…..104
5.6 Effect of Nafion® on glucose sensing films……………………………………...…………108
5.7 Summary……………………………………...…………………………………………….111
CHAPTER SIX: TOWARDS A RELIABLE AND HIGH SENSITIVITY
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O2-INDEPENDENT GLUCOSE SENSOR BASED ON IR OXIDE NANOPARTICLES…….113
6.1 Introduction……………………………………………………………………………...….115
6.2 Results and discussion……………………………………………………….……………...115
6.2.1 Indicators of direct electron transfer and regeneration of GOx………………….115
6.2.1.1 Effect of O2 on glucose signal……………..……………………………115
6.2.2 K’m and imax values as an indicator of GOx regeneration mechanism…………...117
6.3 Film fabrication methods to enhance O2-independence…………………...…………….….121
6.3.1 Effect of film thickness on O2-independence and sensitivity…………………….….121
6.3.2 Effect of sonication of ink on O2-independence……………………………….….125
6.3.3 Spin-coat deposition of sensing films………………………………………….….127
6.4 Summary……………………………………………………………………………………129
CHAPTER SEVEN: DIAZONIUM- IMMOBILIZATION OF GOX IN AN IRIDIUM OXIDE-
BASED GLUCOSE BIOSENSOR………………………………………………………….….132
7.1 Introduction………………..………………………………………………………………..132
7.2 Modification of Au with diazonium salt chemistry…………...………………………….…135
7.2.1 Nitrophenyl grafting and subsequent reduction …………..……………………...135
7.2.2 Degree of Au coverage by diazonium-derived layer to
the number of nitrophenyl groups……………………………………………….………139
7.3 GOx attachment to diazonium-derived aminophenyl layer………….……………….……..142
7.4 Glucose testing……………………………………………………………..……………….143
7.4.1 Glucose testing in the absence of IrOx……………………..…………………….143
7.4.2 Glucose testing at IrOx/Nafion®/immobilized-GOx films….……………………..145
7.5 Further enhancement of glucose sensor response………………………..…………………150
7.5.1 Optimization of amount of Ir in the sensing film…………………………………150
7.5.2 Potentiostatic vs. cyclic voltammetry method for grafting of nitrophenyl
groups on Au………………………………………………………………………...….155
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7.5.3 Increasing glucose response by increasing Au surface roughness………….……160
7.6 Glucose sensors based on an immobilized GOx/Ir oxide/Nafion® film on carbon………....163
7.6.1 Deposition of immobilized-GOx/Ir oxide/Nafion® sensing film on
carbon paper substrate……………….………………………………………………....163
7.6.2 Glucose signal from a CP/Ir/Nafion®/immobilized GOx sensing film……….…...166
7.7 Summary…………………………...……………………………………………………….168
CHAPTER EIGHT: CONCLUSIONS AND FUTURE WORK……………………………….170
8.1 Conclusions………………………………...……………………………………………….170
8.1.1 Conditions that effect the stability of Ir NP sol-based inks……………………….170
8.1.2 Conditions that give the most reproducible glucose signals and film
morphologies……………………………………………………………………………171
8.1.3 Conditions that give the overall highest glucose signals……………………...….172
8.14 Conditions required for direct mediation and/or O2-independent sensing…….….173
8.1.5 Global comparison to other glucose biosensors………..…………………….…..174
8.2 Suggestions for Future Work………………………………..…………………………..…..176
REFERENCES…………………………………………………………………………...…….178
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LIST OF TABLES
Table 5.1. MM parameters for thin films formed from inks containing variable vol. % water in
aerated (O2) and deaerated (Ar) solutions (see Figure 5.5). (Concentration of all other components
was constant in a 4:1:1 volume ratio of IrOx: Nafion®: GOx)………………………………..…102
Table 5.2. Comparison of the average Michaelis-Menten parameters (K’m and imax) for
IrOx/Nafion®/GOx films (ca. 2.5 µm thick) with increasing wt. % Nafion® content in deaerated
(Ar) and aerated (O2), 0.1 M, pH 7 phosphate buffer solution………………………………..…110
Table 6.1. Comparison of the Michaelis-Menten parameters (K’m and imax) for three
IrOx/Nafion®/GOx films (ca. 2.5 µm thick) with Nafion® in aerated and deaerated glucose-
containing 0.1 M, pH 7 phosphate buffer solutions……………………………………………..119
Table 7.1. Michaelis-Menten values for glucose signals at immobilized-GOx/IrOx/Nafion®
sensing films in aerated and deaerated environments (Figure 7.7)………………………………154
Table 7.2. Surface coverage of aminophenyl groups in the diazonium-derived film, deposited on
a Au substrate via either CV cycling or potentiostatically, and Michaelis-Menten values for the the
resulting immobilized-GOx/IrOx/Nafion® sensing films in aerated and deaerated solutions…...158
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LIST OF FIGURES AND SCHEMES
Figure 2.1. (a) The general representation of Nafion® showing the fluorinated carbon backbone
and hydrophilic sulfonate side chain. (b) The configuration of Nafion® in a hydrated environment
such as a blood sample, where water/ion transport channels are formed………………………….27
Figure 2.2. Reaction coordinate diagram, depicting how an enzyme decreases the activation
energy of a reaction, allowing for its acceleration, where G = Gibb’s free energy, E = enzyme, S
= substrate, ES = transition state, and P = product……………………………………………..…33
Figure 2.3. (a) The concentration of all of the components in the Michaelis-Menten equation for
an enzymatic reaction with time (b) A typical Michaelis-Menten plot following the rate of reaction
with increasing substrate concentration…………………………….…………………………….34
Figure 2.4 (a) The Au keyhole pattern deposited on a quartz crystal used in the quartz crystal
microbalance technique, and (b) side view of a quartz crystal disk with an exaggerated view of
the shear distortion resulting from the oscillation1……………………………………………… 36
Figure 3.1. Cyclic voltammetry response of (a) an Ir film and (b) a Au film in a deaerated, stirred,
0.5 M H2SO4 solution, at 100 mV s-1. The purple shading indicates the area used to determine the
real surface area, where the area under the cathodic peak should be equivalent to the area under
the anodic peak……………………………………………………………………………….…..47
Figure 3.2. Electrochemical setup used for (a) substrate cleaning, film characterization, and
glucose additions, and (b) diazonium salt reduction and the subsequent reduction of the nitro- to
an amino group………………………………………………………………………………...…49
Figure 3.3. A photograph of the components of the electrochemical/QCMB cell, in the (a)
disassembled and (b) assembled state………………………………………………………….....53
Figure 4.1. (a) Transmission electron microscopy bight field image of ethanolic Ir ink, (b)
histogram of Ir particle size obtained from (a), and (c) scanning electron microscopy image of an
IrOx film obtained using a 20 kV accelerating voltage..................................................................60
Figure 4.2. (a) Cyclic voltammetry (0.01 V s-1) showing the oxidation of an Ir ink (synthesized in
a 100% EtOH medium) in a stirred 0.5 M H2SO4 deaerated with Ar. (b) Decrease in Hupd peak
charge density with increasing number of cycles between 0 and 1.25 V, as in
(a)………………………………………………………………………………………………...61
Figure 4.3. X-Ray Photoelectron Spectroscopy results, showing the (a) 4f7/2 and 4f5/2 and (b) O 1s
spectra, for an electrochemically oxidized thin IrOx film (ca. 15 µm)………………………..…63
Figure 4.4 X-Ray diffraction pattern of Ir NPs from an ethanolic Ir sol loaded on amorphous
glass in the 2θ range of 0°-90°………………………………………………………………...…64
xiv
Figure 4.5. Average evaporation rate (g h-1) as a function of ethanol concentration (% w/w) for
both the ethanol and water components of a mixture (plot reproduced from data compiled by K.D.
O’Hare et al……………………………………………………………………….…………..….66
Figure 4.6. Visual comparison between (a) an ethanolic Ir ink and a 50:50 vol. % H2O:EtOH Ir ink,
and (b) the resulting IrOx films after drying on a sputtered Au substrate……………………………..67
Figure 4.7. (a) CVs of films (10 mV s-1) formed from a pure ethanolic Ir ink (■) and a 50:50 vol
% water: ethanol Ir ink (▲). Electrochemistry was performed in deaerated, stirred, 1.0 M PBS at
a sweep rate of 10 mV s-1. (b) log of peak current vs. log sweep rate and (c) Ep vs. log ν for the
results shown in (a)……………………………………………………………………………….70
Figure 4.8. HR-TEM images of 50 vol. % H2O containing ethanolic Ir ink on Cu-TEM grid. (a)
Outer region of grid where EtOH evaporated quickly, (b) center of grid where Ir NPs aggregated
in the water-rich droplets, and (c) magnified view of (b)……………………………………..….71
Figure 4.9. Optical images of IrOx films prior to the partial electrochemical oxidation of Ir and
after oxidation, followed by the rinsing of the films with water for 5 seconds. Films were fabricated
using 4 different Ir NP inks: ethanolic, ethanolic with Nafion®, a 1:1 ratio of water and ethanol,
and a 1:1 ratio of water and ethanol with added Nafion®. Each film contained 88 µmol Ir and
respective films contained 1 wt. % Nafion®, when Nafion® was present. Electrochemistry was
performed in deaerated, stirred pH 7, 0.1 M PBS………………………………………………..72
Figure 4.10. (a) Cyclic voltammetry (100 mV s-1) for IrOx/Nafion® films fabricated from ethanolic
(no water) inks containing varying wt. % Nafion® (0%, 1%, 2%, 4%, 8%) at 100 mV s-1 in stirred
pH 7, 0.1 M phosphate buffer solution deaerated with Ar. (b) C1 peak current density (ipc) vs. log
sweep rate (ν) for films examined in (a).........................................................................................74
Figure 4.11. (a) Cyclic voltammetry (100 mV s-1) for thin IrOx films fabricated from dilute
ethanolic Ir (1.1 mM) inks containing 1 wt. % Nafion®, 50 vol.% water, or a combination of both
to compare the effect of Nafion® on the IrOx redox kinetics in stirred, deaerated, 0.5 M H2SO4. (b)
log anodic i vs. log ν plot for the films in (a), and (c) Ep vs. log ν plot for the films in
(a)………………………………………………………………………………………………...76
Figure 4.12 Cyclic voltammetry (100 mV s-1) for a fully electrochemically oxidized (formed by
cycling from 0 to 1.45 V) IrOx electrode with increasing concentrations (0-12 mM) of glucose (1.6
mM additions) added to the cell solution in stirred, aerated 0.1 M, pH 7 PBS…………………..78
Figure 4.13. Cyclic voltammograms (0.1 V s-1) of an IrOx/Nafion® film (after electrochemical
conversion of Ir to IrOx), deposited using a 2 µL aliquot of 20% H2O/80% EtOH Ir sol + 0.04 %
Nafion®, deposited on Au in a deaerated, pH 7, 0.1 M phosphate buffer solution. (a) and (b) show
the CV response when aliquots of 3% H2O2 were added to the solution in (a) deaerated and (b)
aerated environments. H2O2 concentrations are 0 mM (solid line), 0.15 mM (dashed line), 0.3 mM
(dotted line) and 0.9 mM (dot dash line). The insets in (a) and (b) show the plots of the H2O2
oxidation (at 1.2 V) and reduction (at 0.2 V) currents vs. [H2O2] in each
environment……………………………………………………………………………………...79
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Figure 4.14. Cyclic voltammetry (100 mV s-1) for a fully electrochemically oxidized (formed by
cycling from 0 to 1.45 V) IrOx electrode with increasing concentrations (0-12 mM) of
gluconolactone (1.6 mM additions) added to the cell solution in stirred, aerated 0.1 M, pH 7
PBS…………………………………………………………………………………………...….81
Figure 5.1. Michaelis-Menten plot (at 1.2 V) in stirred, pH 7 0.1 M, RT, phosphate buffer solution
(25 mL) at room temperature with 20 µL increment additions of 1.0 M glucose under aerated and
deaerated conditions for a film formed from a IrOx/Nafion®/GOx ink. Inset shows the steady state
current reached after the first glucose additions…………………………………………….……86
Figure 5.2 Four IrOx/Nafion®/GOx films tested for their response to glucose at a constant potential
of either 1.05, 1.10, 1.15 or 1.20 V up to 70 mM of glucose in stirred, deaerated pH 7 0.1 M
PBS…………………………………………………………………………………………...….88
Figure 5.3. High resolution transmission electron spectroscopy images of a typical Ir NP/GOx
film that has been diluted with a 20 vol.% water/ethanol mixture by 50x. (a-c) shows magnified
sections of the film, (d) is a magnified view of the Ir NPs aggregated around the GOx molecules,
and (e) shows the Ir NP distribution at a position in the film that is far from the aggregated GOx
molecules……………………………………………………………………..………………….90
Figure 5.4. Typical cyclic voltammetry (10 mV s-1) of an IrOx/GOx (12 g l-1) glucose sensing film
(20 vol. % H2O) that was electrochemically oxidized by cycling the potential from 0 to 1.25 V in
stirred, deaerated 0.1 M pH 7 PBS. Inset shows the log ipa vs. log ν plot for an IrOx NP film and
an IrOx NP/GOx film…………………………………………………………………………….92
Figure 5.5. XPS data for an IrOx/GOx (12 g l-1) sensing film that was electrochemically oxidized
by cycling the potential from 0 to 1.25 V in stirred, deaerated 0.1 M pH 7 PBS………………..93
Figure 5.6. (a) i/t data showing the baseline current ( ̶ ̶̶ ) (with no glucose present) and the current
produced when 150 mM glucose is present (---) in aerated, stirred PBS (with no EtOH present)
containing 1 g l-1 GOx. The increase in current is a due to the oxidation of H2O2 which is generated
when GOx reacts with glucose, and is regenerated by O2. (b) Charge passed under the same
conditions as (a) except the PBS solution was comprised of varying H2O:EtOH ratios. (c) Glucose
signal achieved after enzyme activity was recovered by decreasing the EtOH content of the GOx
solution from 80 vol. % to 20 vol. % (̶ ̶̶). Reference glucose signals achieved when GOx is
dissolved in a 20 vol. % EtOH (**) and an 80 vol. % EtOH (--) solution………………………..96
Figure 5.7. Ir/GOx (before rinsing) and IrOx/GOx (after rinsing) films fabricated from increasing
water-content inks. Films are shown prior to electrochemical oxidation, and after the films were
electrochemically oxidized (converting Ir NPs to IrOx NPs) and rinsed with H2O………………99
Figure 5.8. Michaelis-Menten (MM) plots for ethanol-based IrOx/Nafion®/GOx films, formed
from inks containing 6 g l-1 GOx and 0 (♦), 10 (■), 20 (●), and 50 wt. % (▲) water and tested for
their glucose response in (a) deaerated (with Ar) and (b) aerated, stirred, neutral phosphate buffer
solution. Error bars are the standard deviation of the glucose signal for three sensing films…….101
xvi
Figure 5.9. (a) Michaelis-Menten plots of electrodes composed of GOx/IrOx mixture with mass
ratios of 0.25 g/g (1, ▼), 0.50 g/g (3, ▲), 1.00 g/g (2, ●) and 2.00 g/g (4, ■). Glucose testing was
performed at 1.2 V in a stirred pH 7 0.1 M phosphate buffer solution under Ar-saturated conditions.
Aliquots consisted of 50 µl; of 2 M glucose, and (b) Plot of imax and K’m values as a function of
GOx:Ir mass ratio based on data in (a), with points calculated using Eadie–Hofstee plots…….105
Figure 5.10. Comparison of the charge in the Hupd peaks (prior to electrochemical oxidation of
Ir to IrOx) and the Ir(III)/Ir(IV) oxide anodic peak current (after complete conversion of Ir to IrOx)
for IrOx/Nafion®/GOx films as a function of increasing amounts of GOx (0 to 17.5 g l-1) in the
inks…………………………………………………………………………………………...…107
Figure 5.11. Glucose response for IrOx/Nafion®/GOx films with increasing Nafion® content (0,
1, 2, 4 and 8 vol. %) in (a) deaerated and (b) aerated, stirred, 0.1 M, pH 7 phosphate buffer
solution……………………………………………………………………………………….…109
Figure 6.1. Chronoamperometry plot at 1.2 V for (a) a (nearly) O2-independent glucose sensor
and (b) an O2-dependent glucose sensing film. Glucose additions (50 µL of 2M glucose) were
made to a stirred, pH 7, 0.1M phosphate buffer solution (25 mL) under Ar-saturated conditions for
the first 7 min. The solution was then fully aerated and no notable change in the sensor response
to glucose was seen……………………………………………………………………………..116
Figure 6.2. Chronoamperometric data obtained from a 2.5 µm thick IrOx/Nafion®/GOx film, tested
for its glucose response in an anaerobic and aerobic environment at 1.2 V. The current was measured
after the addition of 20 µL aliquots of 1.0 M glucose to the stirred pH 7, 0.1 M phosphate buffer
solution (25 mL). Inset shows the glucose response after the addition of the first glucose aliquot in both
aerobic and anaerobic environments………………………………………………………………...118
Figure 6.3. Cyclic voltammetry (0.01 V s-1) showing effect of the IrOx/Nafion®/GOx film thickness
(dictated by the size of the aliquot of ink deposited on the Au surface) on IrOx CV response in
deaerated, quiescent, pH 7, 0.1 M phosphate buffer solution. Estimated film thickness is 0.66 µm (1),
1.30 µm (2), 2.7 µm (3) and 4.0 µm (4). Inset: IrOx anodic peak (A1) current density vs. the film
thickness…………………………………………………………………………………………….122
Figure 6.4. Current density response (imax) of IrOx/Nafion®/GOx films of varying thickness to
glucose, plotted against the IrOx CV peak current (peak A1 in Fig. 6.3) in (a) aerated and (b)
deaerated conditions…………………………………………………………………………….124
Figure 6.5. (a) and (b) CVs (0.1 V s-1) of 2 µm thick IrOx/Nafion®/GOx films in deaerated, stirred,
0.1 M, pH 7 phosphate buffer solution and (c) and (d) corresponding Michaelis-Menten plots in
aerated (●) and deaerated (■) solutions. The inks were either sonicated for 2 hours (a) and (c) or
non-sonicated (b) and (d) prior to film formation. In the Michaelis-Menten plots, each point
represents the addition of a 20 µL aliquot of 1.0 M glucose to the phosphate buffer solution (25
mL)……………………………………………………………………………………………...126
Figure 6.6. (a) Optical image of a spin-coated Ir/Nafion®/GOx film on a sputtered Au substrate,
(b) CV (100 mV s-1) of three spin coated IrOx/Nafion®/GOx films in stirred, deaerated, pH 7, 0.1
M PBS, and (c) Average Michaelis-Menten response to glucose for films in (b)………………128
xvii
Figure 7.1. Cyclic voltammetry of (a) a gold electrode in deaerated 1.0 mM nitrophenyl diazonium
salt + 0.1 M tetrabutylammonium tetrafluoroborate /acetonitrile solution at a scan rate of 100 mV
s-1. Inset: Quartz crystal microbalance data, showing the decrease in frequency with each cycle.
(b) Two cycles (100 mV s-1), depicting the first and second cycle of the electrochemical conversion
of nitrophenyl to aminophenyl groups in deaerated, 0.1 M KCl /10 vol. % MeOH solution are
shown…………………………………………………………………………………………...136
Figure 7.2. (a) Mass gain overlapping the first cyclic voltammogram of a gold electrode in
deaerated 1.0 mM nitrophenyl diazonium salt + 0.1 M tertbutylammonium
tetrafluoroborate/acetonitrile solution (100 mV s-1), and (b) Corresponding plot of mass change as
a function of charge consumed during the first cycle in (a)……………………………………. 138
Figure 7.3. Cyclic voltammogram (0.1 V s-1) for bare Au and three nitrophenyl-modified Au
electrodes (after 1, 3 and 5 cycles in the diazonium salt solution) in deaerated 0.5 M H2SO4…....140
Figure 7.4. (a) Glucose signal (1.2 V) at bare Au substrate (■), aminophenyl-modified Au (●), and
immobilized GOx on Au (▲). Experiments performed in O2-saturated, stirred, 0.1 M, pH 7 PBS
(25 mL) with 0.8 mM additions of 1.0 M glucose, and (b) H2O2 oxidation signal (1.2 V) at bare
Au, and at immobilized GOx on Au. Experiments performed in O2-saturated, stirred, 0.1 M, pH 7
PBS (25 mL) with 2.5 mM additions of 3.0 M H2O2…...………………………………..……..144
Figure 7.5. (a) Cyclic voltammetry of an immobilized-GOx/IrOx film (133 nmol Ir) (dashed line),
and a standard IrOx film (133 nmol Ir) (solid line), and (b) Glucose signal (at 1.2 V) at a bare Au
substrate (■), an IrOx film on Au (●), immobilized-GOx film on Au (▲), and an immobilized-
GOx/ IrOx film on Au (▼). Experiments performed in aerated 0.1 M, pH 7 PBS (25 mL) with 20
µL aliquots of 1 M glucose……………………………………………………………………..147
Figure 7.6. Scanning electron microscopy image of an immobilized-GOx/IrOx/Nafion® film after
glucose sensing………………………………………………………………………………….149
Figure 7.7. (a) Cyclic voltammetry (100 mV s-1) in deaerated, stirred 0.1 M, pH 7 PBS, and (b and
c) Michaelis-Menten glucose response for immobilized-GOx/IrOx/Nafion® containing increasing
amounts of IrOx NPs (ranging from 88, 133, 176 nmol Ir). Experiments performed in stirred, (b)
deaerated and (c) aerated 0.1 M, pH 7 PBS (25 mL) with 20 µL aliquots of 1.0 M glucose……153
Figure 7.8 (a) Cyclic voltammetry (100 mV s-1) for immobilized-GOx/IrOx/Nafion® sensing
films, where the diazonium-derived film was deposited in 1, 3 or 5 cycles (resulting in surface
coverages of 1.4, 2.1 and 3.4 nmol cm-2, respectively), and Michaelis-Menten glucose response in
both deaerated (b) and aerated (c) environments for the films in (a)…………………………….156
Figure 7.9. (a) Cyclic voltammetry (100 mV s-1) of immobilized-GOx/IrOx/Nafion® sensing
films, where the diazonium-derived film was deposited at a constant potential of -0.6 V for 10 to
120 s in the diazonium salt solution, and the Michaelis-Menten glucose response in (b) deaerated
and (c) aerated solutions for films in (a)………………………………………………………...159
xviii
Figure 7.10. (a) The surface coverage of aminophenyl groups vs. the real Au surface area, and (b)
the glucose signal produced by immobilized-GOx/IrOx/Nafion® sensing films fabricated using Au
substrates of increasing roughness……………………………………………………………...162
Figure 7.11. Cyclic voltammetry (100 mV s-1) showing (a) the grafting of the nitrophenyl group
from the diazonium salt solution on a carbon paper substrate, (b) the subsequent conversion of the
nitro to amino groups, and (c) a comparison of the grafting process on carbon paper vs. Au
substrates………………………………………………………………………………………..164
Figure 7.12. Cyclic voltammetry (50 mV s-1) of (a) bare carbon paper (CP), a nitrophenyl-CP film,
and an aminophenyl-CP film in deaerated, stirred, 0.5 M H2SO4 and, (b) CVs of immobilized-
GOx/IrOx/Nafion® film (180 nmol IrOx) on CP in deaerated, stirred 0.1 M pH 7 PBS (inset shows
the IrOx anodic and cathodic peak current vs. the square root of the sweep rate)………………166
Figure 7.13. Michaelis-Menten glucose response in both deaerated and aerated environments for
immobilized-GOx/IrOx/Nafion® sensing films on carbon paper. Experiments were performed in
stirred, pH 7, 0.1 M PBS………………………………………………………………………..167
Scheme 2.1. Electrochemical mechanism for grafting a functionalized aryl group to a substrate
from a diazonium salt solution……………………………………………………………………16
Scheme 2. 2. Common methods of electrochemical detection of glucose based on GOx: (A) first-
generation biosensors based on the use of oxygen, (B) second-generation biosensors based on
soluble, mobile redox mediators (M), (C) third-generation biosensors based on direct electron
transfer between GOx and the electrode, (D) fourth-generation biosensors based on direct electron
transfer between GOx and the electrode via non-mobile mediators, and (E) direct electro-oxidation
of glucose at the electrode……………………………………………………………………..…21
Scheme 2.3. Schematic showing oxidation and reduction of the flavin group of the flavin adenine
dinucleotide FAD redox active site of GOx37……………………………………………………..29
Scheme 5.1 Dual mediation occurring in an IrOx/Nafion®/GOx film. Direct mediation occurs
when GOx is in good electrical contact with the interconnected IrOx NP matrix, while O2-
mediation takes place when GOx is not well-connected to the IrOx NP matrix. The triangle
represents glucose. Note that Nafion® is not represented in this scheme…………………………84
Scheme 5.2 Electron transfer pathway through IrOx matrix when the amount of GOx in the sensing
film is increased…………………………………………………………………………………108
Scheme 6.1 Shows the effect of film thickness on the collection and oxidation of H2O2 at the
sensing film…………………………………………………………………………………………125
Scheme 7.1. (a) Method of depositing the nitrophenyl diazonium-derived layer on sputtered Au,
the subsequent succinylation of the amino group, and immobilization of glucose oxidase (GOx).
xix
(b) Shows GOx immobilized to the Au substrate via a diazonium-derived succinylated
aminophenyl linking chain. IrOx NPs are intercalated within the film, surrounding GOx, allowing
for direct electron transfer from the active site of GOx to the underlying Au substrate.…………134
Scheme 7.2 Ir NP intercalation models for the deposition of the Ir ink into the diazonium-derived-
immobilized GOx film, prior to the electrochemical conversion of the Ir NPs to
IrOx.…………………………………………………………………………………….............150
Scheme 7.3. Schematic representation of single IrOx nanowire (left) and a well-interconnected
and thicker IrOx nanowire, resulting from a higher IrOx loading. The structure on the right would
be expected to have a lower IrOx surface area (per gram) than that on the left…………………..151
Scheme 7.4. Direct electron transfer (via IrOx) pathways with increased film IrOx content
(assuming tight IrOx NP packing, Scheme 7.2b)………………………………………………..155
xx
LIST OF ABBREVIATIONS
Abbreviation Definition
% v/v H2O Volume % of H2O added to absolute ethanol Ir sol
ACN Acetonitrile
ADA American Diabetes Association
BGL Blood glucose levels
BSE Back scattered electron
CD Carboxymethylated-cyclodextrin
CE Counter electroded
CGM Continuous glucose monitor
CNT Carbon nanotube
CP Carbon paper
CV Cyclic Voltammetry (or cyclic voltammogram)
DMF Dimethylformimide
dNB Diazonium derived nitrobenzene monolayers
EDC ethyl(dimethylaminopropyl) carbodiimide
EtOH Absolute ethanol
FAD Flavin adenine dinucleotide (oxidized form)
FADH2 Flavin adenine dinucleotide (reduced form)
GC Glassy carbon
GDH Glucose dehydrogenase
GL Glucose levels
GOx Glucose oxidase enzyme
H2O2 Hydrogen peroxide
HRP Horse radish peroxidase
xxi
Hupd Hydrogen Underpotential Deposition
Ir Metallic Iridium
IrOx Iridium oxide
IrOx/GOx Iridium oxide / glucose oxidase composite films
IrOx/Nafion®/GOx Iridium oxide/ Nafion®/ glucose oxidase composite films, fabricated using
aliquot deposition
ISF Interstitial fluid
Immob-
GOx/IrOx/Nafion®
Iridium oxide/ Nafion®/ glucose oxidase composite films, fabricated by
immobilizing GOx to a diazonium-derived film and then intercalating the
ethanolic Ir/Nafion® ink into the film
MM Michaelis-Menten
NAD Nicotinamide adenine dinucleotide
NHS N-hydroxysulfosuccinimide
NP Nanoparticle
NPD Nitrophenyl diazonium salt
OER Oxygen evolution reaction
ORR Oxygen reduction reaction
PBS Phosphate buffer solution
Ph-NH2 Aminophenyl group
Ph-NO2 Nitrophenyl group
PQQ pyrroquinolinequinone
QC Quartz crystal
QCMB Quartz crystal microbalance
RE Reference Electrode
RHE Reversible Hydrogen Electrode
RT Room Temperature
SA Succinic anhydride
SAM Self assembled monolayer
xxii
SE Secondary electron
SEM Scanning Electron Microscopy
TBA Tertbutylammonium tetrafluoroborate
tNB Thiol derived nitrobenzene monolayers
TEM Transmission Electron Microscopy
WE Working Electrode
XPS X-Ray Photoelectron Spectroscopy
XRD X-Ray Diffraction
xxiii
LIST OF SYMBOLS
Symbol Value Units Definition
∆f Hz Crystal frequency change
∆m μg Mass change
A cm2 Area
A1/C1 Ir(III)/Ir(IV) oxide redox peaks
Å 1*10-10 m Angstrom
β Parameter used to describe the effect of GOx
loading and film thickness on the apparent Km
value
E V Electrode potential
E0 V Standard reduction potential
E1/2 V Redox potential
E+ V Upper potential applied
E- V Lower potential applied
EB eV Binding energy
EK eV Kinetic energy
Ep V Peak potential
[E]total Total enzyme concentration ([ES] + [E])
f0 Hz Crystal initial frequency
ΔG‡ Activation energy
i mA cm-2, or µA
cm-2
Current density
I mA Current
ip mA Peak current
i/t Chronoamperometry
xxiv
imax µA cm-2 MM parameter: Maximum rate of enzyme
activity; proportional to the amount of active
enzyme
Km mM MM parameter: substrate concentration
where the enzymatic rate is equal to half the
maximum rate of the enzyme reaction with a
substrate (½ imax)
K’m mM Apparent Km obtained from experimental data
k1 Forward rate constant for E + S ES
k-1 Reverse rate constant for ES E + S
k2 Forward rate constant for catalysis (ES E
+ P)
k3 Forward rate constant for Ered Eox
M g mole-1 Molar mass
n mole Number of moles
q mC/cm2 charge density
[S] Substrate concentration for enzyme catalytic
reaction
[S0] Initial substrate concentration
θ ° X-ray incident angle
λ 1.5506 Å Cu Kα radiation wavelength
μq 2.947×1011 g cm-1 s-2 The crystal shear modulus
ρ g cm-3 Density
ϕ Spectrophotometer work function
ʋ V s-1 Sweep rate or scan rate
ʋo V s-1 Onset of irreversibility behaviour, or
reversibility parameter
QA C Anodic charge density
QC C Cathodic charge density
Vmax Maximum rate of reaction
CHAPTER ONE: INTRODUCTION
1.0 Project Background
Worldwide, it is expected that in less than 20 years, the number of persons with diabetes will
reach 438 million2. In Canada alone, diabetes is a contributing factor in the untimely death of
approximately 42 thousand persons each year2. Defined as a group of chronic diseases, there are
three main types of diabetes3, Type 1, the more common Type 2, and gestational diabetes. Type 1
diabetes is an autoimmune response that occurs when the pancreas does not produce insulin. T-
cells attack the body’s beta cells, which are located in the islets of Langerhans of the pancreas. The
beta cells produce insulin, and if they are damaged, cannot do so properly3,4. Type II diabetes,
which has been linked to obesity, occurs when the body becomes resistant to the effects of insulin,
or no longer makes enough insulin to maintain normal blood glucose levels (BGLs)5. Gestational
diabetes is typically temporary and occurs during pregnancy6. This inability leads to either hypo-
or hyperglycemic levels that can result in a wide variety of medical complications, including nerve
damage, as well as heart, kidney and eye disease. A typical blood glucose range for a non-diabetic
patient can range from 5.6 (fasting) to 7.8 mmol/l (after a meal)7, while levels can deviate to as
low as 2 mmol/l (unconsciousness) and as high as 33 mmol/l (Diabetic hyperosmolar syndrome)7.
Good management of diabetes considerably reduces the risk of developing complications
and prevents premature death, as such, it is important that blood glucose levels (BGLs) are properly
monitored through the use of glucose monitors (GMs) and medications. GMs are used to measure
a person’s BGLs continuously throughout the day to alert the person if their glucose levels have
fallen out of the normal, healthy range.
There are many different types of GMs on the market, most of which do not fall under the
2
American Diabetes Association’s (ADA) maximum inaccuracy values of 5%. In 2009, the most
accurate GMs at the time only achieved 5% accuracy 63% of the time2. Ginsenberg wrote that,
“As insulin delivery improves, the need for more accurate measurement of glucose becomes more
important2”. Thus, the research and development of glucose biosensors is an extremely well-
funded field. In 2012 alone, the ADA invested nearly $34.6 million into diabetes research3.
Even with all of this scientific exploration and funding being directed towards sensor
research, current sensor technology has proven to be inadequate in terms of accuracy and stability,
especially in the field of continuous and implantable sensors. The main reasons for these shortfalls
are that many of these devices utilize “old generation” technology that suffers from four main
sources of inaccuracies, including testing strip, physical, patient and pharmacological factors2. A
major source of inaccuracy is the O2-sensitivity, or, O2-dependence, of the sensor. This occurs
when the glucose signal is affected by the O2-deficit (in blood) and will thus vary substantially as
blood oxygen levels fluctuate throughout a normal day.
In this thesis work, a “new generation” glucose biosensing film, comprised of IrOx
nanoparticles (IrOx NPs) and glucose oxidase (GOx), is characterized and optimized to provide a
glucose signal that is accurate and not affected by the factors that affect “old generation” sensors.
This IrOx/Nafion®/GOx glucose sensing film is novel and unique in that it utilizes IrOx NPs as an
electron relay "wire" to regenerate GOx after the flavin adenine dinucleotide (FAD) active site is
reduced to FADH2 through the oxidation of glucose. Although the use of IrOx NPs and “enzyme
wiring” is not itself original, the combination of these two approaches is completely innovative.
In collaboration with the Department of Biomedical Engineering at the University of Calgary, this
sensing film will be incorporated into an externally applied patch, dubbed “the electronic
3
mosquito”8. The technology behind the mosquito will allow for painless and continuous
monitoring of a person’s BGLs, likely using our glucose biosensing film.
1.1 Research objectives
The primary goal of this thesis work has been to understand the effect of the Ir oxide
(IrOx)/Nafion®/glucose oxidase (GOx) sensing film components on the distribution of each
species in the sensing films and on the glucose response, and then use this knowledge to improve
the film sensing characteristics. Categorically, the focus of this thesis is:
To understand the effect of water, ethanol, the IrOx NPs, GOx molecules, and Nafion®,
on sensor performance and how they dictate the regeneration mechanism of GOx.
To fabricate a uniform and reproducible glucose sensing film morphology.
To enhance sensor sensitivity.
To achieve direct electron transfer through an IrOx NP matrix, independently of the
partial pressure of O2 in the sample solution.
1.2 Organization of thesis
This thesis is divided into eight chapters. In Chapter 1, the motivation behind this thesis
work is discussed, as well as the overall thesis objectives. Chapter 2 delivers a comprehensive
review of all pertinent background information regarding the thesis topic, including a review of
the literature required to understand the theories and models discussed in this thesis. A summary
of the experimental methods used throughout the thesis work is provided in Chapter 3.
In Chapter 4, the morphology and redox kinetics of Ir and Ir oxide (IrOx) nanoparticle (NP)
thin films, derived from ethanolic Ir NP sol-based inks, were characterized. The effect of the
water:ethanol content of the Ir NP ink on subsequent dried film morphology, stability and redox
kinetics, was investigated, and a small amount of Nafion® was found to enhance IrOx
4
interconnectivity and film stability. It was also found that glucose, and the products of its reaction
with glucose oxidase (GOx), do not affect the IrOx redox kinetics. Section 4.5.2 is published in
Biosensors and Bioelectronics (2013)9, where Dr. H. Elzanowska (a visiting scientist from Warsaw
University) assisted in suggesting experimental directions, discussion of the results and reading
over of paper drafts.
In Chapter 5, the effect of GOx, water and ethanol, the IrOx NPs, and Nafion® on dual-
mediation (O2- and direct-mediation of electrons from the GOx active site to the underlying Au
substrate) of our glucose sensor was studied. It was shown that the O2-medation route is favored
when the GOx:IrOx mass ratio in the film was non-optimal, or when an Ir ink water-content of >
20 vol. % was used, considered to be due to GOx/IrOx NP aggregation. GOx was found to require
an ink water content of at least 20% for optimal glucose sensing (performed at a potential of 1.2
V), and Nafion® was required for film stability. Section 5.2.1 was published in Biosensors and
Bioelectronics (2013)9, and Sections 5.2.2 and 5.5 were published Electrochimica Acta (2010)10,
where A. Jhas, a previous M.Sc. student in the Birss group, initiated the project, and Dr. H.
Elzanowska (a visiting scientist from Warsaw University) assisted in suggesting experimental
directions, discussion of the results and reading over of paper drafts.
In Chapter 6, the concept of O2-independent sensing is introduced and the two GOx
regeneration pathways were shown to exhibit different Michaelis-Menten parameters. The film
was optimized for O2-independent sensing by manipulating the film thickness, and sonicating the
ink prior to film fabrication. Good reproducibility, sensitivity and small K’m values were achieved.
However, complete O2-independence could not be realized. Sections 6.2.2, 6.3.1 and 6.3.2 were
published in Biosensors and Bioelectronics (2013)9 and Section 6.1.1 was published in
Electrochimica Acta (2010)10, where Dr. H. Elzanowska (a visiting scientist from Warsaw
5
University) assisted in suggesting experimental directions, discussion of the results and reading
over of paper drafts.
In Chapter 7, a new approach to film fabrication was taken, where GOx was immobilized
to the Au substrate by covalently binding the enzyme to a diazonium-derived aminophenyl layer
bound to the Au. Many parameters, including the amount of Ir assumed to be intercalated into the
films, the nitrophenyl surface coverage, and the substrate surface roughness, were optimized to
achieve the highest possible glucose signals, and O2-independent signals.
Lastly, the global overall conclusions of this thesis work, and the recommended future
work that would supplement the findings of this thesis, are given in Chapter 8.
6
CHAPTER TWO: RELEVENT BACKGROUND
2.0 Introduction to glucose biosensors
The goal of this research has been to optimize the construction of a glucose biosensor for the
monitoring of blood glucose levels of diabetic persons. This chapter will provide a short overview
of diabetes, and what causes it, as well as the sensors that have been developed to help monitor
this disease. The different types of sensing film components that are typically used in glucose
biosensors, along with theory behind the fabrication of the sensors will be presented. A brief
introduction to the film components used in this thesis work will be given, as well as the theory
behind the techniques used to characterize and fabricate the sensing films.
2.1 Development of glucose sensors for the management and monitoring of diabetes
2.1.1 Types of biosensors
A biosensor is a device with the capability of quickly producing analytical readings of either
a single, specific analyte or multiple analytes, depending on the sensor’s purpose. The glucose
biosensor, in particular, monitors (measures) the concentration of glucose in blood alerting a
person if their blood glucose levels are outside of the normal range.
Biosensors are typically comprised of two components, a bioreceptor and a transducer. The
bioreceptor is a biological molecule that recognizes a specific target analyte and is, in some way,
bound to the transducer. Many different bioreceptors are used in biosensors, such as enzymes,
antibodies, nucleic acids or even whole cells11. The transducer converts the bioreceptor recognition
event into a measurable signal12. The six most common classes of transducers include
piezoelectric, optical, thermometric, pyroelectric, calorimetric and electrochemical13.
Piezoelectric sensors are typically antibody-based sensors14 (immunosensors), where an
antibody is immobilized to the piezoelectric transducer. A piezoelectric material has the unique
7
property where, when an electrical field is applied across the crystal, it vibrates at a specific
frequency. When the antibody’s complementary antigen binds to it, there is an increase in mass on
the sensing surface. This increase in mass is detected by the change in frequency at which the
piezoelectric material vibrates, which allows for the detection of antigens in the picogram range.
In comparison, optical sensors typically use fluorescence-based detection, which requires
the analyte to be labeled. Some types of optical sensors are capable of label-free detection (analyte
does not have to be tagged, and is detected in its natural form) by studying changes in the refractive
index, optical absorption, or by Raman spectroscopic detection15.
Thermometric, pyroelectric, and calorimetric sensors all fall under a similar category of
sensors, as they measure the enthalpy change due to a reaction13. The benefit to these sensors is
that the analyte does not require a label (as some optical sensors require). However, these sensors
are not typically appropriate for measuring small concentrations (nano- or picomolar), and as such
are not ideal for diagnostic testing.
Of the six transducers, the electrochemical sensor is the most commonly employed for
glucose biosensing, especially since many of the bio-recognition elements that react with glucose
are electrochemically active. As such, the number of electrons transferred during the reaction of
the bio-recognition element and glucose can be quantitatively measured to give a signal which is
proportional to the concentration of glucose. The most common types of electrochemical sensing
include cyclic voltammetry, chronoamperometry, chronopotentiometry, impedance spectroscopy,
and various field-effect transistor based methods.
In the present work, the focus is on electrochemical transducers (or electrodes), and, in
particular, on amperometric transducers, due to their high sensitivity and suitability for mass
production16. Amperometric sensors function by producing a current when a constant potential is
8
applied between two electrodes. When the bio-recognition element that is immobilized at the
electronically conductive transducer reacts with its analyte, electrons are either transferred to or
removed from the transducer. The number of electrons transferred is proportional to the
concentration of the analyte. There are many different types of amperometric sensors on the market
and in the literature for the detection of analytes, such as glucose17-20, alcohol16,21, biomarkers22,23,
and cholesterol24.
2.1.2 Glucose sensors in clinical trials and currently on the market
With such a high population of people who suffer from diabetes, scientists have been
developing novel techniques in which a patient can easily detect and monitor their blood glucose
levels (GLs). There are three types of glucose sensors, invasive (finger pricking, blood drawing,
painful), semi-invasive (minimal pain), and non-invasive (no drawing of blood, pain or puncturing
the skin). Continuous glucose monitoring (CGM) allows for the optimal treatment of diabetes by
providing a continuous measurement of glucose concentrations throughout the day. There are both
invasive and non-invasive types of CGMs.
The invasive CGMs typically involve inserting a small sensor either subcutaneously or
intravenously into the body. The sensor then continuously measures the blood glucose levels and
relays the information to an external, wireless controller via radio frequency or optical signaling.
These sensors have a wide list of drawbacks, as they can easily be fouled, the sensor can migrate
within the body, and they can result in tissue inflammation. Typically minimally invasive and non-
invasive glucose monitoring involves measuring interstitial fluid (ISF) glucose concentrations.
Interestingly, there does not appear to be consensus as to whether or not there is a relationship
between glucose levels from ISF and blood. One study25 found that, if blood GLs were manipulated
so as to rapidly change, there was no significant lag in readings observed among the ISF, capillary
9
and venous glucose levels. A second group26 found that, in dogs, there was a time delay established
of only 5-12 minutes between glucose concentration in plasma and ISF, which was concluded to
be insignificant. Other groups believe that the relationship between GLs in the blood vs. ISF is too
significant to consider detecting glucose levels from ISF24, 25.
Non-invasive devices have been deemed “the holy grail” of glucose sensing. Interestingly,
however, a leading scientist in glucose biosensors and head of the glucose biosensor company,
Glucovation, Robert Boock, recently stated in an interview with MobiHealthNews27 that non-
invasive technologies are difficult, if not impossible, to produce reliably. He stated “That (invasive
sensing technologies are) the price you have to pay if you want real science”27. A second leader in
the field, Dexcom’s Executive Vice President of Strategy and Corporate Development, Steve
Pacelli, agrees with this statement, saying that, “Ultimately, I think both non-invasive intermittent
glucose sensors (not requiring finger sticks) and continuous glucose monitoring will be the only
technologies some years from now17.” However, novel non-invasive devices are always highly
anticipated, due to the painlessness of the measurement.
The “Biometric watch” is a non-invasive device, recently published by the Zalevsky group28,
where the wearable glucose sensor uses changing patterns of scattered light to directly monitor
blood GLs. The device, however, suffers from a high margin of error from a medical glucometer.
According the The Wall Street Journal29, the iWatch, which may operate in a similar fashion to
the biometric watch, may shortly be introduced to the public by Apple.
Recently, the glucose-sensing contact lens by Google has been making headlines30,
praising the ability of the contact lens to painlessly monitor GLs in the patient’s tears. This sensor,
published by Yao et al., in 2012 is an electrochemical sensor based on glucose oxidase31. An issue
with this sensor is the fouling of the electrode, as tears contain a large number of enzymes and
10
proteins. Fouling leads to poor electron transfer resistance and complex impedance changes for
the electrochemical cell.
The majority of glucose sensors currently on the market are either invasive, or minimally-
invasive. There are many high profile companies that manufacture glucose biosensors currently
on the market, including Yellowsprings Instruments, Nova Biomedical, Bayer AG, Roche
Diagnostics AG, Dexcom, LifeScan, and Neogen Corporation32. For a sensor to be viable on the
market, it must be versatile and inexpensive. These companies are constantly trying to solve the
global technical issues of sensor lifetime, H2O2 production, matrix interference, O2-dependent
sensing, sensor fouling, signal drift, and microbial contamination32.
Nova’s StatStripTM Glucose Monitor33 has a very short analysis time (6 s) that accurately
measures blood glucose levels for persons (such as neonatals) with hematocrit abnormalities. This
sensing technology also corrects for electroactive interferences and eliminates oxygen interference
to ensure accurate glucose test results. The Glucometer Elite® Diabetes Care Sytem34 by Bayer
AG is an invasive, electrode-based sensor technology that uses capillary action at the end of a test
strip to draw in blood to the detection chamber. Results are not as rapid as with the StatStripTM,
as this system requires 30 s. Roche Diagnostics offer a minimally invasive monitoring system
called the Accu-Chek Plus Glucose Meter35, which is preloaded with 17 diabetes test strips that
monitor blood glucose levels continuously. LifeScan (Johnson and Johnson) sells disposable test
strips that give rapid test results (5 s), but require blood to be drawn from either the finger or
forearm.
LifeScan36 has, however, developed the Ultra SoftTM Adjustable Blood Sampler that allows
the patient to adjust the puncture depth of the lancet, theoretically decreasing the pain associated
with finger pricking. Dexcom37 has manufactored a continuous blood glucose meter that inserts a
11
small sensor just beneath the skin and a transmitter wirelessly sends the information to a receiver
that alerts the patient if their glucose levels are within a good range or not. This device is one of
the longest wearing 7-day sensors and is the only continuous monitoring system approved for
toddlers.
A final example of a minimally-invasive sensor is the sensing device we are developing in
collaboration with the Department of Engineering at the University of Calgary. Termed “the E-
Mosquito”, it is externally positioned and continuously measures GLs from fluid obtained from
the skin tissue (blood or ISF) in a painless manner8 The mosquito is a patch consisting of a
microneedle array. Each microneedle is attached to two piezoelectric strips that push the needle
below the skin, allowing for the painless extraction of a small blood sample (0.6 µl) from the
capillaries8. GLs are then determined when the sample comes into contact with the sensing film
developed in this thesis work.
2.1.3 Existing challenges of glucose biosensors
Accuracy is one of the most important requirements of any glucose monitoring device.
According to the International Organization for Standardization guidelines (ISO 15197), when
measuring glucose concentrations below 4.2 mM, the measurement is considered accurate if the
error is within 0.8 mM of the actual concentration. At blood glucose concentrations above 4.3 mM,
an error of up to 20% is acceptable38.
There are many potential sources of inaccuracy in an electrochemical biosensor, especially
as the sensor’s ability is based on a biological element that requires a specific set of environmental
conditions for proper operation. One of the greatest sources of error in many enzymatic
electrochemical sensors is the O2-dependent response. The use of exogenous mediating
compounds can also lead to inaccurate sensing, as the mediator can be lost from the film, changing
12
the mediator concentration. Enzymes can be denatured or degraded over time, resulting in a
decrease in stability and activity. Many enzymes also suffer from poor specificity for the target
analyte, leading to false, hyper-glycemic readings.
The sensing films can also be affected by electrochemically interfering species, such as
ascorbic or uric acid. It was shown that interfering species can result in up to a 193% error in the
accuracy of a reading39. Finally, as explained by Vaddiraju et al., calibration of the device is often
difficult as it often requires the patient to use the finger-prick test strips40.
Other factors that should be considered for a good glucose sensor include reliability, with a
high expected uptime, good durability, and a quick response time. It has been shown that, if the
glucose levels in the ISF are measured, there can be a delay of up to 10 minutes between the blood
and interstitial glucose concentrations, with the ISF glucose levels lagging behind BGLs 81% of
the time 41. This is especially problematic if a patient engages in strenuous physical activity or
consumes a sugary meal.
Biocompatibility of the sensing film is a challenge when fabricating sensors that will be
implanted either subcutaneously, or intravenously. For a film to be biocompatible, it must meet a
wide range of regulations. For example, the reactions that take place in the films must not generate
heat, evolve gas or change the pH of the surrounding environment. The film should be corrosion
resistant, and should also be flexible/soft, so as to not damage any surrounding tissues.
2.2 Glucose biosensor components
2.2.1 Glucose sensing enzymes
There are many different enzymes available for amperometric sensing, as the main
qualification is that the enzyme is redox active. For glucose sensing, there are two main families
of redox active enzymes, including glucose oxidase (GOx) and glucose-1-dehydrogenase (GDH).
13
Each of these enzymes has different active sites and cofactors and therefore has different redox
potentials, turnover rates (number of subsrate molecules converted to product per catalytic
site/second), and selectivity for glucose (vs. other sugars)42.
Of the two enzyme families, GOx enzymes are more commonly used for a multitude of
reasons, including that GOx is extremely robust and capable of withstanding a range of pH, ionic
strength and temperatures43,44. GOx has the greatest selectivity, as the dehydrogenase family tends
to react with other interfering sugars, such as maltose, xylose and galactose45.GOx maintains a high
activity and stability for a relatively long period of time43. The flavin adenine dinucleotide (FAD)
cofactor (Scheme 2.3), used by GOx is tightly bound to the active site43, whereas the cofactors for
GDH enzymes (pyrroquinolinequinone (PQQ) or nicotinamide adenine dinucleotide (NAD)) are
not tightly bound to GDH, and therefore must be added exogenously.
The most notable benefit of using GDH vs. GOx is that the enzymatic reaction of GDH with
glucose is independent of the oxygen concentration (Reactions (2.4-2.7)), whereas GOx requires
O2, or an external mediator, to regenerate its active site (Scheme 2.2a-d). The cofactor is
regenerated by diffusing through solution and transfering electrons to/from the transducer.
Unfortunately, the mobility of the cofactor can lead to sensitivity issues if it is lost to solution
instead of reacting at the current collector. GDH-PQQ is not as commonly used in glucose
biosensors as is GDH-NAD, as the water-soluble GDH-PQQ has low selectivity and poor thermal
stability, while membrane-bound GDH-PQQ requires detergents for solubilization in aqueous
solutions45. Due to its excellent stability and selectivity, GOx is the most commonly used enzyme
for glucose biosensing.
GOx-FAD: Glucose + GOx(FAD) Gluconolactone + GOx(FADH2) (Reaction 2.1)
GOx(FADH2) + O2 GOx(FAD) + H2O2 (Reaction 2.2)
14
GDH-PQQ: Glucose + PQQ(OX) gluconolactone + PQQ(RED) (Reaction 2.4)
PQQ(RED) PQQ(OX) + 2H+ + 2e- (Reaction 2.5)
GDH-NAD: Glucose + NAD+ gluconolactone + NADH (Reaction 2.6)
NADH NAD+ + H+ + 2e- (Reaction 2.7)
The other major component of amperometric glucose biosensors is the matrix, or film, in
which the enzyme is immobilized. There are many different ways in which to immobilize an
enzyme (see Section 2.2.1.1), and there are many different materials used as immobilization and/or
electron transfer matrices. Nanomaterials make up the majority of matrix materials, which can be
further categorized into metals, metal oxides, carbon nanotubes (single and multiwall), graphene,
graphene oxide and polymers46.
2.2.1.1 Enzyme immobilization techniques
Perhaps one of the most difficult aspects of fabricating an enzymatic biosensor is the
immobilization of the enzyme on the transducer, while maintaining high enzymatic activity and
stability. There are many different strategies for enzyme immobilization, including physical
adsorption, physical entrapment, and covalent attachment, to name a few. Immobilization should
not affect the structure or function of the enzyme, access of the analyte to the active site of the
enzyme, or the removal of any products.
Physical adsorption is one of the simplest and quickest methods to immobilize enzymes to a
surface. However, it is not always the most effective method, as the stability of a physisorbed
enzyme is poor. The interactions during adsorption are limited to Van der Waals forces, which are
relatively weak in comparison to those formed during covalent bonding. Stability depends on the
type of enzyme, as well as a combination of environmental factors, such as temperature, surface
tension, electrostatic interactions, ionic strength, turbidity and pH of the solution. Non-optimal
15
conditions can cause an enzyme to desorb from the surface or become denatured/inactive. There
are a few examples of enzymes being successfully physically adsorbed to ZnO nanostructures.
Umar et al47., fabricated a sensitive cholesterol biosensor by physically adsorbing cholesterol
oxidase to the surface of “flowerlike” ZnO NPs. Wang et al48., designed a diffusion-controlled
glucose biosensor in which GOx was physically adsorbed on a ZnO nanocomb-modified Au
electrode.
Physical entrapment within a solid or hydrous matrix is a commonly used method for
enzyme immobilization, as it allows the enzyme to maintain its native configuration and therefore
high activity, while concurrently increasing the stability of the enzyme. The signal magnitude and
response time can suffer, however, as it can be difficult to transport the analyte, ions, electrons and
products through the matrix. As such, it is important that the selected matrix is quite porous or
permselective. A common method for encapsulation is using electropolymerized non-conducting
films, due to their resistance to fouling and their ability to repel interfering species that may be
present in solution. Li and Lin49 fabricated a glucose sensor by electrosynthesizing a composite of
Pt nanoclusters, embedded in polypyrrole nanowires, on a glassy carbon electrode. They then
immobilized GOx in an electropolymerized poly(o-aminophenol) film, deposited on top of the
Pt/polypyrrole composite.
Covalent bonding (a form of chemisorption) is another effective immobilization strategy,
where specific amino acids or glycosyl sites on the outside of the enzyme are bound to a functional
group attached to the transducer surface. Wan et al50., fabricated an glucose biosensor by linking
GOx to the hydroxyl groups of chitosan (which was bound to a polyaniline-coated gold electrode).
Covalent attachment of enzymes to organic layers formed from diazonium salts or self-assembled
monolayers (Section 2.2.1.2 and 2.2.1.3) is also a popular technique.
16
2.2.1.2 Diazonium salt chemistry
There are many examples in the literature where diazonium-derived films have been utilized
for immobilizing enzymes and other biological compounds51-53. The most common method of
grafting the aryl group of a diazonium salt to a surface is via electrochemical reduction by cyclic
voltammetry51-57. However, it is also possible for the reaction to occur spontaneously or
photochemically58. Delmar et al. were one of the first groups to electrochemically reduce an aryl
diazonium salt to form a covalent attachment to the surface of carbon in 199254. This is a two-step
process where a phenyl radical is formed, and then, the radical attacks the surface, forming a
covalent bond between the phenyl group and a surface atom of the substrate (Scheme 2.1).
Scheme 2.1. Electrochemical mechanism for grafting a functionalized aryl group to a substrate
from a diazonium salt solution.
This grafting process can be performed on a wide array of surfaces, including silicon59,
organic polymers60, carbon54,57,61,62, and metals, such as Fe63, and Cu64. Recently, it has been
demonstrated by Lauentius et al., that a carbon-gold covalent bond can be formed65. Belanger et
al., were among the first groups to deposit diazonium-derived films on a gold substrate66. Since
then, many groups have deposited aryl films on Au substrates66-69.
The thickness of the modified film formed by diazonium salt chemistry can also be
controlled by optimizing the grafting conditions. The formation of multilayers occurs when a
second radical attacks a phenyl group already grafted to the surface. It has been documented,
however, that it is difficult to control the formation of multilayers, often resulting in a film with
17
low packing density70. It is, however, possible to slow down the formation of multilayers by
placing sterically hindering blocking groups71 on the diazonium salt being reduced.
Liu et al72., fabricated an amperometric immunosensor for glycosylated hemoglobin by
attaching Au NPs to a glassy carbon surface by first depositing an aryl film to the carbon from a
diazonium salt, and then reducing the Au NPs to form a C-Au bond with the aryl film. Another
group, Radi et al., immobilized horseradish peroxidase to a screen-printed carbon electrode by first
modifying carbon with a layer of 4-nitrophenyl groups from a diazonium salt and crosslinking
horseradish peroxidase to the aminophenyl group by glutaraldehyde73. In contrast, Polsky et al74.,
first immobilized horseradish peroxidase to a 4-carboxyphenyl diazonium salt, and then attached
the enzyme-aryl diazonium salt conjugate to the transducer.
2.2.1.3 Thiol-derived Self Assembled Monolayers
Alkane-thiol chains will reproducibly chemisorb to a substrate to form highly organized
layers (Self assembled monolayers, or SAMs). Often SAMs are formed on Au substrates, with the
Au-S having a very high energy, and therefore, the S head group is always oriented toward the
surface in the 3-fold hollows in the Au lattice. The chains are tilted at an angle of ca. 30° in order
to maximize the Van der Waals interactions between chains75. The strength of these interactions
induces the alkane tail to align in a trans conformation. However, if the tail is > 11 carbons in
length, the tails become less ordered. An advantage to thiol-derived vs. diazonium-derived films
is that a single, uniform monolayer is formed.
Shewchuk and McDermott53 performed a thorough comparison to determine whether thiol
derived nitrobenzene monolayers (tNB) or diazonium derived nitrobenzene films (dNB) exhibit
better stability. Three tests were performed, where each of the films were subjected to either
refluxing in acetonitrile for one hour, sonication for 30 minutes in acetonitrile, or being immersed
18
in a “chemical displacement” solution of ethanolic octadecane thiol for 24 hours. Infrared
reflection absorption spectroscopy was used to monitor the number of remaining NO2 molecules
and cyclic voltammetry to monitor the electrochemical blocking of Au after each of the three
treatments. It was found that, in the case of sonication and refluxing in acetonitrile, more material
was lost from the multilayered dNB film in comparison to the monolayer tNB film. However, a
substantial amount of material remained on the Au for both films. It was hypothesized that the
material lost from the diazonium-derived film may have been disordered, strongly physisorbed
material (such as nitrobenzene dimers). In contrast, the entire tNB layer was lost during the
chemical displacement experiment, while ca. 25% of the dNB film was retained, indicating a
greater stability of the diazonium vs. the thiol-derived films on Au.
Diazonium-derived films have a greater degree of stability over time53,76 and a wider
potential window over which they are stable76, in comparison to thiol-derived films. As well,
diazonium derived aryl films can form covalent bonds with both carbon and metal substrates. As
a result of these findings, in this work, glucose oxidase was immobilized to the Au substrate via
diazonium-derived films (Chapter 7), as opposed to the use of self-assembled monolayers.
2.2.2 Enzyme immobilization matrices
Glucose biosensors that are enzyme-based have the problem of poor long-term stability, as
enzyme activity decreases with time. As such, non-enzymatic glucose sensors were fabricated to
address the problem of insufficient sensor stability. These sensors typically use noble metals, such
as Au77, Pt78, or Ni79,80, due to their high electrocatalytic activity and high sensitivity for direct
electro-oxidation of glucose (Scheme 2.2 e). Without the bioreceptor component, however, the
sensing film is not selective, and electroactive interfering species (such as other sugars, or drugs)
in the sample solution can react at the metal. It is also known that the reaction intermediates of the
19
breakdown of glucose (such as CO), and ions in solution (such as Cl-)81, can foul the metal surface,
decreasing the active surface area for glucose reaction.
Metal and metal oxide nanomaterials are commonly used as matrices due to their good
electronic conductivity, low cost (compared to bulk metals), high biological activity, and accute
sensitivity (due to their large surface:volume ratio)82. A downside to the use of metals and metal
oxides as immobilization matrices is that they are not all biocompatible. Metallic nanomaterials,
such as Ir83,84, Au85, Ag86, Ru87 and Pt87, can be used to decorate various carbon structures, such
as nanotubes (CNTs), or are dispersed within polymers, such as in polyaniline. Examples of oxides
used as matrices include IrO288-90, RuO2
19, TiO2 91
, SiO2 91
, ZnO92, CeO2 93 and ZrO2
94.
Metal and metal oxides can be fabricated into different nanostructural contructs, including
spheres, nanoparticles, nanotubes, nanoclusters, nanorods, nanowires, sol-gels, dendrites, etc.95
The ability to fabricate such a wide array of nanostructures results in different sensor constructs,
each with their own advantages and disadvantages. As an example, one of the major benefits of
sol-gel technology is that it is an inherantly low temperature process, and, as such, heat-sensitive
biological materials (such as enzymes, proteins and antibodies) can be encapsulated without
denaturation. Denaturation is the process of enzyme unfolding from its tertiary structure to a
disordered peptide, resulting in the improper alignment of the chief amino acids required for
enzyme functionality/activity.
Silica or titania sol-gels are commonly used as they are inert and stable, do not swell
substantially in aqueous solution, are biocompatible, and physically rigid. A disadvantage to the
use of sol-gels is cracking with time. In order to prevent cracking, polymers are mixed into the sol-
gel to form an organic/inorganic hybrid sol. Choi et al91., fabricated an amperometric glucose
biosensor based on the encapsulation of the enzyme glucose oxidase (GOx) in a sol-gel-derived
20
metal oxide and Nafion® composite film. They found that the film was stable over a long period
of time (80% of initial activity after 4 months) and did not crack. For comparison, Zang et al96.,
designed a stable amperometric glucose sensor by immobilizing GOx on ZnO nanowires, 1-D
nanomaterials, with unique catalytic and electronic properties. ZnO has a high isoelectric point,
which allows for excellent immobilization of GOx (an acidic enzyme) and simultaneous electron
transfer mediation to the current collector. With a high GOx loading, a high maximum current
density signal of 114 uA cm-2 was achieved.
Many glucose biosensors are carbon-based, as carbon is biocompatible, conductive, and
stable. Although carbon-based nanostructures are electonically conductive, CNTs, for example,
have a higher electrocatalytic activity in comparison to activated carbon or pyrolytic graphite97.
CNTs, in particular, are commonly used to immobilize enzymes during amperometric glucose
detection. Often the enzyme is covalently bound to the carbon walls through the use of a cross-
linking agent, such as gluteraldehyde98. A disadvantage of using CNTs is that they are insoluble
in most solvents and require the addition of a binding agent. Common binders include Nafion®97,
methacrylates99, or Teflon100, to name a few.
2.3 Four generations of amperometric glucose biosensors
With the progression of research in the field of glucose biosensors, four different
generations of amperometric glucose biosensors have emerged, all using glucose oxidase (GOx).
Each of these generations has enhanced the method of enzyme regeneration by utilizing different
mediators for electron transfer from the active site to the underlying transducer. Scheme 2.2 shows
the electron transfer processes for all four generations, as well as the direct reaction of the analyte
at the transducer.
21
Scheme 2. 2. Common methods of electrochemical detection of glucose based on GOx: (A) first-
generation biosensors based on the use of oxygen, (B) second-generation biosensors based on
soluble, mobile redox mediators (M), (C) third-generation biosensors based on direct electron
transfer between GOx and the electrode, (D) fourth-generation biosensors based on direct electron
transfer between GOx and the electrode via non-mobile mediators, and (E) direct electro-oxidation
of glucose at the electrode.
First-generation sensors (Scheme 2.2a) detect the concentration of hydrogen peroxide
(H2O2) produced during GOx regeneration by O2 (Reaction 2.2). H2O2 is normally oxidized at the
sensor electrode, producing an amperometric signal (current) that is proportional to the
concentration of glucose in the sample7. There are three main problems associated with first-
22
generation sensors, as outlined in a review by Wang8. The first is the issue of “oxygen deficit”,
where the oxygen levels in blood are an order of magnitude lower than the physiological blood
glucose concentrations, and, as such, there is not enough oxygen to efficiently regenerate the
electrode. A second problem is that, in order to measure the H2O2 concentration, a high operating
potential is required to achieve the desired selectivity. Finally, while working at such high
operating potentials, interfering electroactive species in blood, such as ascorbic acid,
acetaminophen, and uric acid, may also react at the electrode, causing inaccuracy of the sensor.
Second-generation sensors (Scheme 2.2b) operate at potentials where endogenous
electroactive species do not interfere8,9. This generation of sensors uses a soluble, mobile mediator
other than oxygen, e.g., organic and organometallic redox compounds, such as quinones,
ferrocenes, and organic conducting salts10. The use of a mediator other than oxygen alleviates the
obstacle of variable blood oxygen concentrations observed in first-generation sensors. Kutner et
al.11, used four different poly-pyridine complexes of ruthenium as mediating compounds for their
second-generation glucose biosensor. GOx and an anionic carboxymethylated-cyclodextrin (CD)
polymer film was cast on either a Pt or a glassy carbon (GC) electrode and the Ru complexes were
incorporated by inclusion in the CD molecular cavities and by ion exchange with CD. There are
many problems associated with the use of soluble and mobile mediators, including toxicity levels,
and the loss of the mediator to solution.
The third-generation of biosensors (Scheme 2.2c) involves the regeneration of the active site
directly at the surface of the transducer, eliminating the need for redox mediators. Often, this
involves deglycosylizing GOx to minimize the distance between the active site and the transducer
surface101, or the use of enzymes with short distances between the active site and the exterior of
the enzyme102.
23
The fourth-generation of biosensors12 (Scheme 2.2d) abolishes the use of mobile redox
mediators. Instead, this sensor design allows for direct electron transfer or “wiring” between the
enzyme and the electrode through a stationary, non-soluble mediator. Stationary mediators
(“wires”) are extremely efficient, because they are immobilized in the sensing film, and the
response is no longer dependent on the diffusion rate of mediators. Additionally, fourth-generation
sensors are generally more biocompatible, without the presence of toxic mediators, and they also
exhibit better selectivity to glucose. The work done in this thesis is based on this generation of
sensor. Heller et al.13, also developed a fourth-generation sensor, in which GOx was “wired” to
the metal substrate with a conductive relay chain. GOx was chemically modified by attaching
conductive relays (~ 12 molecules of ferrocene carboxylic acids) to lysine or tyrosine side chains
on the enzyme periphery. Those relays that were sufficiently close to the active site (<10 Å) were
able to transfer electrons from FADH2 to the substrate.
The direct oxidation of glucose (Scheme 2.2e) at an electrode surface does not give accurate
glucose signals, as other electroactive species may react at the electrode as well95. These types of
glucose sensors also have poor sensitivity due to the adsorption and accumulation of interfering
species to the active site95.
2.4 Iridium oxide/Nafion®/GOx-based glucose biosensor components
In this work, the glucose sensor is comprised of three main components, an iridium oxide
nanoparticle matrix, glucose oxidase, and a binding agent, Nafion®. In this section, pertinent
background information regarding these three sensing film components is provided.
2.4.1 Ir and Ir oxide nanoparticles
Ir belongs to a unique family of transition metals, commonly referred to as the “platinum
group”, which includes Ru, Rh, Pd, Os, Ir and Pt. This family of metals has been well studied, as
24
they exhibit excellent catalytic properties, have a high resistance to chemical attack, and are
electrochemically stable103. Many transition metal oxides exhibit pseudocapacitance, where
electrochemical energy is stored in the faradaic redox reactions at their surfaces104. Ir readily forms
a hydrous oxide (IrOx) film when exposed to oxidative potentials. This hydrous oxide is an
excellent material for a wide variety of technologies, including electrocatalysts105, as well as
electrochromic106,107 and biomedical devices108,109. There are several oxidation states of Ir in IrOx,
ranging from 2+ to 6+.
Pickup and Birss proposed a model describing the electrochemical conversion of Ir to
hydrous Ir oxide that outlined the five key stages of growth110. Interestingly, once a hydrous oxide
has formed, it can no longer be reduced. The electrochemical growth of IrOx (from Ir) is also
highly dependent on the aqueous growth medium in which electro-oxidation takes place111. In the
present work, Ir NPs are typically electrochemically converted to IrOx in a neutral solution. In a
neutral solution, IrOx follows a 60 mV pH dependence, where one H+ (or H3O+) is injected per
electron per Ir site during oxide reduction (and vice versa during oxidation), according to Reaction
(2.13). In comparison, a ~90 mV pH dependence is observed in an acidic solution.
Ir(IV)oxide • x H2O + e- + H+ Ir(III)oxide • x H2O (Reaction 2.13)
In our earlier work112, glucose oxidase (GOx) was deposited simultaneously with an IrOx
film formed electrochemically on a bulk Ir wire or plate substrate. However, most of the GOx
resided in the outer regions of the IrOx film, thus losing the intended stabilizing benefits of the
IrOx matrix, while also leaving costly, unused Ir metal beneath the IrOx film. For these reasons,
we have directed our efforts towards the encapsulation of GOx into films composed of IrOx
nanoparticles (NPs)88,90,113, formed using methods developed earlier114,115. Prior to film fabrication,
the Ir NPs are in a sol state, suspended in an ethanolic solution. Once deposited as a film, the Ir
25
NPs are electrochemically oxidized to IrOx, as described above, which increases the
interconnectivity between particles and forms an excellent, porous matrix for the immobilization of
GOx. IrOx is also known for its biocompatibility116-118, high surface area, and good conductivity119.
Ir and IrOx NPs are very versatile materials, as they can be used for a wide variety of
applications, with IrOx NPs being excellent catalysts for the oxygen evolution reaction (OER)120-
125, oxygen reduction reaction (ORR)126 and hydrogenation127-129 reactions. The production of
hydrogen from water is done via water-splitting124,130 or electrolysis131,132. Ir can also be combined
with other metals, such as Pt120, Ru122 or Au124, in order to fabricate a catalyst with an enhanced
OER activity. Zhao et al.124, fabricated IrOx/Au composite NPs that exhibited very good OER
turnover frequency (10.9 s-1). The electrocatalytic activity of the composite was reported to be
significantly enhanced (one order of magnitude) in comparison to IrOx NPs, attributed to the high
oxidation states of Ir caused by the transfer of electronic charge from the oxide to the gold
substrate. Meanwhile, Nakagawa121 demonstrated that their IrOx NP films exhibited only small
OER overpotentials of 0.15 V.
IrOx NPs have also been used frequently as highly conductive interfaces for other
applications, such as sensing. An ultra-sensitive electrochemical immunoassay based on Au-IrOx
NP assemblies22 was fabricated by Tang et al., for targeting biomarkers in serum. The role of the
IrOx-coated Au NPs was to generate protons (by splitting water), which, in turn, were used to
reduce the product of the sensing reaction. In comparison, Mayorga-Martinez et al., fabricated a
disposable IrOx NP-based tyrosinase biosensor133 for the detection of pesticides, such as catechol
and chlorpyrifos, in water samples. In this case, the IrOx NPs were used to transfer electrons, as
opposed to splitting water. An aliquot of suspended IrOx NPs (in aqueous solution) was deposited
on a screen printed carbon electrode, followed by an aliquot of glutaraldehyde and finally a
26
solution of tyrosinase and bovine serum albumin was deposited on top. At negative potentials, the
quinone product of pesticide oxidation was electrochemically reduced at the IrOx NPs, giving very
low detection limits.
It is also possible to fabricate IrOx NPs of different shapes and sizes. By manipulating the
shape of the NPs, it is possible to change both the chemical and physical properties of the transition
metal134,. The first highly dispersed Ir NPs were reported in a Nature paper in 1977 by Anderson
and Howe135. Many different shapes of Ir and IrOx NPs have been fabricated since then. Baida
and Diao136 prepared disc-like, ellipsoidal, and spherical NPs. Zhao et al., fabricated nanoflower-
shaped IrOx composite124 particles. Lee et al., fabricated Ir NPs with dendrites137 that were found
to exhibit enhanced catalytic activity toward OER, as opposed to NPs without dendrites. Zhao et
al., developed a novel template-assisted deposition and etching strategy for fabricating IrO2
nanotube arrays138 on conductive substrates.
IrOx is also a commonly used material for biomedical applications, such as for nerve
stimulation139,140 and pacemakers141, due to its excellent biocompatibility. Meyer et al140.,
electrodeposited IrOx films onto a range of metals (Au, Pt, PtIr, and stainless steel substrates) to
fabricate a mechanically stable, low impedance and high charge capacity coatings for neural
stimulation and recording electrodes. IrOx has also been used as an electrode (implanted in a 72-
year-old male) to detect blood pH changes in the right atrium141. The changes in pH were then
relayed to a pH-triggered pacemaker, where in the case of persistent acidosis, the pacemaker would
increase the pumping rate. IrOx has also been used for the fabrication of cardiovascular stents109,
defibrillation electrodes142 and cochlear implants143.
27
2.4.2 The role of Nafion® in sensing films
Nafion® is a copolymer composed of a hydrophobic tetrafluoroethylene back-bone and
hydrophilic perfluorosulfonate side chains, which can vary in length (Fig. 2.1a). When Nafion®
is in a hydrating environment, the sulfonate side chains will arrange such that they form
hydrophilic pockets or channels (Fig. 2.1b). These pockets can vary in size, depending on the
degree of hydration (1-5 nm)144. Nafion® is not electronically conductive, but facile proton
conduction does occur through the polymer as protons can hop from sulfonate pocket to pocket
when Nafion® is sufficiently hydrated145. Nafion® also allows for the facile diffusion of both ions
and molecules and ions through the polymer matrix146.
Figure 2.1. (a) The general representation of Nafion® showing the fluorinated carbon backbone
and hydrophilic sulfonate side chain. (b) The configuration of Nafion® in a hydrated environment
such as a blood sample, where water/ion transport channels are formed144.
Fluoropolymers are commonly used as binding agents in biosensors and biofuel cells, as they
are chemically inert and biocompatible147-150. Nafion®, in particular, has been used as a
binder/stabilizing agent in many different glucose sensor designs20,147,151. It has even proven to
prolong the lifetime of the sensor in comparison to other binding agents147. Nafion® has been
shown to act as an excellent matrix to protect glucose oxidase (GOx) from thermal deactivation
28
during glucose sensing using the temperature pulse amperometry technique, where the temperature
can reach up to 68 °C152, and GOx would lose activity.
Nafion® is also a favorable binding agent for biological systems, as it prevents biological
fouling of electrodes 147,150 and helps repel electroactive species present in the sample that may
otherwise react at the electrode150,153. Lee et al.,154 compared three different glucose sensing films,
with and without Nafion®, and found that, when the films were tested in human plasma and whole
blood solutions, the sensors with chemically bonded Nafion® in the film showed the best sensitivity
and the largest glucose signal, indicating that Nafion® was preventing the sensing film from fouling
in the bio-fluidic solution. Common interferents include uric acid, ascorbate, urea,
acetaminophen150, cysteine and glutathione153. These interferents can react at the electrode, but the
reaction intermediates can also foul the film. Moatti-Sirat et al155., specifically tested five glucose
sensors, with and without Nafion®, in rat and human patients for interfering signals from
acetaminophen. It was observed that those films containing Nafion® had a much lower sensitivity
to acetaminophen. However, some signal from acetaminophen was still observed.
Another benefit of Nafion® is that it has been shown to stabilize and immobilize enzymes.
The Minteer group modified Nafion®, where the size of the hydrophilic channels of the native
Nafion® structure were enlarged (Fig. 2.1b)146. In native Nafion®, the sulfonic acid side chains are
very acidic, which is not favorable for enzyme immobilization. To remedy these issues, the
Minteer group neutralized the sulfonic acid groups by mixing Nafion® with an excess of
hydrophobic alkyl ammonium salts, resulting in large, favorable channels for enzyme
immobilization. Moore et al.,156 found that the addition of quaternary ammonium bromides to
Nafion® helped stabilize a variety of dehydrogenase enzymes immobilized in the Nafion®
membrane, resulting in a high enzyme activity and long enzyme lifetimes of > 45 days.
29
2.4.3 Glucose Oxidase
Clark and Lyons157 were the first to design a commercially successful first-generation
(Scheme 2.2a) glucose sensor (the Yellow Springs Instrument Company Analyzer, Model 23A
YSI) using glucose oxidase (GOx). GOx is an extremely robust glycoprotein commonly extracted
from fungus, typically Aspergillus niger. It is found as a dimer of two identical subunits, with a mass
of 160 kDa and an average size of 5.5 x 7 x 8 nm. Each of the subunits contains a tightly bound
flavin adenine dinucleotide (FAD) prosthetic group located deep within the protein shell (ca. ≥ 1.3
nm from the surface)158. FAD consists of a flavin group (Scheme 2.3) attached to ribitol (a sugar
alcohol), which is bound to the phosphate group of an adenosine diphosphate molecule.
Scheme 2.3. Shows the oxidation and reduction of the flavin group of the flavin adenine dinucleotide
FAD redox active site of GOx37.
FAD is a redox active cofactor that is commonly used to transfer electrons in the metabolism
pathway. Reaction (2.1) shows the two electron transfer between the flavin at the active site of
GOx, and glucose. Once a single reaction has taken place, the FAD site is reduced to FADH2 (as
depicted in Scheme 2.3), rendering the enzyme completely inactive. It is therefore important that an
electron mediating compound is present to re-oxidize the FAD site, thereby reactivating the enzyme
for further glucose detection. Reaction (2.2) shows the regeneration of FAD in the presence of the
mobile mediator, O2.
30
2.4.4 Early glucose sensing work in the Birss group
The Birss group has been focused on sol-derived nanoparticulate Ir oxide (IrOx) films,
demonstrating how their unique properties can be employed as a matrix for glucose detection
(Fourth-generation, Section 2.3). In 2000, Andreas et al159., found that IrOx films with high charge
densities could be formed by depositing an Ir-containing sol solution on a Au substrate. The
Ir(III)/Ir(IV) redox kinetics were rapid, and substantially higher than for IrOx films formed on
standard bulk Ir electrodes. The IrOx nanoparticle (NP) films were found to be highly capacitive
and to have good adhesion with the underlying Au substrate. The NP films differed from bulk IrOx
films in that the hydrogen adsorption/desorption (Hupd) peaks (Section 3.2.1.4) for
nanoparticulate films decrease in size with potential cycling due to the conversion of Ir to IrOx.
At bulk Ir, the Hupd peaks are omniscient, indicating that the underlying metallic Ir sites are still
exposed to solution.
To fabricate the first IrOx NP-based glucose biosensor, Abu-Irhayem et al119,160.,
immobilized glucose oxidase (GOx) within an IrOx matrix by either one of two ways. In the first
method, thin repeating layers of IrOx and then GOx were deposited on a Au substrate. In the
second method, IrOx film growth and GOx immobilization was performed simultaneously. The
resulting electrodes generated H2O2 in the presence of glucose and O2, giving good glucose signals.
Sensing films were later found to produce a small glucose signal in deaerated environments161,
indicating that direct electron transfer through the IrOx NPs was possible. The ability for GOx to
be regenerated via the O2-mediation route and directly through the IrOx NP matrix was termed
“dual-regeneration”162.
Later, GOx powder was mixed directly into an aqueous Ir sol and the resulting ink was
deposited on a sputtered Au substrate by dipping the substrate into the ink and slowly withdrawing
31
it163. It was found that the IrOx redox response was stable in the presence of glucose, and was not
affected by the addition of GOx.
A theoretical model was proposed for the diffusion of glucose into the IrOx/GOx films from
the bulk solution to the active sites embedded deep within the film. The resulting concentration
gradient observed was found to skew the MM plot, resulting in higher Km values. A parameter
similar to the Thiele’s modulus117, (β), was used to describe the effect of GOx loading and film
thickness on the apparent Km value. For example, a large β value was obtained for high loadings
of GOx and when the film thickness was large164.
It was also shown that Nafion® is an important component of sensing films, proposed to
increase the amount of Ir that remains suspended in the aqueous/ethanolic Ir sols with ageing164,
therefore improving IrOx redox kinetics over time. Nafion® was also shown to prevent
electroactive interfering species such as uric acid, ascorbic acid and acetaminophen, from reacting
at the sensing film165, while allowing the oxidation of glucose to take place, similar to work of
others154,166.
2.5 Enzyme kinetics
There are over 6000 currently known enzymes. Some enzymes have single substrates that
they react with, e.g., glucose oxidase (GOx) specifically reacts with glucose. Other enzymes have
low specificity and react with many different substrates167. The specificity of an enzyme depends
on the shape of the enzyme’s active site, and the number of interactions between the amino acids
that make up the active site and the substrate that binds it. In an optimal environment, the enzyme
active site will display maximal activity167, whereas in a non-optimal environment, it can denature
and unfold, rendering it inactive or severely decreasing the enzyme’s activity167.
32
Enzyme activity is defined in terms of enzyme units per volume of enzyme. An enzyme unit
is the amount of substrate that is converted to a product per unit time. The units of enzyme activity
are therefore typically written as nmol per min per ml. The higher the enzyme activity, the faster
the substrate will be converted to product167. Therefore, a high enzyme activity will provide a
sensing film with high sensitivity. As such, when fabricating an enzyme-based biosensor, it is
important to ensure that the enzyme environment within the film is optimal. To ensure an ideal
enzyme environment, many factors must be taken into consideration, including, but not limited to,
pH, water content, temperature, and salt content.
An enzyme acts as a catalyst to accelerate a reaction by decreasing the activation energy
(ΔG‡) of the process, thereby facilitating the formation of the transition state (Figure 2.2). The
activation energy is decreased as a result of the formation of a large number of weak interactions
that exist between the enzyme (E) and the substrate (S). The overall rate of reaction depends on
the activation energy, according to the following equation167:
𝑉 = 𝑘𝐵𝑇
ℎ[S]𝑒−∆𝐺‡ 𝑅𝑇⁄ (Equation 2.1)
where V is the rate of reaction, h is Plank’s constant, kB is the Boltzmann constant, R is the gas
constant, T is the temperature, and [S] is the concentration of substrate.
33
Figure 2.2. Reaction coordinate diagram, depicting how an enzyme decreases the activation
energy of a reaction, allowing for its acceleration, where G = Gibb’s free energy, E = enzyme, S
= substrate, ES = transition state, and P = product.
The general Michaelis-Menten (MM) model for an enzymatic reaction is as follows167:
(Reaction 2.14)
where ES is the enzyme substrate complex, P is the product, and k is a rate constant.
As the enzyme initially binds with the substrate, the concentration of free enzyme decreases
along with the concentration of the substrate with time (Figure 2.3a). Eventually, all of the enzyme
active sites are bound to the substrate and a steady-state is reached. At this steady-state, the
concentration of the enzyme-substrate complex remains constant, the amount of substrate is
depleted, and the amount of product formed increases. It is at this steady-state that a maximum
rate of reaction (Vmax) is achieved, as is shown in the Michaelis-Menten (MM) plot exhibited in
Figure 2.3b.
34
Figure 2.3. (a) The concentration of all of the components in the Michaelis-Menten equation for
an enzymatic reaction with time (b) A typical Michaelis-Menten plot following the rate of reaction
with increasing substrate concentration167.
From the general MM equation, the following equation is given for the rate of reaction:
𝑉0 =𝑉𝑚𝑎𝑥[S]
𝐾𝑚+[𝑆] (𝐸𝑞𝑢𝑎𝑡𝑖𝑜𝑛 2.3)
where Vo is the initial velocity and Km is the Michaelis-Menten constant. When Vmax is reached,
all of the active sites are occupied with substrate, and the addition of more substrate will not affect
Vmax. Km can be defined as the concentration of substrate that will give 1/2 Vmax when reacted with
an enzyme in an optimal environment (Fig. 2.3b). K’m can be calculated by taking the slope of an
Eadie-Hofstee plot168 (V0[S] vs. V0) and can be derived from the general MM equation as being:
𝐾𝑚 =𝑘−1+𝑘2
𝑘1 (𝐸𝑞𝑢𝑎𝑡𝑖𝑜𝑛 2.4)
The experimentally observed Km value is normally represented by the apparent Km value
(i.e., K’m), as for sensors, Km can be additionally influenced by factors such as the rate of diffusion
of substrate to the electrode and mediator diffusion rates169. In this work, the velocity (or rate) of
reaction is denoted by i, or current density, as the glucose concentration is proportional to the
35
number of electrons collected by the transducer. The rate at which the electrons are collected (the
current passed, i) is equivalent to the reaction velocity.
2.6 Techniques used in this work
2.6.1 Quartz crystal microbalance (QCMB) technique
The quartz crystal microbalance (QCMB) technique is an extremely sensitive tool for the
measurement of small mass changes per unit area (as low as in the nanogram range)1. This
technique can be used for studying interfacial interactions, such as oxide formation, corrosion, the
adsorption of thin films, or the immobilization of a biological element to a substrate. The QCMB
technique can also be used as a method of detection of biological analytes (biomarkers, cells,
pathogens, sugars, etc.), as well as metals, pollutants, or gases, etc. Tang et al.170 designed a
displacement-type assay sensor where concanavalin A was immobilized on a dextran/carbon
substrate, deposited on a quartz crystal (QC). Glucose then competes with dextran to bind the
concanavalin A sites, thereby displacing dextran, leading to a change in mass (with a detection
limit of 5.0 µM glucose). Uluda and Tothill used Au NPs and antibodies deposited on a QCMB
probe for the detection of prostate-specific antigen in serum23. In some other examples biological
components immobilized to SAMs on a QC have been used for the specific detection of analytes,
such as pathogenic bacteria171, or epidermal growth factor receptors172.
For QCMB analysis, a metal is deposited on both sides of the QC, in a “keyhole pattern”, as
depicted in Fig. 2.4a so that an alternating potential can be applied to both sides of the crystal,
resulting in shear deformation of the crystal, or an oscillation (Fig. 2.4b). QCs are typically AT-
cut as these cuts have a nearly zero temperature coefficient. The QCs used in this work are 1-inch
in diameter and oscillate at 5 MHz, and the Sauerbrey equation (2.5) can be simplified so that for
a 56 HZ change in frequency, 1 µg cm-2 has been lost or deposited on the crystal1.
36
Figure 2.4 (a) The Au keyhole pattern deposited on a quartz crystal used in the quartz crystal
microbalance technique, and (b) side view of a quartz crystal disk with an exaggerated view of
the shear distortion resulting from the oscillation1.
Sauerbrey demonstrated that the frequency at which a QC oscillates will depend on the
change in the mass on the crystal surface173. If a voltage is applied to a quartz crystal, it will
oscillate at a specific frequency due to the piezoelectric effect. As the mass of the crystal changes,
the frequency also fluctuates, according to the Sauerbrey equation173:
∆f(Hz) = −2f0
2
(μqρq)12⁄×∆m
A= −Cf (constant) ×
∆m (μg)
A (cm2) (𝐸𝑞𝑢𝑎𝑡𝑖𝑜𝑛 2.5)
where ∆f represents the change in frequency, f0 is the resonant frequency of the crystal, μq is the
shear modulus, ρq is the quartz density, ∆m/A is the mass change per unit area, and Cf is the
sensitivity factor. As can be seen in Equation 2.5, the addition of a small mass on the crystal surface
results in a decrease in the frequency at which the crystal oscillates, whereas, the removal of mass
would increase the oscillation frequency.
2.6.2 Cyclic voltammetry and amperometry
Cyclic voltammetry (CV) is one of the most commonly utilized electrochemical techniques.
This technique can be used in many different areas of study, including for the analytical detection
or quantification as well as identification of redox species in a solution or immobilized to an
37
electrode surface. CV can also be used for the determination of both kinetic and thermodynamic
properties of specific electrochemical systems.
In this work, a potentiostat and a three electrode (a working (WE), reference (RE) and
counter electrode (CE)) circuit is used (Section 3.2). A particular potential (E) is applied to the
WE vs. the RE. The purpose of the third electrode, the (CE), is to pass current to the WE. A
triangular potential waveform is applied to the WE between a lower potential, E-, and an upper
potential, E+, at a specific sweep rate. Cyclic voltammetry at varying sweep rates can give
information regarding the kinetics and mode of mass transport of an electrochemical system.
When the potential is swept from E- to E+, oxidation takes place at the WE and an increase
in the oxidative current is observed, and vice versa in the negative sweep, where reductive
processes occur. The resulting CV, i.e., the plot of current vs. WE voltage, then depicts at which
potentials specific redox chemistry occurs, and the charge passed can give the number of moles of
redox-active material in the electrochemical reaction 174.
In amperometry, the potential of the WE is held constant (vs. the RE) and the current is
monitored as a function of time. In this work, the potential at the WE, and therefore the sensing
film, was fixed at 1.2 V, while aliquots of glucose were added to the cell solution. As the
concentration of glucose increases (with time), the oxidative currents were tracked, and from this
i/t plot, the Michaelis-Menten (MM) plot can be derived. The MM plot is commonly used in to
study the kinetics of enzyme-substrate binding and product formation (Section 2.5).
2.6.3 Scanning and transmission electron microscopy
The scanning electron microscope (SEM) involves a high-energy electron beam focused on
a sample surface, and when the electrons bombard the sample, energy is dissipated as secondary
electrons (SEs) or back-scattered electrons (BSEs), as well as in other forms, such as Auger
38
electrons and X-Rays. SEs escape from near the surface of the sample, giving topological and
morphological information. BSEs, on the other hand, are omitted (reflected primary electrons)
from deeper within the sample, and the yield of the collected BSEs increases monotonically with
the atomic number of the sample. The greater the energy of the primary electron beam, the larger
the number of SEs and BSEs emitted175.
Transmission electron microscopy (TEM) is a technique that can be used to view objects at
the order of a few angstroms in size (at a higher resolution than SEM images). Samples being
imaged by TEM are typically deposited on a carbon/copper TEM grid, and must be thin enough to
allow for electron transmission through the sample176. A light source at the top of the microscope
emits electrons, which then travel through a vacuum in the column of the microscope before being
focused to form an electron beam by an electromagnetic lens. The electron beam then travels
toward the specimen and, depending on the density of the specimen, some electrons will be
scattered when they come into contact with the sample, while others go directly through. The
electrons that were not scattered during the process hit a fluorescent screen below the sample. This
gives rise to a shadow image of the specimen, which is photographed by a camera.
2.6.4 X-Ray photoelectron spectroscopy
X-Ray Photoelectron Spectroscopy (XPS) was first developed in the mid-1960’s by the
Siegbahn group. Later in 1981, Dr. Siegbahn was recognized for his contribution and awarded
with a Nobel Prize in Physics177. XPS is a non-destructive technique, and can be used for
determining the elemental composition of sample surfaces for a wide array of applications. XPS
is commonly used in the fields of corrosion, semiconductors, thin film coatings, dielectric
materials, catalysts and polymer surfaces, to name a few.
39
XPS involves irradiating a sample surface (in an ultra-high vacuum) with a monoenergetic
X-Ray beam, either Mg Kα (1253.6 eV) or AlKα (1486.6 eV). These X-Rays bombard the core
electrons of the atoms to a depth ca. 1 micrometer, resulting in the ejection of photoelectrons.
Typically, however, the information garnered from the XPS spectrum is only from the outer 1 –
10 nm of the sample. The core electrons are closest to the nucleus and have binding energies (BE)
that are characteristic of each atom.
The BE is defined as the amount of energy that is required to remove a core electron from
the nucleus of an atom, and is calculated with respect to the Fermi level (0 energy of attraction
between the electron and the nucleus). There is a simple relationship that relates the kinetic energy
(KE) of the detected core electrons and the BE, as shown in equation 2.6 below178:
𝐾𝐸 = ℎ𝜈 − 𝐵𝐸 − 𝜙 (𝐸𝑞𝑢𝑎𝑡𝑖𝑜𝑛 2.6)
where hν is the photon energy from the X-Ray source and 𝜙 is the spectrometer work function.
The KE of the emitted electrons is measured by a cylindrical mirror analyzer. The cylindrical
mirror analyzer has two concentric metal cylinders, one at a positive voltage and another at 0 V,
which generates an electric field between the two cylinders. Electrons with the correct velocity
will pass through the cylinders to the detector. If an electron’s velocity is too slow or too fast, they
will collide with the cylinders, and will not pass through to the detector. By manipulating the
voltage of the cylinders, the field can be varied so that only electrons of a specific KE will pass
through the detector. The number of electrons collected provides a quantitative analysis of the
element of interest. It is also possible to determine the oxidation state of an element, due to the
shift in the BE. Atanasoskaa et al179., used XPS to study the surface composition and ratios of the
Ir-O σ and Ir=O π bonds, in particular, of an IrO2 coating for a cardiovascular stint.
40
2.6.5 X-Ray diffraction
In X-ray diffraction (XRD), monochromatic X-rays (generated by a cathode ray tube) of a
particular wavelength are used to bombard a crystalline sample. When the incident X-rays
bombard the sample, there will be constructive interference between the diffracted rays that will
satisfy Bragg’s Law180:
d = nλ
2 sinθ (𝐸𝑞𝑢𝑎𝑡𝑖𝑜𝑛 2.7)
where n is an integer, λ is the X-ray wavelength, θ is the angle between the incident X-ray and the
scattering planes, and d is the inter-planar spacing of the crystalline sample.
It is possible to identify crystalline materials by XRD, as each crystalline material has a
unique d-spacing (unit cell dimensions), like a finger-print, which can be calculated from Bragg’s
Law and compared to a database. By scanning the sample over a wide range of 2θ angles, it is
possible to obtain all of the lattice d-spacings of the sample.
41
CHAPTER THREE: EXPERIMENTAL METHODS
3.1 Preparation of Ir sols and sensing inks
3.1.1 Preparation of ethanolic Ir sol-based inks
Ir sol-based inks were made by mixing one mole of Ir (III) chloride (IrCl3, Alfa Aesar,
99.99 % pure) with three moles of sodium ethoxide (NaOC2H5, Aldrich, 96% pure) in 10 mL of
absolute ethanol (Aldrich, 99.5% pure) and refluxing at 75 °C under stirred, deaerated conditions
for two hours. The resulting ink was then cooled to room temperature and allowed to stir (under
deaerated conditions) for ca. 18 hours. The ink was then filtered through a fritted funnel,
transferred to a sealed glass vial, and stored in the fridge at 4 °C to minimize the time the NPs
are exposed to air, light, or heat112,159. Ir inks were stored for a maximum of 6 months before a
fresh sol was prepared. The Ir ink was always sonicated for at least 45 minutes prior to each use.
3.1.2 Preparation of Ir/Nafion®/GOx sensing film via aliquot deposition
3.1.2.1 Preparation of GOx solution
Glucose oxidase (GOx) solutions (60 g l-1) were typically prepared by dissolving small
amounts of lyophilized GOx powder (Sigma, 17 300 U/g) in either a 0.1 M, pH 7 phosphate buffer
solution (PBS) or a 0.1 M acetate buffer (pH 4) solution. Solutions were typically 60 g l-1, and
were stored at 4 ºC until use. Lyophilized enzyme was stored in the freezer.
3.1.2.2 Preparation of Nafion® solution
Nafion® solutions (Typically 1 wt. % Nafion®) were prepared by mixing liquid Nafion® (15
wt. % Nafion®, Ion Power Inc, 950 EW) with absolute ethanol. Solutions were stored in a sealed
vial at 4 ºC.
42
3.1.2.3 Aliquot deposition of sensing ink on Au substrate
Aqueous glucose oxidase (GOx), ethanolic Nafion®, and ethanolic Ir sol were gently mixed
together (typically in a 1:1:4 µL ratio, or a 1:50 mole ratio of GOx:Ir) in a small vial by drawing
the solution in and out of an Eppendorf pipette, taking care to not produce bubbles. In Section 5.5,
different ratios of GOx:Ir were also used, ranging from 1:25 to 1:200. In some experiments,
additional aliquots of 0.1 M, pH 7 PBS or absolute ethanol were added to the sensing inks in order
to alter the ratio of ethanol to water. Also, in some cases, 0.1 M, pH 7 PBS was added to an ink in
place of GOx, or absolute ethanol was added to the ink in place of Nafion® or Ir sol. Inks were
deposited on the Au substrate immediately after mixing.
The ink mixtures (typically 4 µL) were deposited on a substrate (Au-sputtered slides, carbon
or polished Au foil) by either aliquot deposition via Eppendorf pipette, or by the diazonium salt
reduction method (Section 3.1.3). The aliquot deposition method consisted of depositing an aliquot
(1-4 µl) of either the Ir sol, Ir/Nafion®, or the Ir/Nafion®/GOx inks on the Au slide over a total
surface area of ca. 0.25 cm2 (typically 2.7 µm thickness, with thickness ranging from 0.66 µm to
4.0 µm). The films were then covered and air dried at room temperature for at least 15 hours.
3.1.3 Sensing film fabrication using diazonium salt chemistry
3.1.3.1 Diazonium salt grafting and GOx immobilization
Diazonium salt solutions (1.0 mM nitrophenyldiazonium tetrafluoroborate, Sigma Aldrich,
97 % purity) were prepared using 0.1 M tetrabutylammonium tetrafluoroborate (Sigma Aldrich,
99 % purity) as the electrolyte. The first and most commonly used solvent was acetonitrile (ACN,
Sigma-Aldrich, 99.9 % purity). The nitrophenyl group was grafted to the Au or carbon
(electrochemically cleaned) substrate by cycling the potential reductively (0.6 to -0.2 V vs. pseudo
Ag/Ag2O), or applying a constant potential (-0.2 V vs. pseudo Ag/ Ag2O) in the diazonium salt
43
solution. After grafting, the electrode was sonicated in ACN, then rinsed with acetone (EMD, ACS
grade), isopropanol (EMD, ACS grade), methanol (EMD, ACS grade) and triply distilled water.
The nitro group was subsequently converted to an amine by a 6 electron electrochemical reduction
in 0.1 M KCl (EM Science, ACS grade) / 10 vol. % methanol or ethanol solution181,182.
A succinic anhydride linker was then attached to the phenyl group, via amide bond
formation, by soaking the aminophenyl modified Au or C electrode in 0.1 M succinic anhydride
(Sigma Aldrich, > 99 % purity) in dimethylformamide (DMF, Sigma-Aldrich¸ ACS grade) under
deaerated, stirred conditions for two hours181. The carboxylic acid functional group was then
activated by soaking in 1.0 mM sulfo-N-hydroxysuccinimide (sulfo-NHS, Sigma Aldrich, > 98.5
% purity) and 10 mM ethyl-3-[3-dimethylaminopropyl] carbodiimide hydrochloric acid (EDC,
Sigma Aldrich, >99 % purity) in 0.1 M 2-(N-morpholino)ethanesulfonic acid buffer (pH 5.5) for
one hour under deaerated, stirred conditions181,183,184. The film was then transferred to a solution
of glucose oxidase (typically 1.0 g l-1 in pH 4, 0.1 M acetate buffer) and allowed to soak for at
least 17 hours. Electrodes were then rinsed several times with distilled water, dried in air at room
temperature, and characterized.
3.1.3.2 Infiltration of IrOx/Nafion® into immobilized GOx layer
The ethanolic Ir/1 wt. % Nafion® inks (4:1 volume ratio of Ir sol:Nafion®) were sonicated
for 45 minutes before a typical aliquot of 4 µL was deposited on the surface of the immobilized
glucose oxidase film. The film dried for at least 1 hour in air, at room temperature before the Ir
film was rinsed, electrochemically converted to Ir oxide (IrOx), and then tested for its response to
glucose.
44
3.2 Electrochemical characterization
3.2.1 Working electrodes (WEs)
3.2.1.1 Au working electrodes
Au working electrodes consisted of sputter-coated Au slides or Au foil substrates, coated
with a thin film (of varying thicknesses) of the sensing ink. Au-coated microscope slides were
purchased from Deposition Research Lab, Inc. The Au-coated slides consisted of a 10 nm Ti
adhesion layer was sputter-coated on the clean borosilicate slide prior to the deposition of a 100
nm thick Au sputtered layer. The microscope slides (1” x 3” x 1.1 mm) were then cut into 0.5 cm
x 1 inch strips using a diamond glass cutting saw. The same Au-coated microscope slides were
also fabricated in house. Microscope slides (Corning) were treated with a 3:1 solution of
concentrated H2SO4 and 30 wt. % H2O2 (Aldrich, ACS grade) for 2-3 hours to remove any residual
organic impurities. Slides were then rinsed with triply distilled water and isopropanol (Sigma, >
95 % purity) before drying in air. The Ti (10 nm) and Au (100 nm) layers were then sputter-coated
on top of the slides using a Denton DV-502A High Vacuum with a Sorensen DCS 600 - 1.7 Power
Supply unit. All slides were cut into ca. 0.5 cm x 2.5 cm pieces, referred to as the “Au substrate”
in this thesis.
Circular Au foil samples were supplied by the University of Warsaw. Circular Au foil
attached to a thin 0.25 mm diameter Au wire were made by heating the Au wire (Sigma, 99.99 %
purity) over a Bunsen burner to form a Au globule at the end of the wire. The ball was then flattened
to a diameter of ca. 3.0 mm and a width of 0.5 mm. The surface area of the Au foil was increased
by anodizing the Au to form Au black185 in stirred, 0.3 M oxalic acid. Linear sweep voltammetry
was performed from 0.27 to 2.77 V vs. RHE at a sweep rate of 0.5 mV s-1, and the Au electrodes
45
were subsequently held at 2.77 V vs. RHE for 1250 to 2500 s. Au foil strips (ca.0.3 cm x 2.5 cm,
0.127 mm thick Sigma Aldrich, 99.99% purity) were also used, but were not anodized.
All Au substrates were suspended in solution using Cu clips which were soldered to Cu wires
encased in a sealed glass tube. The Cu did not come into contact with solution as it was both kept
above the meniscus and it was tightly wrapped in Parafilm™. Creeping of solution onto the Cu was
not observed.
3.2.1.2 Carbon working electrodes
Various types of carbon were used as the working electrode for the immobilization of
glucose oxidase (via a diazonium-derived film). Carbon paper (CP, Fuelcellstore.com, Spectracarb
2050 A) was cut into 0.5 cm x 0.5 cm strips and held securely in place on a sputtered Au substrate
on a borosilicate slide (Section 3.2.1.1) using carbon tape. CP consists of graphitized, resin-
bonded, carbon microfibers pressed together to form a compact, high surface area surface (380 μm
sheets with a porosity of 78%) material. These paper-thin sheets are commonly used in proton
exchange membrane fuel cells and electrolysers186.
The sputtered Au substrate was used as a rigid, conductive substrate, to supply a sturdy
support for the fragile CP.
3.2.1.3 Electrochemical cleaning of working electrodes
Au substrates were cleaned prior to film deposition by first rinsing with isopropanol,
methanol, and water. The Au was then subjected to electrochemical cleaning in 0.5 M H2SO4 by
cycling from 0.05 V to 1.7 V vs. RHE until the Au oxide reduction peak no longer fluctuated in
size or shape.
46
Carbon substrates were cleaned in a similar manner by cycling the potential in 0.5 M H2SO4
from 0.05 V to 1.0 V vs. RHE until the carbon cyclic voltammetry features remained constant in
size and shape.
3.2.1.4 Real Ir and Au surface area determination
The real Ir surface area was determined from the charge passed in the hydrogen
underpotential deposition (Hupd) region (0.05 to 0.3 V vs. RHE) of the cyclic voltammograms in
0.1 M PBS at a sweep rate of 100 mV s-1 (Fig. 3.1a), according to the equation below:
𝑄𝑐 = (i x E) / ʋ (𝐸𝑞𝑢𝑎𝑡𝑖𝑜𝑛 3.1)
𝑆𝐴𝑅𝐸𝐷 =Q𝑅𝐸𝐷
x mC 𝑐𝑚−2 (𝐸𝑞𝑢𝑎𝑡𝑖𝑜𝑛 3.2)
The electrochemical active surface area was estimated from the charge passed (Qc) in the
Hupd region 0.137 mC (for 65% of 1 monolayer) for 1 real cm2 of Ir (Equation 3.2)187. A typical
roughness factor for an Ir film is ca. 50.
The real Au surface area was determined from the reduction peak of the Au oxide (Fig. 3.1b)
in 0.5 M H2SO4 at a sweep rate of 100 mV s-1, using Equations 3.1 and 3.2 where an average value
of ~400 µC cm-2 was used for the reduction of a monolayer of gold oxide (Au2O3) on
polycrystalline Au, when the Au was cycled negatively from 1.7 V vs. RHE188,189. A value of 216
µC cm-2 was calculated for the reduction of one monolayer of Au2O3 when the potential was cycled
negatively from 1.5 V vs. RHE. The real surface area, or roughness factor was calculated to be ca.
1.5 times the calculated geometric area of the Au surface for sputtered Au substrates.
47
Figure 3.1. Cyclic voltammetry response of (a) an Ir film and (b) a Au film in a deaerated, stirred,
0.5 M H2SO4 solution, at 100 mV s-1. The purple shading indicates the area used to determine the
real surface area, where the area under the cathodic peak should be equivalent to the area under
the anodic peak.
3.2.2 Counter Electrodes (CE)
A Pt black counter electrode was used for all electrochemical experiments. Pt black was
prepared by platinizing a Pt gauze (99.9 % pure, Alfa Aesar) electrode using a Yellow Springs
Instruments 3139 platinizing kit. The platinizing solution consisted of 3.47 g H2PtCl6•6 H2O
(Sigma Aldrich, 99.9%), and 0.015 g Pb(C2H3O2)4 (Sigma Aldrich, ACS grade) dissolved in 60
mL of 3X distilled H2O. Two Pt gauze electrodes were placed in the platinizing solution (ca. 5 cm
apart) and the polarity of the current between the electrodes (constantly 50 mA) was reversed every
30 s for a total of 3 min.
3.2.3 Reference Electrodes (RE)
A reversible hydrogen electrode (RHE) was used for most of the electrochemical studies in
this work. The RHE consisted of a platinized Pt black electrode (identical to the CE) immersed in
H2-saturated 0.5 M H2SO4 or 0.1 M, pH 7 PBS. All of the electrochemical data in this thesis are
referenced to the RHE, excluding the work related to the electrochemical grafting of the diazonium
0.0 0.4 0.8 1.2
-30
-20
-10
0
10
20
30
IrO2 + e
-+ H
+ H
2O
+ Ir
(a)
i /
mA
cm
-2
E / V vs. RHE
H+ + e
- H
(ads)
H(ads)
H+ + e
-
H2O
+ Ir IrO
2 + e
-+ H
+
0.0 0.4 0.8 1.2 1.6-0.5
-0.3
-0.2
0.0
0.2
2Au + 3H2O Au
2O
3 + 6H
+ + 6e
-
Au2O
3 + 6H
+ + 6e
- 2Au + 3H
2O
I /
mA
cm
-2
E / V vs. RHE
(b)
48
salts and the subsequent reduction of the grafted moiety, as the RHE requires a large solution
volume, and a smaller electrochemical cell could be used with a Ag/Ag2O pseudo-RE instead.
A Ag/Ag2O pseudo-reference electrode was used when grafting the nitrophenyl groups to a
conductive substrate, as well as for the subsequent reduction of the nitro to an amino group (in 1.0
mM nitrophenyldiazonium tetrafluoroborate/0.1 M tetrabutylammonium tetrafluoroborate/ACN,
and 0.1 M KCl/10 vol. % MeOH solutions, respectively)181. The Ag/Ag2O pseudo-reference
electrode was fabricated by dipping a 1.0 mm Ag wire (Sigma Aldrich, 99.9 % purity) in a 1:1
solution of nitric acid (EMD, ACS grade):H2O for 15 s prior to each usage. This pseudo-reference
electrode was found to be extremely stable during the course of an experiment, and also between
experiments.
3.2.4 Electrochemical methods
3.2.4.1 Electrochemical cell
A three-electrode two compartment cell setup was employed for the oxidation of Ir to Iridium
oxide, anodization of Au, and the electrochemical electrode cleaning and characterization of the
films. The CE, WE and gas bubbler all entered the cell from the top of the main compartment (25
mL), while the RE was placed in a second compartment, connected to the main compartment via
a Luggin capillary. The main compartment was placed above a stir plate. Glucose additions were
performed through a sealed needle port on the side of the cell that allowed glucose to be injected
directly into the path of the stir bar. The electrochemical cell setup is shown in Figure 3.2a.
The electrochemical cell for the grafting of the diazonium salt to a conductive substrate and
the subsequent reduction of the grafted moiety is shown in Figure 3.2b. Four airtight holes were
drilled into a rubber stopper, and the CE, WE, RE and a gas needle were inserted through them.
The stopper was then inserted into a 20 mL vial containing 5 mL of either the grafting diazonium
49
Figure 3.2. Electrochemical setup used for (a) substrate cleaning, film characterization, and
glucose additions, and (b) diazonium salt reduction and the subsequent reduction of the nitro- to
an amino group.
salt solution or the aqueous KCl/MeOH reduction solution (Section 3.1.3.1). Agitation of the
solution was accomplished by bubbling of Ar through the solution.
50
3.2.4.2 Solutions
All solutions in this work were made with triply distilled water. Phosphate buffer solution
(pH 7, 0.1 M, PBS) was used for glucose concentration analysis and the oxidation of Ir to form
IrOx. Neutral PBS was made by mixing 0.2 M Na2HPO4 (J.T. Baker Chemical Co., ACS grade)
and 0.2 NaH2PO4 (Fisher Scientific Co., ACS grade) together according to the Henderson-
Hasselbalch equation167. Sulfuric acid (0.5 M) was used to electrochemically clean all of the
sputtered Au electrodes, and in some cases, it was used for Ir oxidation as well. 0.5 M sulfuric acid
was made by diluting concentrated sulfuric acid (EMD, ACS grade). In Section 7.5.3, 0.3 M oxalic
acid (J.T. Baker Chemical Co., ACS grade) was used to anodize Au foil to Au black. β-D-Glucose
(Sigma, 99.5% purity) solutions for glucose concentration determination experiments were made
up to 2.0 M concentration. The solutions were made 2 hours prior to testing to ensure proper
mutarotation to the preferred β-D-glucose anomer. Any remaining glucose was disposed of after
testing to prevent bacterial contamination. Aliquots of glucose (typically 10 µl) were added to the
PBS using a 10 µL Hamiltonian needle.
3.2.4.3 Environment
To ensure accurate and reproducible potential values, H2 was purged through the RHE
solution for at least for 15 minutes prior to any electrochemical experiments. Similarly, for
deaerated and aerated experiments, Ar and O2, respectively, were purged through the testing
solution for at least 15 minutes prior to any electrochemical experiments. To ensure that solutions
were completely deaerated, a cyclic voltammogram of Pt electrochemistry in the Ar-saturated
solution was performed to confirm the absence of an O2. A second experiment, performed to
exhibit O2-independence, showed that during a glucose addition experiment in a deaerated
51
environment, when O2 was bubbled into solution mid experiment, the glucose signal was not
affected.
3.2.4.4 Electrochemical instrumentation and software
All of the electrochemical experiments were carried out using a Multistat 656 Potentiostat
(Solartron). All data were analysed and recorded using CorrView™. Microsoft Excel and Origin
Pro 8.0 (Student version) were used to analyze data and produce the figures presented in this thesis,
as well as to integrate the areas under peaks of cyclic voltammograms.
The K’m and imax Michaelis-Menten (MM) values (Section 2.5) were calculated from the
chronoamperometric data using an Excel program (with the solver add-in)190, which performs
iterations in combination with the MM equations to solve for the kinetic values. K’m values were
compared to those calculated from Eadie Hofstee plots (Section 2.5).
3.2.5 Glucose addition experiments
Typically, all of the electrochemical experiments were performed under a constant flow of
Ar (Praxair, Research grade 6.0, ≥ 99.9 % pure, O2 < 0.2 ppm). For glucose testing in aerated
environments, O2 (Praxair, Research grade 5.0, ≥ 99.9 % pure) was constantly flowed through the
PBS. Experiments were performed under constant stirring conditions using a magnetic stir plate
and stir bar. Prior to each experiment, the solution was saturated with either gas for a minimum of
15 minutes to ensure saturation. Typically, 10 µL of 2.0 M D-+-Glucose were injected into 25 mL
0.1 M, pH 7 PBS at time intervals of 50 s.
3.3 Other techniques
3.3.1 Transmission electron microscopy (TEM)
All Transmission Electron Microscopy (TEM) work was carried out on a Tecnai TF20 G2
FEG-TEM (FEI, Hillsboro, Oregon, USA) in the Microscopy and Imaging Facility (Health
52
Sciences Centre) at the University of Calgary. The samples were prepared by suspending diluted
Ir sols (50x dilutions) in EtOH or EtOH/H2O and sonicating the resulting inks for 40 min. Samples
containing enzyme were not sonicated to ensure that any enzyme-metal/metal oxide conjugates
were not disturbed. An aliquot (0.5 µl) of the resulting suspensions was deposited on top of a
carbon-coated Cu 400 TEM grid (EMS) (2 mm). Imaging was performed in bright field mode.
ImageJ software was used for the analysis of TEM images to determine the size of the Ir
nanoparticles.
3.3.2 Scanning electron microscopy (SEM)
A Philips/FEI XL 30 unit in the Microscopy and Imaging Facility (Health Sciences Centre)
at the University of Calgary was used for to study the morphology of the Ir/Nafion®/glucose
oxidase films. The working electrodes were glued to Al stubs using double-sided carbon tape.
Graphite paint was then used to electrically connect the films to the Al stub. Images were collected
with an accelerating voltage of 15-20 kV, at a working distance of ca. 10 mm. Samples did not
exhibit charging effects.
3.3.3 Quartz Crystal Microbalance (QCMB) technique
In these experiments, the WE was a 2.5 cm (diameter) AT cut quartz crystal (Valpey-Fisher),
first sputter-coated with a Ti adhesion layer (ca. 10 nm), followed by Au (ca. 100 nm), all using a
Denton DV-502A high vacuum sputterer. The electrochemically active geometric surface area of
the Au WE was ca. 0.48 cm2. Electrical contact was made to each side of the crystal using
conducting silver epoxy and Au wires (as shown in Fig 3.3a).
A three electrode, two-compartment glass cell was used for the quartz crystal microbalance
experiments (QCMB), similar to the electrochemical setup discussed in Section 3.2.4.1. The
working electrode (WE, quartz crystal) was inserted between two rubber O-rings at the base of the
53
Figure 3.3. A photograph of the components of the electrochemical/QCMB cell, in the (a)
disassembled and (b) assembled state.
main cell compartment, as shown in Fig. 3.3b, allowing for one side of the WE to be exposed to
solution, while the other side was open to air. The Pt-black counter electrode (CE) was placed in
the solution above the WE and the reference electrode (RE) was placed in a second reference
hydrogen electrode (RHE) compartment, identical to what is depicted in Fig 3.5b.
54
An EG&G PARC 173 potentiostat, combined with a PARC 175 function generator, was used
for QCMB measurements. The mass changes accompanying the electrografting of nitrophenyl
groups to the Au/C substrates from the diazonium salt and the subsequent immobilization of GOx
were monitored using both the in-situ and dry QCMB techniques, employing a Philips PM6654C
high resolution counter-timer and a Pierce-type oscillator. Powerlab 400 was used for the
frequency change and electrochemical data collection, while Chart for Windows v. 5 was used to
plot the data.
3.3.4 X-Ray photoelectron spectroscopy (XPS)
X-Ray photoelectron spectroscopy (XPS) using a PHI VersaProbe 5000 (XPS-Physical
Electronics, and Catalysis Surface Science Laboratory, University of Calgary) was used to
determine whether Ir had completely converted to IrOx after electrochemical oxidation in the
presence and absence of GOx. CasaXPS software was used to fit the XPS spectral data.
3.3.5 X-Ray Diffraction (XRD)
X-ray diffraction (XRD) using a Rigaku Multiflex X-ray Diffractometer (Department of
Geoscience, University of Calgary) with a CuKα radiation source (λ = 1.5406 nm) and operating
at 40 kV and 20 mA, was employed for the determination of the crystal structure and size of the Ir
NPs in the ethanolic ink, as deposited from an ethanolic ink. The ethanolic Ir ink was dried and
ground into a powder. The sample was then loaded on an amorphous glass holder (with a 2.5 cm
diameter). The XRD pattern of the ink was recorded over 0 °- 90 ° 2θ values at a rate of 1 °/min.
3.4 Error analysis
The primary source of error encountered in this thesis work is the measurement error
associated with electrochemical instrumentation, film fabrication reproducibility, and
reproducibility between enzyme batches and sol synthesis.
55
For all electrochemical experiments, the reported results are the average value of
experiments replicated in triplicate. The error (standard deviation) was always < 15 %, unless
otherwise stated. Cyclic voltammetry (CV) was performed at multiple sweep rates, however, the
majority of the experiments were performed at 100 mV s-1. To ensure this sweep rate was not too
fast, CVs were performed at 10 mV s-1 and the peak currents were compared to ensure a 10-fold
increase was observed between the CVs performed at 10 vs. 100 mV s-1.
For the measured currents, an error of 0.2 % ± 0.1 % (error in measured current ± error in
potentiostat current multiplier) was estimated. Glucose concentrations were estimated to be within
± 2 % of the calculated concentration. Error due to electrical noise during glucose testing was ca.
0.3 µA cm-2. The signal to noise ratio was found to remain constant at roughly 30 with increasing
glucose concentration.
Solution volumes from 5.0 to 50 ml were measured using 10 ± 0.1 ml, 25 ± 0.1 ml or 50 ±
0.4 mL graduated cylinders. Solution volumes less than 5.0 ml, were dispensed using Eppendorf
pipettes of varying volume ranges (1-10 μL ± 2.5 % and 100-1000 μL ± 1.2 %). Glucose injections
were performed with a 10 μL Hamiltonian pipette with a calculated uncertainty of 4.96 %.
In the quartz crystal microbalance experiments, the geometric surface area of the exposed
Au working electrode was 0.48 ± 0.02 cm2. A literature-based conversion factor of -56.6 Hz µg-1
cm2 was used to calculate the mass change from the frequency changes, however, no calibration
experiments were performed to confirm the accuracy of this conversion factor. There is a ± 5 %
error in the transmission electron spectroscopy scales, therefore, there is also a ± 5 % error in the
calculation of the average nanoparticle size. The X-ray diffraction instrument has an instrument
alignment within ± 0.01° for 2 theta over the whole angular range191. Calibration was performed
56
by using NIST corundum standard SRM 1976a. The X-ray photoelectron spectroscopy instrument
has a typical error in peak position in the ranges of ±0.2 eV192.
57
CHAPTER FOUR: UNDERSTANDING IRIDIUM-BASED INK DRYING AND
ELECTROCHEMICAL PROPERTIES*
*Section 4.5.2 was published in Campbell, H.B.; Elzanowska, H.; Birss, V. I. Biosensors and
Bioelectronics 2013, 42, 7.
4.1 Introduction
Recently, the fabrication of electrochemical biosensors has been focused on designing
supporting matrix architectures for the immobilization of redox-active enzymes, such as glucose
oxidase (GOx). As examples, Au193, Pt194 and Ni195 nanoparticles (NPs), as well as carbon
nanotubes196, and conductive organic polymers, such as poly(thiophene)s197, have been used.
Characteristics of a good matrix for a biosensor electrode include stability, a high surface area for
maximal enzyme deposition, high porosity (to allow access to the analyte), and good electronic
conductivity.
Nanoparticulate Ir and Ir oxide (IrOx) films have been proven to be exemplary matrices for
fourth-generation, glucose biosensing applications (Section 2.4.1), as regeneration of the glucose
oxidase (GOx) active site after the reaction with a glucose molecule can occur directly via the
interconnected IrOx matrix, or via the O2-mediation route9,10 (dual-mediation, Section 2.4.4). To
achieve reproducible glucose sensing, the IrOx NP film morphology must be uniform and well-
interconnected (fast electron transfer kinetics), and all GOx molecules must be in good contact
with the IrOx NP matrix. In order to prevent GOx or IrOx aggregation in the sensing film,
knowledge must be garnered concerning how to fabricate the optimal, well-interconnected sensing
film matrix. The purpose of Chapter 4 is therefore to understand how the interactions between the
IrOx NPs, the water/ethanol solvent and Nafion® affect the IrOx NP interconnectivity and redox
kinetics, as well as the sensing film morphology.
58
The benefits of fabricating IrOx NP films from Ir NP suspensions (inks) include the very
high surface area that can be achieved112,198, as well as the flexibility of the support materials that
can be employed. One disadvantage to using an Ir NP ink, however, is that it must be synthesized
in pure ethanol to maintain small NP sizes. GOx is insoluble in most organic solvents199, and, as
such, water is essential for enzyme activity. As a consequence, dissolving a hydrophilic enzyme
in an organic solvent (such as the ethanolic Ir ink) can lead to protein unfolding, or
denaturation200,201.
There are a few papers in the literature that describe how a colloidal suspension of metallic
NPs will interact with varying water/ethanol mixtures during NP synthesis 202,203 and Nafion®204,205
during NP synthesis204, in a NP ink203, and during the process of NP film fabrication205. This
chapter also focuses on how water/ethanol mixtures affect film morphology, stability and the IrOx
redox kinetics of films fabricated from these inks, and how the addition of Nafion® to the ink
affects these film characteristics. To the best of our knowledge, the work described here is the first
to provide a detailed analysis on how 1-2 nm metal NPs are affected by the addition of water to an
ethanolic ink (after NP synthesis).
It is shown here that water addition to an ethanolic ink does not significantly affect the
overall redox kinetics of the IrOx matrix, while it does affect the adhesion of the film to the
underlying Au substrate. The addition of an optimal amount of Nafion® to the film aids in
enhancing IrOx interconnectivity and the robustness of the film on the Au substrate. It is also
shown that glucose, or the products of the reaction of glucose with GOx, do not cause fouling of
the IrOx matrix or affect the IrOx redox kinetics.
59
4.2 Characterization of IrOx nanoparticles synthesized from 100% ethanolic sols
Ir nanoparticle (NP) inks are typically synthesized in a 100% ethanolic solution. Once
deposited and dried in air at room temperature on a substrate such as Au, these thin Ir NP films are
exceptionally stable. In the ethanolic ink, chloride ions likely act as an electrostatic stabilizing
agent to ensure that the Ir NPs remain as a colloidal suspension. The addition of AgNO3 to an
aliquot of Ir ink immediately results in the precipitation of the Ir NPs out of solution, due to the
complexation of the chloride ions, forming AgCl(s). In the Ir ink, the ionic strength is relatively
low, as the chloride ions are not very soluble in ethanol206. When the ionic strength of the solution
is low, this ensures that the double layer thickness around the NPs is large, causing electrostatic
repulsion of the NPs and preventing the NPs from interacting with each other, thus preventing
aggregation207.
High resolution transmission electron microscopy (HR-TEM) was used to determine the
particle size distribution of the suspended Ir NPs, formed by the reduction of iridium trichloride
by sodium ethoxide in dry ethanol. Figure 4.1a shows that, after room temperature air-drying of
an aliquot deposited on a carbon/Cu TEM grid, the particles are primarily in the range of 1-2 nm
in diameter (Fig. 4.1b). The Ir NPs are seen to be uniformly dispersed, a characteristic required
for the formation of an effective, well-interconnected electron transfer matrix. The SEM image
(Fig. 4.1c) shows that after electrochemical oxidation, the IrOx NPs grow together to form large,
well-interconnected, IrOx globules ca. 1 µm in diameter. These globules form a well-
interconnected electronically conductive IrOx matrix. It has been shown in the literature that the
IrOx NPs are ca. 5 times larger than the Ir NPs, depending on if they were oxidized in neutral
phosphate buffer solution or H2SO4114.
60
Figure 4.1. (a) Transmission electron microscopy bight field image of ethanolic Ir ink, (b)
histogram of Ir particle size obtained from (a), and (c) scanning electron microscopy image of an
IrOx film obtained using a 20 kV accelerating voltage.
Consistent with Fig. 4.1c, Fig. 4.2a confirms that the Ir NPs can be easily electrochemically
converted to Ir oxide (IrOx) NPs when subjected to potential cycling in 0.5 M H2SO4, with the
upper limit successively increased up to 1.25 V vs. RHE, similar to what is done with bulk110,111
and electrodeposited208 Ir. The potential was then cycled 65 times (to ensure no further oxidation)
from 0 to 1.25 V to electrochemically oxidize the Ir to IrOx (Fig 4.2a). For scans up to only 1.0 V,
the Ir deposit is maintained in the metallic form. At more positive potentials, the hydrogen
1.2 1.60
3
6
9
Num
ber
of
Part
icle
s
Ir particle diameter / nm
Number of
Particles = 50
(b)(a)
(c)
61
underpotential deposition (Hupd) peaks are seen to diminish in size as the exposed metallic Ir
surface area decreases and the cyclic voltammetry features associated with IrOx209,210, particularly
the peaks centered at ca. 0.8 V, related to the Ir(III)/Ir(IV) oxide redox reaction (A1/C1,
respectively), begin to develop (Fig 4.2b). IrOx is the preferred form of Ir for glucose sensing
purposes, as it exhibits very rapid redox kinetics and is both physically and chemically stable. It
should be noted that the Ir NPs would slowly transform to IrOx under the constant potential
conditions (1.2 V) used for glucose oxidation, and thus it is logical to perform this conversion to
IrOx before the glucose detection experiment commences.
The A1/C1 peak currents in the IrOx cyclic voltammogram (CV) in Fig. 4.2a are proportional
to the potential sweep rate, even at relatively high sweep rates, as expected for the kinetically rapid
IrOx redox reaction. This is similar to what has been reported previously for IrOx materials formed
from 100% ethanolic Ir sols112,198. At potentials > 1.2 V, the IrOx film can be oxidized further to
Ir(V,VI) states, with their redox kinetics being even more rapid than the Ir(III)/(IV) redox
process112,198.
Figure 4.2. (a) Cyclic voltammetry (0.01 V s-1) showing the oxidation of an Ir ink (synthesized in
a 100% EtOH medium) in a stirred 0.5 M H2SO4 deaerated with Ar. (b) Decrease in Hupd peak
charge density with increasing number of cycles between 0 and 1.25 V, as in (a).
0.0 0.5 1.0
-0.2
0.0
0.2
A1
I /
mA
E / V vs. RHE
Hupd
C1
(a)
0 10 20 30 40 50 60
100
200
300
Hu
pd
ch
arg
e, C
Number of Cycles 0 - 1.25 V vs. RHE
(b)
62
When the Ir NPs have been completely electrochemically converted to IrOx, the charge in
the Hupd peaks decreased to zero. Typically, films must be cycled to potentials > 1.25 V, such as
1.45 V vs. RHE, to fully convert Ir to IrOx. To determine if the absence of Hupd peaks implies
complete Ir oxidation, X-ray photoelectron spectroscopy (XPS) analysis was performed. It was
assumed that the Ir-containing sample was homogenous with no preferred distribution of particles
(Ir or IrOx) at the surface of the sample. Figure 4.3a shows the Ir 4f7/2 and Ir 4f5/2 peaks for a fully
electrochemically oxidized IrOx thin film (ca. 15 µm) at 62.3 and 65.2 eV, respectively. These
values are close to the expected values of 62.7 and 65.7 eV211 for IrO2. With peak fitting, it was
found that there are no features that would indicate the presence of metallic Ir in the film (4f7/2 and
4f5/2 peak energies would be expected at ca. 61.1 and 64.1 eV)211. The third minor peak at the
higher binding energy side of the 4f5/2 peak obtained after deconvolution is assumed to be due to
the conduction band interaction during the photoemission process212. A similar peak was observed
by Atanasoska et al213., when characterizing RuO2/IrO2 mixed oxide layers.
The XPS O 1s spectrum (Fig. 4.3b) shows one main peak at 531.6 eV, with a second shoulder
peak at 533.2 eV. The O peak energy at 531.6 eV is close to the value seen for oxygen in powdered
Ir oxide, which is between 530.5 and 532.2 eV for Ir:O atomic ratios of 1:4 and 1:3,
respectively211,214. The shoulder peak at 533.2 eV is likely due to some retained ethanol from the
Ir sol215.
63
Figure 4.3. X-Ray Photoelectron Spectroscopy results, showing the (a) 4f7/2 and 4f5/2 and (b) O 1s
spectra, for an electrochemically oxidized thin IrOx film (ca. 15 µm).
X-Ray Diffraction (XRD) was used in an attempt to determine the crystallite size of the Ir
NPs in the ethanolic sol. XRD analysis was performed on a thin Ir NP film (ca. 300 nm thick)
which was loaded on amorphous glass. As seen in Figure 4.4, no peaks related to any of the Ir
crystal facets are seen, as Ir peaks should be observed at ca. 41 ° (111), 47 ° (200), 69 ° (220), 84
° (311) and 88 ° (222)216. It is well known that, if the crystallite size of the sample is too small (30-
70 68 66 64 62 60 580
500
1000
1500
4f5/2
, 65.2 eV
Inte
nsity /
CP
S
Binding Energy / eV
4f7/2
, 62.3 eV
(a)
958 957 956 955 954 953 952 951 950
800
1200
1600
2000
EtOH 633.2 eV
Inte
nsity /
CP
S
Binding Energy / eV
(b)
O 1s 631.6 eV
64
50 Å), the peaks will be very broad217. It is possible, then that the Ir crystallite size is too small (<
2 nm, according to TEM, Fig. 4.1 a and b) to be measurable, or that the sample is amorphous.
The peaks in the XRD spectrum correspond to the reactants and products of the Ir NP
synthesis reaction (Section 3.1.1). Sodium ethoxide is seen at ca. 10 ° (reactant) and sodium
chloride is detected at ca. 31 ° and 45 ° (product), while the large broad peak at ca. 22 ° is due to
the amorphous glass.
Figure 4.4 X-Ray diffraction pattern of Ir NPs from an ethanolic Ir sol loaded on amorphous
glass in the 2θ range of 0°-90°.
These results, taken together, show that the Ir NPs in the ethanolic sol are very small (1-2
nm in diameter) and, when electrochemically oxidized (by potential cycling to ca. 1.45 V), the
NPs are converted completely to IrOx (of varying oxidation states). If a film fabricated from an
ethanolic Ir sol is only potential cycled to 1.25 V, only partial Ir oxidation will occur, and some of
the film will remain in the metallic state. Although there is typically some unreacted Ir metal
present in any Ir-containing film after electrochemical oxidation, films will be referred to, for
0 10 20 30 40 50 60 70 80 90
0
200
400
600
800
Gla
ss
Na
Cl
CH
3C
OO
Na
Inte
nsity /
Co
un
ts
2 Theta / O
Na
Cl
65
simplicity, as IrOx films. The reason for this simplification is that the IrOx NPs are the dominant
phase, as they are large and more interconnected to each other due to the hydrous oxide187, whereas
the Ir NPs smaller and more isolated.
4.3 Effect of % water in Ir ink
The two major components in the glucose sensor being developed here are glucose oxidase
(GOx), for specificity, and IrOx NPs for accuracy, signal intensity, enzyme regeneration and
electron transfer to the underlying substrate. However, each of these essential components requires
different solvents for optimal functioning. The biological element, GOx, requires an aqueous
environment to ensure that the enzyme remains in its native, active protein configuration218. The
colloidal Ir NPs, in contrast, are synthesized in an ethanolic environment (where the ethanol acts
both as the solvent and reducing agent) to ensure small NP formation88,219. As such, it was
imperative that the effect of water in the Ir ink and its subsequent electrochemistry was
investigated.
4.3.1 Effect of water content of Ir inks on ink and film properties after drying
Water and ethanol are known to form an azeotrope when the ethanol content is quite high
(95.63 wt. % ethanol and 4.37wt. % water). At this specific mass ratio, the boiling point of the
mixture is lower than that of either ethanol or water. At any other composition, however, water
and ethanol form a zeotropic mixture (Fig. 4.5), and ethanol vaporizes at a lower boiling point than
water due to differing vapor pressures220. This faster rate of ethanol evaporation likely results in
uneven rates of film drying, resulting in aggregation and the appearance of “coffee stains”, arising
when regions of the film dry at different rates.
66
Figure 4.5. Average evaporation rate (g h-1) as a function of ethanol concentration (% w/w) for
both the ethanol and water components of a mixture (plot reproduced from data compiled by K.D.
O’Hare et al)221.
In this work, the water content of the Ir inks was varied from 10 to 50% by volume,
corresponding to a water content of 12 to 56 % by mass. These water contents were chosen because
the Ir ink must be fabricated in pure ethanol, and as such, diluting the concentration of Ir by greater
than 50% will result in too low a concentration of the Ir NPs for good glucose oxidase (GOx)-Ir
oxide (IrOx) interconnectivity when fabricating the sensing film. Lower water contents were not
considered, as a 10% ethanol content resulted in a substantial loss of enzyme activity in the sensing
film. The high water content of these inks means that, unfortunately, ethanol vaporizes from the
mixture at a faster rate than water (Fig. 4.5). When an aliquot of a water-containing ethanolic Ir
ink is deposited on a Au substrate, the resulting film thus should not dry uniformly. It is imperative
for this work, however, that the Ir (nanoparticles) NPs are as uniformly distributed as possible to
ensure that, when GOx is present, each enzyme molecule is uniformly surrounded by
interconnected IrOx NPs capable of direct electron transfer from GOx to the Au substrate.
0 20 40 60 80 1000
5
10
15
20
EtOH
H2O
Evapora
tion R
ate
/ g
hr-1
[Ethanol] / w/w%
67
As seen in Figure 4.6a (left), a typical freshly synthesized ethanolic Ir ink is dark brown in
color, due to the suspension of a high concentration of 1-3 nm Ir NPs. When the Ir ink is mixed
with 50% water (Fig. 4.6a, right), the NPs remain suspended, and the dark brown color persists.
When these two inks (100% ethanol, and 50% ethanol/ 50% H2O) are each cast on a sputtered Au
substrate and allowed to dry in air at room temperature for 2 hours, the resulting films, however,
have very different morphologies. The pure ethanolic ink exhibits good wettability, spreading
readily over the Au surface to ultimately form a smooth, uniform film (Fig. 4.6b, left). This ink
dries rapidly in air, due to the high volatility of ethanol. The 50 vol. % water-containing ethanolic
Ir ink (Fig. 4.6b, right), however, shows poor wettability of Au, initially forming a bead, or droplet,
on the surface. The outer edges of the droplet evaporate quickly, decreasing the size of the bead.
After a period of time, the droplet dries completely, resulting in a non-uniform film containing
drying rings (or “coffee stains”), and aggregated IrOx NPs.
Figure 4.6. Visual comparison between (a) an ethanolic Ir ink and a 50:50 vol. % H2O:EtOH Ir
ink, and (b) the resulting IrOx films after drying on a sputtered Au substrate.
68
It is possible that the water/ethanol droplet may undergo surface-induced separation on the
surface of Au, causing ethanol to move toward the periphery of the droplet, where it volatilizes.
Then, as the water droplet evaporates, this leaves behind rings of Ir NPs at the contact lines. The
aggregation of Ir NPs within the increasingly water-rich droplet may also be related to the changing
concentration of chloride ions present in the droplet. Chloride ions act as a stabilizing agent for
the Ir NPs, keeping them suspended in the ink. As the ethanol evaporates from the ink, however,
the concentration of chloride ions in the water droplet increases. This increase in ionic strength
would result in a thinner NP double layer, which destabilizes the NPs, causing the total surface
energy of the NPs to increase, and the NPs to aggregate (Fig. 4.6b).
4.3.2 Effect of water content Ir ink on subsequent IrOx film properties
The cyclic voltammetry (CVs) for films fabricated from a purely ethanolic Ir ink and a 50:50
vol. % water:ethanol Ir ink (Fig. 4.7a), followed by partial or full electrochemical oxidation of the
Ir nanoparticles (NPs) to Ir oxide (IrOx), reproducibly indicate that, despite their different
morphologies (Fig. 4.6b) water has only a small effect on the IrOx redox kinetics, after drying.
The addition of water to the ink, does, however, appear to enhance the interconnectivity of Ir
present in the film, as the charge under the Hupd peaks is ca. 3x larger (3.6 vs. 1.2 mC cm-2) for
the film fabricated from the water-containing vs. ethanolic ink. The anodic IrOx peak current
density (ipa) at 1.1 V is also larger for the film fabricated from the water-containing vs. ethanolic
ink (0.13 vs. 0.10 mA cm-2). Ir and IrOx NP interconnectivity can be determined by the charge in
the Hupd peak current, and in the Ir(III)/Ir(IV) oxide peak, respectively. The greater the
interconnectivity and connectivity to the electrode, the larger the charge will be observed.
In diffusion controlled systems, the slope of a log ip vs. log ν plot should be equal to 0.5, the
exponent of the sweep rate variable in the Randles-Sevcik equation (Equation 4.1)174. In a surface
69
controlled system, where there are no diffusional limitations, the slope should be equal to 1, the
exponent of the sweep rate variable in Equation 4.2174.
𝑖𝑝 = (2.69 𝑥 105)𝑛3
2𝐴𝐷0.5𝑣0.5𝐶 (Equation 4.1)
𝑖𝑝 = (9.39 𝑥 105)𝑛2𝐴𝑣𝐶 (Equation 4.2)
In Equations 4.1 and 4.2, n is the number of electrons transferred in the redox reaction, A is the
area of the electrode (cm2), F is the Faraday Constant, D is the diffusion coefficient (cm2 s-1), C is
the concentration (mol cm-3) and υ is the scan rate (V s-1).
The sweep rate analysis in Figure 4.7b shows that the slope of the log i vs. log ν plot is
equal to one for both the anodic and cathodic Ir(III)/Ir(IV) oxide peaks for IrOx films formed from
both the water-containing and pure ethanolic inks. A slope of one indicates that the electron
transfer process is surface controlled, as opposed to diffusion controlled, the result of facile
movement of ions (protons, Na+, or HPO4-) and electrons through the IrOx matrix fabricated from
either ink.
As another method to study electron transfer kinetics, Armstrong et al.222, and Jhas164
plotted peak current potentials (Ep) vs. the log of the scan rate (ν) using a Butler-Volmer model
(fitting the points to a typical Tafel slope of 0.120 V/ log ν) to obtain the “reversibility parameter”,
νo223, which is the sweep rate at which the electron transfer kinetics switch from reversible to
irreversible. At slow sweep rates, the peak potential should not shift dramatically, as the redox
reactions have a long time-scale to occur and a constant Ep demonstrates reversible electrochemical
behavior. If the sweep rate is increased to a point where the redox reaction cannot occur rapidly
enough for charge transfer to take place (ν > νo), this electrochemical behavior is considered
kinetically irreversible. Figure 4.7c shows, however, that, νo is smaller for the film fabricated from
a water-containing vs. ethanolic Ir NP ink. Films fabricated from inks containing 50% water show
70
Figure 4.7. (a) CVs of films (10 mV s-1) formed from a pure ethanolic Ir ink (■) and a 50:50 vol
% water:ethanol Ir ink (▲). Electrochemistry was performed in deaerated, stirred, 1.0 M PBS at
a sweep rate of 10 mV s-1. (b) log of peak current vs. log sweep rate and (c) Ep vs. log ν for the
results shown in (a).
irreversible IrOx electron transfer kinetics at > 45 mV s-1, while the kinetics of films fabricated
from ethanolic inks become irreversible at > 90 mV s-1, although this is not a significant difference.
To determine the effect of water in the ink and also during film drying on the nanostructure
of the Ir-based films, a 50% water-containing Ir ink was sonicated and a film was cast on a Cu grid
for high resolution transmission electron microscopy (HR-TEM) analysis. The HR-TEM images
of the 50 vol. % water-containing ink show that, near the outer edge of the film (Fig. 4.8a), the Ir
NPs are uniformly distributed and are all 1-3 nm in size. Near the center of the grid (in the “drying
rings224” where the Ir NPs tend to aggregate during drying), large (microns in size) non-uniform
0.0 0.2 0.4 0.6 0.8 1.0 1.2 1.4-0.5
-0.4
-0.3
-0.2
-0.1
0.0
0.1
0.2
0.3
C1
i /
mA
cm
-2
E / V vs. RHE
EtOH Ir sol
50 vol.% H2O Ir sol
(a)
A1
Hupd
-2.5 -2.0 -1.5 -1.0 -0.5 0.0-6
-4
-2
0
2
4
6
Slope: 1 log A / log
log
ip /
A
log / Vs-1
(b)
-2.5 -2.0 -1.5 -1.0 -0.5 0.0
0.75
0.80
0.85
0.90
0.95
1.00
Ir/H2O,
0 = 44 mVs
-1
EP v
s.
RH
E
log () / Vs-1
Ir, 0 = 89 mVs
-1
(c)
71
clusters of individual Ir NPs are seen at a higher magnification. Although clustered, these NPs are
1-3 nm in diameter (Fig. 4.8c), indicating that the addition of large amounts of water to the ink
does not affect the size of the NPs, but does cause Ir NP aggregation. It is seen that, surrounding
the NP aggregates, there are many individual NPs that are uniformly distributed and not clustered
together.
Figure 4.8. HR-TEM images of 50 vol. % H2O containing ethanolic Ir ink on Cu-TEM grid. (a)
Outer region of grid where EtOH evaporated quickly, (b) center of grid where Ir NPs aggregated
in the water-rich droplets, and (c) magnified view of (b).
Although the addition of 50 vol. % water to the ethanolic Ir sol enhances the
interconnectivity of the IrOx NPs (Fig. 4.8a), the film morphology is non-uniform (Fig. 4.6b and
4.8b and c). Nafion® was added to the ethanolic ink to determine how it would affect NP
interconnectivity in the absence of water. Nafion® was then added to the 50 vol. % water-
containing Ir inks, to determine if the high Ir NP interconnectivity observed with the high water
content (Fig. 4.8) could be retained, while stabilizing the Ir NPs, preventing aggregation.
72
4.4 Effect of Nafion® in Ir inks
4.4.1 Effect of 1 wt. % Nafion® in Ir ink on Ir film morphology
When an ethanolic Ir sol is deposited on a sputtered Au substrate, it forms a uniformly
dispersed Ir film after solvent evaporation (Fig. 4.9, i). When the Ir NPs are then partially
electrochemically converted to Ir oxide (IrOx), the resulting IrOx film is very stable and adheres
to the surface well, as is evidenced by the optical images shown in Figure 4.9i. In this work, we
show (Fig. 4.6b) that the addition of water to ethanolic Ir inks result in non-uniform Ir films with
large degrees of aggregation. The formation of these non-uniform films results in poor adhesion
of the Ir nanoparticles (NPs) to the Au substrate, and with rinsing, the Ir NPs aggregate, with some
then being lost to solution (Fig. 4.9, iii). With the addition of Nafion® to the ethanolic ink (Fig.
4.9, ii), the Ir NP dispersion is not as uniform as for the ethanolic-Ir film. When Nafion® is added
to the water-containing ink (Fig. 4.9 iv), drying rings are observed.
Figure 4.9. Optical images of IrOx films prior to the partial electrochemical oxidation of Ir and
after oxidation, followed by the rinsing of the films with water for 5 seconds. Films were fabricated
using 4 different Ir NP inks: ethanolic, ethanolic with Nafion®, a 1:1 ratio of water and ethanol,
and a 1:1 ratio of water and ethanol with added Nafion®. Each film contained 88 µmol Ir and
respective films contained 1 wt. % Nafion®, when Nafion® was present. Electrochemistry was
performed in deaerated, stirred pH 7, 0.1 M PBS.
73
4.4.2 Effect of Nafion® content in ink on subsequent IrOx electrochemistry
One of the challenges encountered in this work was the fabrication of films of uniform
morphology. As was shown in Fig 4.6b and 4.9i, a purely ethanolic Ir ink will form a uniform Ir
oxide (IrOx) nanoparticle (NP) film on a smooth Au substrate. With the addition of water to the
ink (a component required to retain high glucose oxidase (GOx) activity), however, the film dries
at an uneven rate (due to the different drying properties of ethanol and water), resulting in a less-
uniform or “coffee stained”224 (Fig. 4.6b, 4.9iii) film morphology. Ideally, for glucose sensing, the
Ir NPs should be uniformly dispersed throughout the Ir morphology, as this should produce the
best matrix for GOx/IrOx NP interactions.
It is known that the addition of a polymeric binder, such as Nafion®, can aid in the
stabilization of enzymes and metallic NPs in suspension204. Zhang et al. found that a small amount
of Nafion® was effective as a stabilizing agent in the synthesis of Ir colloids (ca. 1 nm dia.) for use
in fuel cells and electrolyzers. The addition of Nafion® to their sols resulted in very stable colloids
with excellent dispersion204. It is common knowledge that Nafion® is not electronically conductive
and thus an excess of the polymer could be detrimental to glucose sensing225. Too much Nafion®
could potentially hinder electron transfer by blocking the formation of a well-interconnected IrOx
NP network, obstructing GOx/IrOx NP interactions, and potentially blocking glucose access to
GOx active sites.
To determine the concentration of Nafion® that should be added to the ink to produce an
IrOx film that is stable and has rapid electron transfer kinetics, increasing concentrations of
Nafion® were added to ethanolic Ir inks (0, 1, 2, 4 and 8 wt. %), and the physical and
electrochemical properties of the resulting Ir and IrOx NP films were studied. Figure 4.10a shows
the CV for a film fabricated from an ink containing Ir NPs and Nafion® (no GOx). The size and
shape of both the cathodic and anodic Ir(III)/Ir(IV) oxide peak current densities (A1 and C1,
74
respectively) do not change dramatically when the amount of Nafion® in the films is increased
from 0 to 2 wt. %. The addition of more Nafion® (4-8 wt. %) to the ink, and hence in the resulting
films, however, is seen (Fig. 4.10a) to decrease the hydrogen underpotential deposition (Hupd)
peak size (0 to 0.4 V), as well as the cathodic and anodic Ir(III)/Ir(IV) oxide charges. The addition
of 8 wt. % Nafion® substantially diminishes the Hupd and IrOx peak currents, but does not affect
the overall IrOx character (Hupd peaks and Ir(III)/Ir(IV) oxide peaks are still observed). This
decrease in electroactive IrOx on the Au substrate is likely due to the electronic blocking effect of
Nafion®, causing some of the IrOx NPs to become electrically isolated, and the overall CV currents
to decrease.
Figure 4.10. (a) Cyclic voltammetry (100 mV s-1) for IrOx/Nafion® films fabricated from ethanolic
(no water) inks containing varying wt. % Nafion® (0%, 1%, 2%, 4%, 8%) at 100 mV s-1 in stirred
pH 7, 0.1 M phosphate buffer solution deaerated with Ar. (b) C1 peak current density (ipc) vs. log
sweep rate (ν) for films examined in (a).
The slope of the log peak current (ip) vs. log sweep rate (ν) plots for the IrOx/Nafion® films
(with increasing Nafion® content) in Figure 4.10b is linear and equal to 1 for low Nafion®-content
films (0, 1 and 2 wt. %). As the Nafion® content in the film is increased (4 to 8 wt. %), however,
the slope remains linear (and equal to one) at slow sweep rates, but begins to drop to below one at
faster sweep rates. IrOx/Nafion® films containing > 4 wt. % Nafion® exhibit a slope between the
0.0 0.5 1.0
-2.4
-1.2
0.0
1.2
Hupd
C1
i /
mA
cm
-2
E / V vs. RHE
0%
1%
2%
4%
8%
(a)
A1
-2.5 -2.0 -1.5 -1.0 -0.5 0.0-2.0
-1.5
-1.0
-0.5
0.0
0.5
1.0
0%
1%
2%
4%
8%
log ipc /
mA
cm
-2
log (Vs-1
)
(b)
75
accepted values of 0.5 (diffusion controlled process) and 1, i.e., ca. 0.85 for 4 and 8 wt. % films.
This suggests that portions of the film became diffusion controlled with a higher Nafion® content,
while the majority of the film remains surface reaction rate controlled.
It is possible for specific regions of the film to become diffusion controlled, as the
morphology is non-uniform (Fig. 4.9ii) and likely some areas of the film are highly aggregated
and dense. As such, higher concentrations of Nafion® (≥ 4 wt. %) are not recommended for
achieving good IrOx interconnectivity and kinetics. Direct mediation of electrons from the enzyme
active site to the current collector (underlying Au substrate) via the IrOx NP matrix (Section 2.4.4)
will also be affected by the presence of large quantities of Nafion®. Using this hypothesis, 1 wt. %
Nafion® was added to ethanolic and 50 vol. % water-containing Ir inks, and the IrOx redox kinetics
were compared for the two films to determine how Nafion® will affect the subsequent films, as
discussed below.
As the glucose sensing films are fabricated from inks that contain water, and as it was
shown in Fig. 4.7a that water in the ink also enhances the amount of active IrOx in the film, the
IrOx redox kinetics were compared for films fabricated from 50 vol.% water-containing sols vs.
50 vol.% ethanol-containing sols, with or without 1 wt. % Nafion®. It can be seen in Figure 4.11a
that both the charge in the double layer capacitance region of the IrOx CV (ca. 0.4 to 0.7 V vs.
RHE) and the IrOx anodic peak current (ip) (ca. 0.9 V vs. RHE) for films without Nafion® are
much larger (ca. 40%, and 34%, respectively, for films fabricated from water-containing and
ethanolic inks) than for films without Nafion®. The overall current enhancement indicates that the
addition of Nafion® to the ink results in an increased number of active IrOx NPs in the film, or
improved interconnectivity between the IrOx NPs. Interestingly, the charge in the hydrogen
underpotential.
76
Figure 4.11. (a) Cyclic voltammetry (100 mV s-1) for thin IrOx films fabricated from dilute
ethanolic Ir (1.1 mM) inks containing 1 wt. % Nafion®, 50 vol.% water, or a combination of both
to compare the effect of Nafion® on the IrOx redox kinetics in stirred, deaerated, 0.5 M H2SO4. (b)
log anodic i vs. log ν plot for the films in (a), and (c) Ep vs. log ν plot for the films in (a).
deposition (Hupd) peaks (Section 3.2.1.4) is largest for the film containing water and Nafion®,
indicating that Nafion® also enhances interconnectivity between Ir NPs
A plot of the peak anodic potential (Epa) vs. log ν (Fig. 4.11c) reveals that the Ir(III)/Ir(IV)
oxide redox reaction for the film fabricated from a pure ethanolic Ir ink (no water or Nafion®)
exhibits reversible electron transfer kinetics up to a sweep rate of ca. 17 mV s-1. With the addition
of 50 vol. % water to the Ir ink, νo for the resulting IrOx film was found to decrease marginally to
ca. 14 mV s-1. Interestingly, when Nafion® was added to either ink, the νo for the resulting IrOx
0.0 0.4 0.8 1.2
-2
0
2
Hupd
C1
I /
mA
cm
-2
E / V vs. RHE
Ir
Ir/Nafion
Ir/50 vol.% H2O
Ir/Nafion/50 vol.% H2O
(a)A
1
-2 -1 0
-4
-2
Ir/50 vol.% H2O
Ir/Nafion/50 vol.% H2O
Ir
Ir/Nafion
log i /
mA
cm
-2
log / Vs-1
(b)
-2 -1 0
0.84
0.96
1.08
Ir/50 vol.% H20
Ir/Nafion/ 50 vol.% H2O
Ir
Ir/Nafion
Ep / V
Log / Vs�-1
(c)
77
films decreased only to 12 and 9 mV s-1, respectively. These slight changes in νo indicate that the
presence of water and small amounts of Nafion® (or a combination of water and Nafion®) do not
significantly affect the reversibility of the IrOx redox kinetics.
4.5 Effect of glucose, H2O2 and gluconolactone in solution on the IrOx electrochemistry
To guarantee that the Ir oxide (IrOx) nanoparticle (NP) sensing matrix characterized in this
chapter is appropriate for specific and accurate glucose testing, it is important to perform “blank
experiments” to determine how glucose, or the products of the GOx/glucose reaction (H2O2 and
gluconolactone) react at the IrOx NP matrix. In particular, it is imperative that glucose and
gluconolactone do not react non-specifically at the IrOx NPs, whereas, in the case of first-
generation sensing, it is central that H2O2 undergoes rapid oxidation at the IrOx NP matrix.
4.5.1 Effect of glucose in the cell solution on IrOx redox kinetics
It has been shown in the literature that glucose will be oxidized at bare Au surfaces226 and
at colloidal Au NPs227. As such, it was important to determine if glucose would also react, non-
specifically, at the Ir oxide (IrOx) nanoparticle (NP) matrix. To observe the effect of glucose on
the IrOx NP redox kinetics, ca. 1.5 mM glucose additions were made to the cell solution and the
potential of the fully oxidized IrOx film was cycled from 0-1.25 V in a stirred, aerated
environment.
As can be seen in Figure 4.12, with increasing glucose concentration, the IrOx anodic peak
currents are not substantially affected. There appears to be a slight decrease in the peak Ir(IV)
oxide current during the positive potential scan, indicating that a small amount of glucose or the
product of glucose oxidation may be adsorbing to the film. The negative potential scan, however,
showed a significant loss of current (as compared to the positive potential scan) in the potential
78
Figure 4.12 Cyclic voltammetry (100 mV s-1) for a fully electrochemically oxidized (formed by
cycling from 0 to 1.45 V) IrOx electrode with increasing concentrations (0-12 mM) of glucose (1.6
mM additions) added to the cell solution in stirred, aerated 0.1 M, pH 7 PBS.
windows of ca. 1.0 to 0.8V and 0.5 to 0.2 V. These changes to the IrOx NP film are, however, negligible
in comparison to the high pseudocapacitive IrOx peak currents (Section 2.4.1).
4.5.2 Effect of H2O2 in the cell solution on IrOx redox kinetics
In addition to the very good electronic conductivity228 and biocompatibility116-118
characteristics required for a glucose biosensing matrix, Ir oxide (IrOx) has been reported to have
very good electrocatalytic action towards H2O2. As H2O2 is generated in a first-generation glucose
sensor (Reactions 2.1 and 2.2), where O2 is used to regenerate GOx, it was of interest to determine
whether the IrOx films would also be catalytic towards H2O2 oxidation/reduction. Figure 4.13 (a
and b) shows their response to various concentrations of H2O2 in neutral solutions under both
anaerobic (Fig. 4.13a) and aerobic (Fig. 4.13b) conditions.
Similar to IrOx films formed electrochemically on bulk Ir wires and foils228, Fig. 4.13 (a
and b) shows that H2O2 oxidation and reduction is quite facile on the sol-derived IrOx surfaces,
with the reactions commencing at ~0.9 V and ~0.65 V, respectively, and approaching current
0.0 0.4 0.8 1.2-4
-2
0
2
4
6
i /
mA
cm
-2
E / V vs. RHE
Increasing [glucose]
79
Figure 4.13. Cyclic voltammogram (0.1 V s-1) of an IrOx/Nafion® film (after electrochemical
conversion of Ir to IrOx), deposited using a 2 µL aliquot of 20% H2O/80% EtOH Ir sol + 0.04 %
Nafion®, deposited on Au in a deaerated, pH 7, 0.1 M phosphate buffer solution. (a) and (b) show
the CV response when aliquots of 3% H2O2 were added to the solution in (a) deaerated and (b)
aerated environments. H2O2 concentrations are 0 mM (solid line), 0.15 mM (dashed line), 0.3 mM
(dotted line) and 0.9 mM (dot dash line). The insets in (a) and (b) show the plots of the H2O2
oxidation (at 1.2 V) and reduction (at 0.2 V) currents vs. [H2O2] in each environment.
plateaus at 1.2 V and 0.2 V, respectively. Notably, the main CV features of IrOx are not altered,
showing that these IrOx films are very stable in H2O2 environments, an essential characteristic for
the sensing of glucose in the presence of O2.
In deaerated H2O2-containing solutions (Fig. 4.13a), the H2O2 oxidation currents at 1.2 V
are only slightly smaller in magnitude than the reduction currents at ca. 0.2 V (inset in Fig. 4.13b).
0.0 0.5 1.0
-0.4
0.0
0.4
0.8
0.0 4.0 8.0 12.0-0.4
-0.2
0.0
0.2
0.4
C1 (0.2 V)
i / m
A c
m-2
[H2O
2] / mM
A1 (1.2 V)
i /
mA
cm
-2
E / V vs. RHE
(a)
0.0 0.5 1.0
-0.8
-0.4
0.0
0.4
0.8
0.0 4.0 8.0 12.0
-0.2
-0.1
0.0
0.1
0.2
C1 (0.2 V)
i /
mA
cm
-2
[H2O
2] / mM
A1 (1.2 V)
i /
mA
cm
-2
E / V vs. RHE
(b)
80
However, when the solutions were saturated with O2 (Fig. 4.13b), O2 is reduced simultaneously
with H2O2 in the 0 to 0.4 V range of potential and the currents recorded at ca. 0.2 V are thus much
higher and more difficult to interpret than the ones detected without interference from O2, i.e., at
1.0 to 1.2 V (inset in Fig. 4.13b). The electrocatalysis of H2O2 oxidation and reduction by IrOx
takes place in a linear fashion (inset Fig. 4.13 a and b), and, as such, is a valuable material for the
fabrication of a first-generation (O2 and H2O2 dependent) biosensor112, but it is less desirable for
the fabrication of a third-generation biosensor. The sensitivity of IrOx to H2O2 can therefore be
useful in testing the effectiveness of IrOx as an electron transfer catalyst in the case of a dual O2-
and IrOx-mediated sensor response162.
4.5.3 Effect of gluconolactone on IrOx kinetics
It was determined that the Ir oxide nanoparticle (IrOx NP) matrix was a poor catalyst for
glucose oxidation, however, it was an excellent catalyst for H2O2. Consequently, it is important to
determine if the other product of the glucose reaction with GOx (Reaction 2.1), gluconolactone,
reacts at the IrOx matrix. To observe the effect of gluconolactone on the IrOx NP redox kinetics,
ca. 1.5 mM gluconolactone additions were made to the cell solution and the potential of the IrOx
NP film was cycled from 0-1.25 V in a stirred, aerated solution.
When the gluconolactone is dissolved in an aqueous environment, it is hydrolysed to
gluconic acid. There was no literature found regarding the oxidation of gluconic acid or
gluconolactone at a metal surface. As can be seen in Figure 4.14, with increasing gluconic acid
concentration, both the IrOx anodic and cathodic peak currents are slightly increased. These
changes to the IrOx NP film are, however, negligible in comparison to the high pseudocapacitive
IrOx peak currents (Section 2.4.1).
81
Figure 4.14. Cyclic voltammetry (100 mV s-1) for a fully electrochemically oxidized (formed by
cycling from 0 to 1.45 V) IrOx electrode with increasing concentrations (0-12 mM) of
gluconolactone (1.6 mM additions) added to the cell solution in stirred, aerated 0.1 M, pH 7 PBS.
4.6 Summary
The primary goal of this work was to understand the relationship between an ethanolic Ir
nanoparticle (NP) ink, water, and Nafion® when mixed together to form the electron transfer
matrix for the immobilization of GOx. It is shown in Chapter 4, that, the Ir NPs synthesized in
pure ethanol are very small, ca. 1-2 nm in diameter. Ir NP films formed from ethanolic inks have
uniform morphologies (uniform NP distribution), and the NPs can readily be electrochemically
oxidized to IrOx NPs. This conversion from Ir to IrOx can be followed by studying the change in
the charge in the Hupd and Ir(III)/Ir(IV) oxide peak current densities. The electron transfer kinetics
of these IrOx films is also shown to be rapid, exhibiting reversible redox kinetics at fast potential
sweep rates.
It is shown that the addition of water (50 vol. %) to the ethanolic Ir inks results uneven
drying of the ink, and aggregated IrOx films. This aggregation does not affect the film redox
kinetics, and even slightly enhances IrOx interconnectivity in the film. The NP aggregation is a
result of the different evaporation rates of water and ethanol, and possibly changes in the stabilizing
0.0 0.4 0.8 1.2
-4
-2
0
2
4
6
8
i /
mA
cm
-2
E / V vs. RHE
Increasing
gluconolactone
82
agent concentration in the ink, as it dries. This aggregation was found to result in IrOx films with
poor adhesion to the underlying Au substrate.
Nafion® was added to the inks as a binding agent to determine if it could aid in the
stabilization of the Ir NPs in the ink (ethanolic, and water-containing), as well as IrOx film
stability. Too much Nafion® (> 2 wt. %) in the ink severely affected the IrOx interconnectivity in
the resulting films, causing some regions of the film to become diffusion controlled. The optimal
amount of Nafion® (1 wt. %), however, was found to increase IrOx interconnectivity in the film,
as well as aid in adhering films fabricated from water-containing inks on the Au substrate. The
effect of glucose, hydrogen peroxide and gluconolactone (products of the glucose oxidase
(GOx)/glucose reaction, Section 2.2.1) in solution, on the IrOx redox kinetics were also studied,
and it was found that they do not affect the IrOx redox kinetics. It is also shown that the oxidation
signal of glucose and gluconolactone directly at the IrOx film are negligible.
With these new insights, it is important to determine how the addition of the second major
glucose sensing component, GOx, interacts with these water and Nafion®-containing inks. Chapter
5 will introduce how a sensing film (with optimized water and Nafion® content) responds to
glucose in both aerated (O2-mediated) and deaerated environments (direct electron transfer via the
IrOx NPs). The methods for determining the optimal film component ratios will also be discussed.
The ratio of water:ethanol, in the ink will be shown to be important for GOx activity and therefore
glucose signal, while the amount of Nafion® in the inks appears to affect the IrOx NP redox
kinetics, which directly affects the glucose signal. Finally, the effect of GOx on the IrOx NPs in
the ink and in the sensing film will be studied to understand how direct electron transfer may take
place between GOx and the IrOx NPs.
83
CHAPTER FIVE: EFFECT OF SENSOR PREPARATION ON DUAL-
MEDIATION SENSING OF GLUCOSE*
*Section 5.2.1 was published in Campbell, H.B.; Elzanowska, H.; Birss, V. I. Biosensors and
Bioelectronics 2013, 42, 7.
*Portions of Sections 5.2.2 and 5.5 were published in Jhas, A.; Elzanowska, H.; Sebastian, B.;
Birss, V. Electrochimica Acta 2010, 55, 7.
5.1 Introduction
In Chapter 4, the Ir nanoparticles (1-2 nm) that are used as the electron transfer and
immobilization matrix for glucose oxidase (GOx) were examined. It was found that an Ir film
fabricated from an ethanolic ink has a very uniform distribution of Ir nanoparticles (NPs), which
can readily be converted to Ir oxide (IrOx) by electrochemical oxidation. When water was added
to the Ir ink (a component required for high GOx activity), however, the resulting IrOx NPs
aggregated, resulting in poor adhesion to the underlying Au substrate. As such, Nafion® was added
to the inks (ethanolic and water-containing), and 1 wt. % was found to enhance IrOx NP
interconnectivity (as gauged by the charge in the Ir(III)/Ir(IV) oxide peaks in the IrOx cyclic
voltammetry) and adhesion of the NPs to the Au.
With this knowledge of how to fabricate a stable Ir NP matrix that exhibits rapid electron
transfer kinetics (from a water/ethanol sensing ink), it was important to determine how GOx would
be incorporated into the matrix. To enhance the glucose signal obtained from the sensing film, the
effect of the ink components on the film morphology and interconnectivity of the IrOx NPs matrix
must be considered. In fact, the connectivity between GOx and the IrOx NPs are shown to
determine the mechanism by which GOx will be regenerated after reacting with glucose. If the
IrOx NPs are in good contact with GOx, electrons can be transferred directly from the GOx active
site, through the interconnected IrOx NPs, and to the underlying Au substrate (Fourth-generation
sensor, Section 2.3). If GOx is not in good electrical contact with the IrOx NP matrix, electrons
84
will be transferred via the O2-mediation route (First-generation sensor, Section 2.3). In this work,
it has been shown that the IrOx-based glucose biosensors are capable of GOx regeneration via
dual-mediation (via both the O2-and IrOx NP mediated routes (Scheme 5.1)) 10.
This chapter is divided into five sections, with the first section introducing a typical dual-
mediated glucose response of the IrOx/Nafion®/GOx glucose sensing film (Scheme 5.1). An
argument is then made attesting to the claims that the response seen under deaerated conditions is
a result of GOx being regenerated directly by the IrOx NP matrix, as opposed to by trace O2 present
in the solution.
Scheme 5.1 Dual mediation occurring in an IrOx/Nafion®/GOx film. Direct mediation occurs
when GOx is in good electrical contact with the interconnected IrOx NP matrix, while O2-
mediation takes place when GOx is not well-connected to the IrOx NP matrix. The triangle
represents glucose. Note that Nafion® is not represented in this scheme.
The next four sections involve an investigation into the effects of GOx and the other film
components (water and ethanol, the IrOx NPs and Nafion®), on the IrOx redox kinetics, film
morphology, and the regeneration of GOx via dual-regeneration. GOx has been shown in the
85
literature to adsorb to metal NPs229-233, and, as such, the effect of GOx on the sensing film
morphology and IrOx redox kinetics were studied using a range of microscopic and spectroscopic
characterization techniques. It was found that GOx interacts with the Ir NPs, causing them to
aggregate around the GOx molecules. This aggregation prevents the conversion of Ir to IrOx and
slows down the IrOx electron transfer kinetics. The effect of the water:ethanol ratio in the sensing
ink on GOx activity in the ink, the nature of Ir/GOx film drying, and on glucose sensing were also
studied. The effect of GOx on the IrOx electrochemistry and glucose sensing were also studied,
and the optimal ratio of GOx:Ir in the sensing ink for glucose sensing was determined. The last
section is focused on the effect of increasing amounts of Nafion® on glucose sensing.
5.2 Dual-mediation glucose response of IrOx/Nafion®/GOx films
5.2.1 Typical glucose signal from an IrOx/Nafion®/GOx sensing film in aerated and deaerated
solutions
The Michaelis-Menten (MM) plot (Fig. 5.1) shows the effect of added glucose to the current,
recorded with time at 1.2 V for an Ir oxide (IrOx)/Nafion®/glucose oxidase (GOx) film in a pH 7
buffer solution, in an aerated and deaerated solution (saturated with Ar). The background current
is very small, consistent with the capacitive properties of IrOx films, where the oxidation state of
Ir within the oxide film rapidly equilibrates and the current quickly drops to zero at constant
potential. Also, the rapid increase (within 2 s, shown in Fig. 6.1) in the chronoamperometric
currents recorded after each glucose addition indicates the fast response of our sensor.
The application of relatively high positive potentials (1.2 V vs. RHE) does not change the
properties of the IrOx and IrOx/Nafion®/GOx films. In fact, the cyclic voltammetry (CV) response
(Fig. 4.2a) of the sensor remains essentially the same before and after potentiostatic measurements
involving glucose additions. Due to the high pseudocapacitative IrOx currents (Section 2.4.1), the
86
Figure 5.1. Michaelis-Menten plot (at 1.2 V) in stirred, pH 7 0.1 M, RT, phosphate buffer solution
(25 mL) at room temperature with 20 µL increment additions of 1.0 M glucose under aerated (*)
and deaerated (■) conditions for a film formed from a IrOx/Nafion®/GOx ink.
currents related to glucose oxidation at IrOx are relatively small in CV experiments (Fig. 4.12), but
under potentiostatic conditions, the oxide is fully charged and thus only the signal for the catalytic
oxidation of glucose via GOx is seen (Fig. 5.1).
When the glucose addition experiments were performed in deaerated solutions, an excellent
response to glucose is still seen (Fig. 5.1), demonstrating that GOx re-oxidation can also occur by
direct electron transfer to the Ir oxide NPs within the film matrix or via the O2-mediation route
(dual-regeneration, Scheme 5.1). In the deaerated solution, the K’m value (Section 2.5) for this
particular sensor is 2.1 mM, i.e., less than the free enzyme K’m value of 25 mM, and imax (Section
2.5) is 15 µA cm-2, while in aerated media, the K’m and imax values are both larger (7.3 mM and 50
µA cm-2, respectively).
Although the dissolved O2 concentration should be very low in the Ar-saturated cell solution,
the maximum glucose oxidation current observed in Figure 5.1 at saturation (∼25 mM glucose) is
∼70 % less than in air. In fact, negligible currents would have been expected in the absence of
O2234 if the regeneration of GOx were occurring only by O2 (a co-substrate), as observed for other
0 4 8 120
10
20
30
40
50O2 i max
i / A
cm
-2
[Glucose] / mM
70% loss in imax
87
O2-dependent enzymes, e.g., laccase235, which reduces oxygen to water (oxygen is the substrate
for the enzymatic reaction). In the case of laccase235, when the solution changes from O2-saturated
to air-saturated, a significant decrease in the biosensor signal is observed, and the signal is almost
zero (90–95 % loss of signal) when the solution is fully deaerated.
To verify that electron transfer is indeed occurring between the reduced flavin site in GOx
and the surrounding IrOx matrix under deaerated conditions, the amount of oxygen present was
carefully determined. The oxygen reduction current, monitored using a second Pt working
electrode in an O2-saturated solution, was found to be 2.1 mA cm-2 at 0.2 V. After 40 min of
deaeration with Ar, the current at 0.2 V is only 1.3 µA cm-2, i.e., the oxygen content has been
lowered by at least 1000 times10. This would result in a similar decrease in the observed glucose
oxidation currents if the only means of GOx regeneration were through reaction with O2, as seen
for laccase235. As the glucose oxidation currents in deaerated solutions (Fig. 5.1b, < 0.3 µM O2)
are much too high to be explained by trace O2 regeneration of GOx, these results argue strongly
that the IrOx sites are indeed mediating the GOx regeneration process. This is further supported
by the absence of any H2O2 (the product of O2 regeneration of GOx, Scheme 5.1) monitored
electrochemically in the deaerated medium10.
5.2.2 Effect of potential used for glucose sensing on glucose signal
Water is typically oxidized to O2 at 1.23 V vs. RHE234 at Ir, while glucose sensing is typically
performed at 1.2 V. The question could arise as to whether, if in a glucose detection experiment at
1.2 V, O2 is being evolved catalytically on the IrOx electrode, then serving to regenerate the
reduced form of GOx. This supposition is refuted by the fact that the glucose signal is still very
strong at 1.1 V, and a good response was still obtained even at 1.05 V (Fig. 5.2). It is impossible
88
Figure 5.2 Four IrOx/Nafion®/GOx films tested for their response to glucose at a constant potential
of either 1.05, 1.10, 1.15 or 1.20 V up to 70 mM of glucose in stirred, deaerated pH 7 0.1 M PBS.
that the insignificant amounts of O2 that would be generated at these very low potentials could be
sufficient to produce the comparatively high currents (at high glucose concentrations) seen in
Figure 5.1. It can be concluded, therefore, that the IrOx NPs are actively participating in the
electronic communication between GOx and the underlying support (Au), as shown in Scheme
5.1.
This direct electron transfer can be realized through the protein part of the enzyme, allowing
for electron transfer from the enzyme redox site (the flavin) to the IrOx NPs (at which the
Ir(IV)/Ir(V)/Ir(VI) oxide redox transitions occur) surrounding the enzyme molecule. The electrons
can then be transferred through the conductive IrOx layer to the support. This type of mechanism
has been proposed to explain the electrical communication in a well-known biosensor system in
which the enzyme is ‘wired’ by ferrocene/ferrocinium centers attached covalently to the
enzyme236. Similarly, the response in deaerated solutions has been documented in biosensing
layers in which the electron accepting/transferring relays are composed of a dense array of Os
complexes covalently bound to a long flexible polypyridine polymer237 and also, more recently, in
a system based totally on an organic polymer possessing PANI redox centers238.
89
The response at 1.2 V (Fig. 5.2) was found to produce the highest glucose signal, due to the
presence of the greatest number of Ir(IV)Ox sites available to accept electrons from the reduced
FADH2 active site. This potential is considered to be high for glucose sensing, as many interfering
electroactive species can react at the electrode at this potential239. However, it has been shown that
interfering species can be blocked (repelled by the negative charges on Nafion®)165, and as such,
to achieve the highest glucose signal, glucose testing was performed at 1.2 V in the remainder of
this work.
Dual-mediation10,164 is highly dependent on the interactions between the GOx molecules and
IrOx NPs, as well as the IrOx NP interconnectivity (to be discussed in the following section). It
will be shown in this work, that dual-mediation is affected when these interactions are altered, i.e.,
one of the electron transport pathways will be favored over the other. The factors that affect dual-
mediation include the water:ethanol ratio of the ink (Section 5.4), as well as the GOx (Section 5.5)
and Nafion® (Section 5.6) contents of the ink.
5.3 Physical and Electrochemical Characterization of Ir/GOx films
The two fundamental components in our glucose sensing electrodes are the Ir oxide (IrOx)
nanoparticles (NPs) and glucose oxidase (GOx). High resolution transmission electron microscopy
(HR-TEM) was used, to obtain a general idea as to how the Ir NPs dry as a film when they are
mixed together with GOx in the ink. HR-TEM was carried out on a thin film fabricated from an
ink containing GOx and 20% water. It was found that large amorphous structures were present in
the film (> 10 µm in diameter), assumed to be aggregates of GOx, with one such aggregate (of
average size) shown in Fig. 5.3 (a-c). At higher magnification, it is observed that these aggregates
are uniformly covered by Ir NPs, roughly 1-2 nm in diameter (Fig. 5.3d).
90
In the local areas surrounding the aggregates, the concentration of the Ir NPs is much lower
than in regions further away (Fig. 5.3e). Importantly, the Ir NP distribution and size are comparable
to those observed for the Ir sols without GOx (Fig 4.1a). GOx does not, therefore, physically alter
the Ir NPs, but does cause them to accumulate in its vicinity. This indicates that the metallic NPs
and the GOx molecules do interact with each other.
Figure 5.3. High resolution transmission electron spectroscopy images of a typical Ir NP/GOx
film that has been diluted with a 20 vol.% water/ethanol mixture by 50x. (a-c) shows magnified
regions of the film, (d) is a magnified view of the Ir NPs aggregated around the GOx molecules,
and (e) shows the Ir NP distribution at a location in the film that is far from the aggregated GOx
molecules.
91
Periera et al.233, showed that GOx can adsorb to metal NPs (Pd), thus partially blocking the
NP surface. It was seen from their TEM images that the Pd NPs aggregated around GOx, as is also
seen in this work (Fig. 5.3). Ren et al.232, reported that, when GOx interacts with a metal NP, the
NP replaces the aqueous environment at the point of interaction. This causes the GOx active site,
(flavin-adenine dinucleotide (FAD)) to be exposed to the aqueous environment, which acts to
enhance GOx activity. In contrast, Tellechea et al.230, found that GOx activity is negatively
impacted by conjugation to Au NPs, likely due to unfolding and exposure of internal amino acids
(such as cysteines) to the NP. GOx is known to readily interact with metal NPs and the effect of
these interactions on GOx activity appears to be dependent on the environment and the metal.
Sethi and Knecht stated that the level of understanding as to how biomolecules bind and
arrange on the nanomaterial surface is incomplete229, as little is known about peptide/NP surface
interfaces. The interactions between specific amino acids, such as arginine (Arg), lysine (Lys), and
aspartic acid (Asp), to Au and Pd surfaces were studied by Hong et al.231, where they found that
Arg had a weak interaction with the Au(111) facets via interactions with the guanidinium group
of the side chain. As there is still work to be done before the interactions between enzymes and
other biological molecules with NPs are understood, it remains unclear how GOx interacts with
the Ir NPs. However, this likely occurs through the polar amino acids on the exterior of the enzyme.
Under deaerated conditions, the cyclic voltammetry (CV) in Figure 5.4 for an IrOx/GOx film
shows that the peaks due to the Hupd reaction at the Ir NPs (Fig. 3.1a) disappear, and the
Ir(III)/Ir(IV) oxide region (A1/C1) does not have the characteristic IrOx shape (Fig. 4.2). The inset
of Figure 5.4 shows that, with the addition of GOx to an Ir ink, the resulting film has poor electron
transfer kinetics, becoming diffusion controlled at ca. 20 mV s-1. In comparison, an IrOx film
formed from an ethanolic Ir sol has substantially faster redox kinetics, maintaining surface
92
controlled electron transport kinetics upwards of 500 mV s-1 (Fig 4.7b). The poor IrOx redox
kinetics exhibited by the Ir/GOx film is likely due to the aggregation of the Ir NPs seen in the TEM
Figure 5.4. Typical cyclic voltammetry (10 mV s-1) of an IrOx/GOx (12 g l-1) glucose sensing film
(20 vol. % H2O) that was electrochemically oxidized by cycling the potential from 0 to 1.25 V in
stirred, deaerated 0.1 M pH 7 PBS. Inset shows the log ipa vs. log ν plot for an IrOx NP film and
an IrOx NP/GOx film.
images (Fig 5.3). It is, therefore, the goal of this section to determine the effect of GOx on the Ir
NP dispersion within the sensing film.
The CV of an electrochemically oxidized IrOx/GOx film (Fig. 5.4) suggests complete
conversion of Ir to IrOx occurred, as the current due to the Hupd reaction decreased to zero. The
A1 and C1 peaks at ca. 0.8 and 0.6 V, respectively, confirm the presence of Ir(III)/Ir(IV) oxide. In
comparison, electrochemical cycling from 0 to 1.25 V is insufficient to entirely convert a GOx-
free Ir film to IrOx (Fig. 4.2a). As such, the question was posed as to whether Ir is fully converted
to IrOx in the GOx-containing film, or if GOx aggregates the Ir NPs (as observed from the TEM
images, Fig. 5.3) and prevents access of the Ir NPs to the solution.
0.0 0.4 0.8 1.2
-20
0
20
40
60
Hupd C
1
-2 -1 0
-1
0
1
IrOx/GOx,
slope: 0.5
IrOx/GOx,
slope: 1
log i
p / m
A c
m-2
log / Vs-1
IrOx, slope: 1
I /
E / V vs. RHE
A1
93
To determine if any metallic Ir remains in the IrOx/Nafion®/GOx films after electrochemical
oxidation to form IrOx, X-ray photoelectron spectroscopy (XPS) analysis was performed. Figure
5.5 shows the Ir 4f7/2 and Ir 4f5/2 peaks for the film examined in Figure 5.4. It is evident from the
spectra that there is more than one form of Ir present in the film, as the 4f7/2 peak energy typically
has a higher intensity than the 4f5/2 peak energies. As such, the peaks were deconvoluted to identify
the different forms of Ir. The peak energies at 62.9 and 65.9 eV (4f7/2 and e4f5/2, respectively) are
close to the expected values of 62.7 and 65.7 eV211 for IrO2. From the cyclic voltammetry (CV) in
Figure 5.4, it was expected that IrO2 would be the predominant form of Ir in the film. However,
the second set of peaks at 61.9 and 64.8 eV indicate the presence of Ir with its 4f7/2 and e4f5/2 peak
energies of 61.1 and 64.1 eV211. As XPS was able to detect metallic Ir in the film, while the
electrochemistry (Fig. 5.4) did not, this suggests that the Hupd reaction is blocked from taking
place at the Ir NP sites, likely due to interactions between the Ir NPs and GOx.
Figure 5.5. XPS data for an IrOx/GOx (12 g l-1) sensing film that was electrochemically oxidized
by cycling the potential from 0 to 1.25 V in stirred, deaerated 0.1 M pH 7 PBS.
75 70 65 60 55
500
1000
1500
2000
2500
IrOx, 62.9
Ir, 61.9Ir, 64.8
Inte
nsity / C
PS
Binding Energy / eV
IrOx, 65.9
94
5.4 Effect of % water in GOx-containing inks
5.4.1 Effect of % water content on GOx activity
It was shown in Figure 4.7 that the water content of the primarily ethanolic inks does not
have a substantial effect on the Ir oxide (IrOx) nanoparticle (NP) electrochemistry when only Ir,
then converted to IrOx, is present. It is vital, however, to understand how the ethanolic
environment of the Ir ink could affect (glucose oxidase) GOx viability (conformation, activity and
dispersion) in both the ink and ultimately in the final immobilized state in the sensing film. It has
been documented in the literature240,241 that enzyme activity (in general) is negatively correlated
with the concentration of organic solvent in a water/organic mixture, giving three distinct activity
levels as the organic content is increased. At low organic solvent concentrations, the enzyme
activity is not significantly affected, as the enzyme only interacts with the few layers of water that
surround it. However, once a critical organic solvent concentration is reached, the enzyme
becomes denatured, as the organic compounds strip the water molecules from the hydrophilic
enzyme surface. As an example, hen egg lyzosyme was found to be denatured when dissolved in
60% acetonitrile242. Interestingly, if the organic content is increased to as high as 90 vol.%, GOx,
in particular, becomes insoluble (in most organic solvents)243 and cannot undergo any
conformational changes, leaving the enzyme stable, but with a low activity244.
Iwuoha et al.245, found that, when the activity of GOx was tested in pure butan-2-ol, the
apparent catalytic efficiency of GOx decreased significantly compared to that in an aqueous phase,
indicating that butan-2-ol strips water from the enzyme active site. Other groups246 have dissolved
GOx into organic solvents containing only 5-15 vol. % water and then tested the subsequently
fabricated films in aqueous solutions. It was shown that the enzyme activity levels were
unexpectedly high after the earlier exposure to the nearly pure organic solvent. At high
temperatures, GOx was found to be stabilized by the addition of 10-30% organic solvents
95
(methylol, ethyleneglycol, or glycerol)247. To our knowledge, however, there is little literature
regarding the catalytic activity of GOx (in solution) that has been exposed to high ethanol
concentrations, or how well the activity can recover when the water content of the ink is
subsequently returned to 100 %, the typical conditions under which the GOx response (in the
sensing film) to glucose is tested.
As a novel and facile method to compare the relative activity levels of GOx in ethanolic inks
of varying water contents, the rate of glucose consumption by GOx was monitored here using
electrochemistry. GOx was dissolved in stirred ethanol/PBS solutions (0.1 M KCl electrolyte)
ranging from 20-100 vol. % H2O (or 80-0 vol. % ethanol), with a constant rate of O2 influx. The
concentration of H2O2, the by-product of flavin adenine dinucleotide (FAD) regeneration by O2,
was then tracked during its oxidation at an IrOx electrode (at a potential of 1.1 V vs. RHE), giving
a current proportional to the concentration of glucose.
The current was monitored first in the absence of glucose and then again after the addition
of a 150 mM glucose aliquot into the ethanol/ PBS mixture (as depicted in Figure 5.6a). The charge
generated from the reaction of GOx and glucose over 50 s (the oxidation of H2O2 at the IrOx
electrode) was then plotted against the ethanol content in solution (Fig. 5.6b) with charges
corrected to account for the increase in O2 solubility as the ethanol content was increased248. The
correction was carried out by multiplying the glucose signal by the ratio of the Oswald distribution
coefficients for O2 solubility in ethanol at 25 °C, vs. the solubility of O2 in an ethanol-free solution,
as determined by Shchukarev and Tolmacheva. The Oswald distribution coefficient is defined as
the gas volume dissolved in an exact volume of solvent248.
96
Figure 5.6. (a) i/t data showing the baseline current ( ̶ ̶̶ ) (with no glucose present) and the current
produced when 150 mM glucose is present (---) in aerated, stirred PBS (with no EtOH present)
containing 1 g l-1 GOx. The increase in current is a due to the oxidation of H2O2 which is generated
when GOx reacts with glucose, and is regenerated by O2. (b) Charge passed under the same
conditions as (a) except the PBS solution was comprised of varying H2O:EtOH ratios. (c) Glucose
signal achieved after enzyme activity was recovered by decreasing the EtOH content of the GOx
solution from 80 vol. % to 20 vol. % (̶ ̶̶). Reference glucose signals achieved when GOx is dissolved
in a 20 vol. % EtOH (**) and an 80 vol. % EtOH (--) solution.
A similar approach for rapid enzyme activity detection (for FAD or ((nicotinamide adenine
dinucleotide (NAD))-containing enzymes such as GOx, alcohol dehydrogenase, lactate
dehydrogenase and xanthine oxidase) was recently published by Zhang et al.249, where they first
prepared a pre-assay calibration by injecting an exact amount of H2O2 into a glucose-containing
10 20 30 40 50 600
10
20
30
GOx
I / A
t / s
(a)
GOx + Glucose
0 20 40 60 800
2
4
6
8
10
Q / m
C
EtOH content / %
(b)
70 80 90 100 1100.0
0.1
0.2
80% EtOH Reference
20% EtOH Reference
Recovered in 20% EtOH
Cu
rre
nt
/ m
A
Time / s
(c)
97
solution of known volume and then recorded the oxidation current. They then injected GOx into
the solution and recorded the resulting H2O2 oxidation currents, generated by the reaction between
GOx and the added glucose. These results249 were not corrected for the changes in O2 solubility as
a function of ethanol content, however.
As predicted from the literature, Figure 5.6b shows that small amounts of EtOH (up to 20
vol. %) do not negatively impact the GOx enzyme activity (same as in 100% H2O), while with the
further addition of EtOH (> 20 vol. %), the glucose response (H2O2 oxidation signal) drops
significantly. It is likely that, at these concentrations, the “organic threshold” was breached and
the polar solvent began to strip water away from both the exterior and the active site of the protein,
thereby changing the conformation of GOx and affecting the functionality of the active site.
It should be noted in Fig 5.6b, that, as the amount of EtOH was increased to ≥ 60 vol. %, the
degree of GOx aggregation increases (formation of white flakes were seen in the solution) and the
GOx activity level then became very close to zero. This may be indicative of the third stage of
enzyme activity, where the enzyme is unable to undergo any further conformational changes and
the activity levels are greatly diminished. The charge passed during H2O2 oxidation in the 20 vol.
% EtOH solution is seen to be approximately 8x higher than that in the 50 vol. % EtOH solution
(Fig. 5.6b), clearly demonstrating that the more water there is in the ink, the greater the GOx
activity.
Interestingly, it was found that the GOx activity was recoverable if water was then
reintroduced into a highly ethanolic environment. This finding is important, as although GOx is
initially surrounded by a highly ethanolic environment, the sensing film is tested for its response
to glucose in a 100% aqueous environment (PBS). To verify that the enzyme activity could be
restored by “re-hydrating” the enzyme, GOx was dissolved in 80 vol. % EtOH for 30 min (Fig
98
5.6c). The enzyme formed white flakes, indicating denaturation and aggregation. The water
content of the enzyme solution was then increased to a total of 80 vol. % H2O, and was stirred for
45 minutes. Glucose was then injected into the solution and the enzyme response (via H2O2
oxidation) was monitored.
As seen in Figure 5.6c, the enzyme was able to recover approximately 30% of its initial
activity when the ethanol content was decreased from 80 to 20 vol. %. The charged passed from
the oxidation of H2O2 over 40 seconds in 20% and 80% ethanol was 5.0 mC and 0.8 mC,
respectively. Comparatively, the charge obtained after GOx “rehydration” was 2.0 mC. This 30%
increase in activity is significant, as it shows that GOx activity can recover, at least partially, after
being exposed to high ethanol contents. As such, high ethanol contents (80%) in the Ir ink can be
tolerated, as the enzyme activity can partially recover when the film is subsequently immersed in
PBS for glucose sensing. Even so, higher H2O-contents in the inks will lead to higher enzyme
activity. Although high water contents are beneficial for GOx activity, it is important to determine
if these high water contents affect the morphology of the final IrOx/GOx sensing film.
5.4.2 Effect of % water on IrOx and GOx–containing films
Glucose oxidase (GOx) was shown to require a high water content environment to maintain
high enzyme activity, especially in the ethanolic environment of an Ir ink (Section 5.4.1). It is
shown in Figure 5.6b that, as the amount of water in the solution increases, so does the glucose
signal. Unfortunately, in Section 4.3.2, it was shown that, although water in the ethanolic ink does
not have a great effect on the subsequent Ir oxide (IrOx) redox kinetics, it does affect the sensing
film morphology, resulting in an unevenly dispersed Ir nanoparticle (NP) film (Fig. 4.6b). As a
key goal of this work is to achieve uniform film morphologies, where GOx and the Ir oxide (IrOx)
99
NPs are evenly dispersed in an electrically conductive matrix, here we examine the effect of
increasing water content in a GOx-containing Ir ink on the subsequent film morphology.
As can be seen in Figure 5.7 (before rinsing), non-uniform morphologies and drying rings
(Fig. 4.9) are observed for Ir/GOx films fabricated from inks containing ≥ 10 vol. % water. After
electrochemical oxidation of Ir to IrOx, these non-uniform films fabricated from high water
content inks (> 50 vol. %) are not stable and exhibit adhesion issues. The IrOx aggregates have
been found to be easily dislodged from the Au surface (Fig 5.7, after rinsing) by rinsing with water
or during electrochemical experiments. Interestingly, it appears that, for many of these films, the
IrOx NPs are mobile, and thus poorly adhered at the electrode surface (this is very evident for
films fabricated from 50 to 70 vol. % water). A faint outline of the original film can be seen on the
Au substrate for the films after rinsing. The effect of GOx on the sensing film morphology, and
the agglomeration of the IrOx NPs on glucose sensing are of great interest and will be discussed
in the next section of this chapter. The instability that was observed in the films with rinsing is
likely due to the absence of a binder in the film. Similar to what was seen in Section 4.4.1, this
instability was countered by adding Nafion® to the Ir/GOx ink, and the effects of Nafion® on the
films will be discussed in the last section of this chapter.
Figure 5.7. Ir/GOx (before rinsing) and IrOx/GOx (after rinsing) films fabricated from increasing
water-content inks. Films are shown prior to electrochemical oxidation, and after the films were
electrochemically oxidized (converting Ir NPs to IrOx NPs) and rinsed with H2O.
100
5.4.3 Effect of % water in Ir oxide/GOx ink on glucose detection
One of the most difficult tasks when optimizing the response of a biosensor is determining
how all of the sensing components affect each other. In this work, the sensing ink is composed of
two different solvents, ethanol (Ir ink) and water (glucose oxidase (GOx) enzyme). Although these
two solvents are miscible, they have different properties, such as boiling points, and polarity. These
differences result in some challenges when fabricating the glucose biosensor, such as uniform
drying of the film. This section highlights the effect of increasing amounts of water used to prepare
the ethanolic Ir / Nafion®/GOx inks on the glucose response signal after ink deposition and drying,
to demonstrate the problem. The MM parameters, imax and K’m, were therefore compared for Ir
oxide (IrOx)/Nafion®/GOx films fabricated from inks containing varying H2O:EtOH ratios.
The Michaelis-Menten (MM) plots were generated from chronoamperometric measurements
as a function of the glucose concentration in a deaerated (Fig. 5.8a) and aerated (Fig. 5.8b) pH 7,
0.1 M PBS at 1.2 V vs. RHE, where the IrOx/Nafion®/GOx films were fabricated from inks
containing 0, 10, 20 and 50 vol. % water. It is clearly evident from Figure 5.8 (a and b) and Table
5.1 that the glucose signal (imax) achieved for each sensing film is enhanced with increasing water
content in the IrOx/Nafion®/GOx ink. The imax values for films fabricated from inks having a low
water content (10%) are small, ca. 10 µA cm-2, whereas the inks with a high water content (50%)
resulted in films with substantially higher imax values, up to ca. 70 µA cm-2, in deaerated solutions,
and ca. 300 µA cm-2 in aerated solutions. The IrOx/Nafion®/GOx film fabricated from a water-
free ink showed practically no glucose signal at any glucose concentration, showing that a
functional sensing film cannot be made when GOx is subjected to 100% organic solvent conditions
during film preparation.
101
Figure 5.8. Michaelis-Menten (MM) plots for ethanol-based IrOx/Nafion®/GOx films, formed
from inks containing 6 g L-1 GOx and 0 (♦), 10 (■), 20 (●), and 50 wt. % (▲) water and tested for
their glucose response in (a) deaerated (with Ar) and (b) aerated (with O2), stirred, neutral
phosphate buffer solution. Error bars represent the standard deviation of the glucose signal for
three sensing films.
0 5 10 150
25
50
75
0%
10%
20%
i / A
cm
-2
[Glucose] / mM
(a)
50%
0 5 10 150
100
200
(b)
i / A
cm
-2
[Glucose] / mM
0 %
10 %
20 %
50 %
50%
20%
10%
0%
102
Table 5.1. MM parameters for thin films formed from inks containing variable vol. % water in
aerated (O2) and deaerated (Ar) solutions (see Figure 5.8). (Concentration of all other components
was constant in a 4:1:1 volume ratio of IrOx:Nafion®:GOx).
Solution MM parameter a 10% water 20% water 50% water
Ar imax a 11 ± 13% 40 ± 3.0% 66 ± 29%
O2 imax a 11 ± 9% 135 ± 15% 290 ± 11%
imax(O2/Ar) b 1.1 3.4 4.4
Ar K’m a,b 5.4 2.9 2.0
O2 K’ma,b 5.8 5.7 5.9
a. The units for imax are µA cm-2 and the units for K’m are mM.
b. Maximum standard deviation of ± 15 %
It is important to note that, for GOx to be active, it must first be stable in its surrounding
environment. Secondly, under deaerated conditions, GOx must also be in good electrical contact
with a well-interconnected IrOx NP matrix in the dried films (Scheme 5.1) so that the GOx active
site (flavin adenine dinucleotide) may be easily regenerated via direct-mediation after reaction
with a glucose molecule (Reactions 2.2 and 2.8). In an aerobic solution, if GOx and the IrOx NPs
have good inter-connectivity, the imax values should be similar to those obtained in an anaerobic
solution, as the electrons should preferentially be transferred to the IrOx matrix (Scheme 5.1). The
O2 route (Reactions 2.2 and 2.3) is slow in comparison to direct mediation through the IrOx matrix
(Reactions 2.8, 2.11 and 2.9)88. However, if GOx and the IrOx NPs have poor inter-connectivity
(Scheme 5.1), the electrons will be transferred preferentially to O2 and the glucose signal will be
higher in an aerated vs. deaerated solution, giving a high imax ratio between the two solutions, as
seen in Table 5.1 for higher H2O contents in the sol.
In Figure 5.8a, the glucose signal is seen to increase significantly in the deaerated solution
for films fabricated from inks of increasing water content, indicating that there is more active
103
enzyme in the ink, that there is more GOx in good contact with the Ir NPs, or a combination of
these factors. However, the glucose signal ratio (imax(O2/Ar)) in aerated vs. deaerated solutions
(Table 5.1) for these films also increases, indicating that, as the amount of water in the ink
increases, the glucose response of the resulting sensing films will tend to be a result of O2-
mediation. Interestingly, the degree of irreproducibility also increased substantially when the water
content was increased from 20 to 50 vol. %, which could be expected for films with non-uniform
film morphologies (Fig. 5.7) and poor GOx/IrOx electrical connectivity.
Films fabricated from inks containing ≥ 50 vol. % H2O were also tested for their response to
glucose. However, the films were not stable and did not adhere well to the Au substrate (Fig. 5.7)
with rinsing (with water) between experiments. For this reason, the glucose response for films
fabricated from still higher (≥ 50 vol. %) water content films is not shown.
As can be seen in Table 5.1, the increase in water content does not significantly affect the
K’m values in the aerated solution (Fig. 5.8b), while K’m is noticeably affected in the deaerated
solution (Fig. 5.8a). Under these deaerated conditions and with increasing water content, the K’m
value decreases, indicating that the GOx that is in contact with IrOx is regenerated more quickly.
These results contradict the higher imax ratio observed with increasing water content (Table 5.1),
as it would be assumed that, with an increase in O2-mediation, GOx regeneration via direct
mediation via IrOx (Reactions 2.8, 2.11 and 2.9) would be slower. It is likely, however, that this
discrepency (and the signal irreproducibility between films) is due to the non-uniform
morphologies, and poor adhesion to Au that result from the drying of high water:ethanol volume
films (Fig. 5.7).
From the glucose signal data alone (Fig. 5.8 a and b, Table 5.1), it can be concluded that inks
containing 20% water contents form the optimal sensing films. They exhibit the good
104
characteristics displayed by films formed from high water-content inks (high imax) while
maintaining the desired characteristics exhibited by films formed from low water-content inks
(high degree of direct electron transfer mediation through the IrOx NPs, and good reproducibility).
To better understand these results, the effect of GOx on the IrOx dispersion in the sensing film,
the IrOx redox kinetics and the glucose signal were studied.
5.5 Effect of GOx:Ir mass ratio on glucose sensing
In Section 5.3, it was suggested that glucose oxidase (GOx) and Ir form interactions that
prevent the hydrogen underpotential deposition (Hupd) reaction from taking place at the Ir
nanoparticles (NPs) (Fig. 5.4) that are not converted to oxide in the sensing film. If the GOx
molecules are in close proximity with the Ir NPs in the ink/film to prevent the Hupd reaction from
taking place, it is likely that GOx is within close enough contact for direct electron transfer. It is
therefore important to determine the mass ratio of Ir NPs to GOx that is required to achieve direct
electron transfer between the GOx active sites and the Ir oxide (IrOx) NP matrix, thereby giving
the lowest imax ratios. Figure 5.9a shows the Michaelis-Menten (MM) plots for sensors containing
variable GOx:Ir mass ratios, ranging from 0.2 to 2. As expected, imax increases with increasing
GOx loading up to a GOx:Ir mass ratio of 1, as more GOx is available for reaction with glucose.
However, a further increase in the amount of GOx relative to IrOx results in a dramatic decrease
in the biosensor performance (Fig. 5.9b). The signal decreases significantly, from 36 to 6 µAcm−2,
likely due to insufficient IrOx sites available to fully ‘wire’ or connect with the enzyme (i.e., a
fourth-generation biosensor, Scheme 2.2d).
105
Figure 5.9. (a) Michaelis-Menten plots of electrodes composed of GOx/IrOx mixture with mass
ratios of 0.25 g/g (1, ▼), 0.50 g/g (3, ▲), 1.00 g/g (2, ●) and 2.00 g/g (4, ■). Glucose testing was
performed at 1.2 V in a stirred pH 7 0.1 M phosphate buffer solution under Ar-saturated conditions.
Aliquots consisted of 50 µl; of 2 M glucose, and (b) Plot of imax and K’m values as a function of
GOx:Ir mass ratio based on data in (a), with points calculated using Eadie–Hofstee plots.
In Figure 5.9 (a and b), the K’m values, as calculated from an Eadie-Hofstee plot (Section
2.5) range from 5 to 13 mM. Similarly, the ‘wired’ biosensors developed by Heller’s group236-238
were characterized by small K’m values, as low as ca. 6 mM for the Os-based polymer237 and 16.8
mM when using polyaniline. It can be anticipated, therefore, that the right sort of interconnectivity
0.0 0.5 1.0 1.5 2.00
10
20
30
40
GOx:Ir mass ratio
i max /
mA
cm
-2
0
10
20
30
40imax
K' m
/ m
M
K'm
(b)
106
within the film will have a major impact on the pathway of the reaction and can result in a change
in the enzyme kinetics vs. what occurs in the O2 mediated enzyme regeneration mechanism. In
order to get the best interactions or ‘wiring’ between GOx and the IrOx NP matrix it was found
here (Fig. 5.9, a and b) that a GOx:Ir mass ratio of 1.0 is optimal.
Ir NPs (even in the presence of H2O) were found to be primarily in the metallic form in GOx-
free inks, as evidenced by the small 1-3 nm NP size distribution demonstrated in transmission
electron microscopy images (Fig. 4.1a). These NPs were shown to be abundant and uniformly
dispersed throughout the film. Interestingly, when GOx is added to the ink, the NPs tend to form
aggregates near GOx (Fig. 5.3). As such, the electrochemistry of Ir/GOx films formed from inks
with increasing GOx content (0- 17.5 g l-1) were studied.
The Ir/Nafion®/GOx films (of varying GOx concentrations) were electrochemically cycled
to as high as 1.05 V in pH 7, 0.1 M phosphate buffer solution (PBS), so as to not electrochemically
convert the metallic Ir to IrOx (Fig. 5.10). The charge in the Hupd peak of the cyclic voltammetry
(CV) should indicate the degree of interconnected Ir NPs in the sol. It was observed that, as the
amount of active GOx in the Ir/Nafion® ink increased (at a constant H2O content of 20 vol. %), the
charge in the Hupd peaks clearly decreased (Fig. 5.10). Also seen in Figure 5.10 is that, after
complete electrochemical oxidation of Ir to IrOx in the sensing film, the amount of interconnected
IrOx NPs decreased (anodic peak current (ipa) decreased) with an increasing GOx content in the
ink.
107
Figure 5.10. Comparison of the charge in the Hupd peaks (prior to electrochemical oxidation of
Ir to IrOx) and the Ir(III)/Ir(IV) oxide anodic peak current (after complete conversion of Ir to IrOx)
for IrOx/Nafion®/GOx films as a function of increasing amounts of GOx (0 to 17.5 g l-1) in the
inks.
One explanation for the decrease in the IrOx ipa and charge in the Hupd peaks with increasing
GOx content (Fig. 5.10) is that the adsorption of GOx to the Ir NPs blocks the interconnectivity of
the Ir NPs. It has been shown in the literature232 that GOx gives an enhanced glucose signal in the
presence of Ag NPs, suggested to be the result of its adsorption to the Ag NPs and the resulting
decrease in the distance between the (flavin adenine dinucleotide) FAD active site and the NPs. It
was also shown that albumins, immunoglobulins, and cellular proteins (as well as many other
biological molecules) adsorb to metal oxide NPs, such as TiO2, ZnO, SiO2, and Au250,251. The TEM
images (Fig. 5.3) indicate that, in these films, GOx likely causes the aggregation of Ir NPs, thereby
decreasing the amount of Ir available to form the interconnected IrOx matrix.
As shown in Scheme 5.2, when there are fewer GOx molecules in the film, the IrOx matrix
is well-interconnected, and electron transfer from the GOx active site to the Au substrate via the
IrOx matrix is rapid and facile. However, when the amount of GOx in the film is increased, the
number of Ir NPs available for conversion to IrOx in depleted, preventing the formation of a well-
0 5 10 15
0.050
0.075
0.100
0.125
0.150
0.175
[GOx] / gl-1
I pa /
A
cm
-2
0
1
2
3
4
5
Ch
arg
e in
Hu
pd
pe
ak / m
C
108
Scheme 5.2 Electron transfer pathway through IrOx matrix when the amount of GOx in the sensing
film is increased.
interconnected matrix. As seen in Scheme 5.2, this poorly-interconnected matrix prevents the
regeneration of GOx via direct-mediation through the IrOx matrix, thereby resulting in a shift in
dual-mediation to favor O2-mediation over direct-mediation.
5.6 Effect of Nafion® on glucose sensing films
To determine the effect of Nafion® on glucose sensing, increasing amounts of Nafion® (1-8
wt. %) were incorporated into the sensing film and the films were then tested for their response to
glucose. Nafion® and glucose oxidase (GOx) are both large non-conducting, carbon-based
compounds. Nafion® (1 wt. %) makes up ca. 12% of the volume of a typical glucose sensing film,
while GOx occupies ca. 75% of the total IrOx/Nafion®/GOx film volume.
While large amounts of Nafion® were found to decrease the Ir electrochemistry signal for an
Ir oxide (IrOx) film fabricated from an ethanolic Ir sol (Fig. 4.10a), the polymer did not affect the
rate of electron, proton or solution movement through the film. The decrease of charge in the
Ir(III)/Ir(IV) oxide peaks indicate that large quantities of Nafion® blocked IrOx interconnectivity,
and fewer IrOx nanoparticles (NPs) were able to undergo redox chemistry.
109
Accordingly, the effect of increasing Nafion® on the resulting glucose signal of the film
was studied to determine how the changes in the IrOx matrix affected glucose sensing. As expected
(small amounts of Nafion® were found to enhance IrOx interconnectivity in Ir films (Fig. 4.10 a),
small amounts of Nafion® in the ink do not affect the glucose signal (imax) in a deaerated solution
(Fig 5.11a, Table 5.2), as both films (0 and 1 wt. % Nafion®) achieve imax values of ca. 40 µA/cm2.
When the Nafion® content was increased further, however, it is seen that the imax values decreases
pointedly (to as low as 4.7 µA/cm2 for the 7.5 wt. % Nafion®) (Fig. 5.8a).
Figure 5.11. Glucose response for IrOx/Nafion®/GOx films with increasing Nafion® content (0,
1, 2, 4 and 8 vol. %) in (a) deaerated and (b) aerated, stirred, 0.1 M, pH 7 phosphate buffer solution.
The K’m values were seen to increase (> 2x as high, Table 5.2) with increasing Nafion®
content in the ink (from 1 to 8 wt. %), indicating that the regeneration of the enzyme via direct
electron transfer through the IrOx NPs is slower under these conditions. In Chapter 6, it will be
shown that higher K’m values are indicative of O2 regeneration of the flavin adenine dinucleotide
(FAD) active site of GOx, related to inadequate IrOx NP interconnectivity. The glucose signal
obtained from these IrOx-based sensors in an anaerobic solution (Ar-saturated) depends entirely
on the interconnectivity of the IrOx matrix and the proximity of the IrOx NPs to the GOx active
sites (Scheme 5.1). For example, if the distance between an IrOx NP and a GOx molecule were
0 5 100
10
20
30
40
i / A
cm
-2
[Glucose] / mM
0 %
1 %
2 %
4 %
8 %
(a)
0 5 100
40
80
120
160
i / A
cm
-2
[Glucose] / mM
0 %
1 %
2%
4%
8%
(b)
110
Table 5.2. Comparison of the average Michaelis-Menten parameters (K’m and imax) for
IrOx/Nafion®/GOx films (ca. 2.5 µm thick) with increasing wt. % Nafion® content in deaerated
(Ar) and aerated (O2), 0.1 M, pH 7 phosphate buffer solution.
MM
parameter
0%
Nafion®
1%
Nafion®
2%
Nafion®
4%
Nafion®
8%
Nafion®
Ar imax a
40 39 16 8.8 4.7
Ar K’m b
2.6 2.0 4.9 5.5 4.5
O2 imax a
170 138 89 31 17
O2 K’m b
5.9 5.9 5.2 6.7 6.6
imax(O2/Ar) b 4.3 3.5 5.6 3.5 3.6
a. The units for imax are µA cm-2.
b. The units for K’m are mM.
c. All errors are ≤ 15%.
to increase from 8 to 17 Å, this would result in a decrease in the electron transfer rate by 104
times232.
The significant loss of glucose signal in the aerated solution with increasing Nafion® content
(Table 5.2), however, indicates that the blocking of the IrOx matrix by Nafion® may not be the
only reason for this loss in signal. Although the current would be expected to decrease with a
poorly-interconnected matrix, H2O2 is mobile, and therefore, will diffuse to and oxidize at the
electronically conductive IrOx pathway, shifting dual-mediation toward O2-mediation (resulting
in an increase in the imax ratio with increasing Nafion® content). It is possible that Nafion® blocks
glucose from reaching the enzyme active site, resulting in a smaller than expected glucose
response, or that it somehow interacts with GOx, inactivating it.
From these results, it can be said that small amounts of Nafion® (≤ 1.0 wt. %) shift dual-
mediation toward direct-mediation, while large amounts of Nafion® cause a shift towards O2-
mediation. Likely, the increase in glucose signal via direct-mediation with the addition of 1 wt. %
Nafion® can be attributed to the increase in the IrOx NP interconnectivity in the IrOx films
observed in Fig. 4.11a with the addition of 1.0 wt. % Nafion® to the Ir NP ink. The optimal amount
111
of Nafion® that should be added to the films is 1 wt. % if direct-mediation is the preferred
mechanism for GOx regeneration.
5.7 Summary
It was found in this thesis work that the active site of glucose oxidase (GOx) in the Ir oxide
(IrOx)/Nafion®/GOx sensing films can be regenerated by two different mechanisms (dual-
mediation), where electrons can be transferred directly from the active site to the adjacent IrOx
NPs, or to O2 (Scheme 5.1). The focus of the work in this chapter was to understand the effect of
the sensing film components on the preferential electron transfer pathway used to regenerate GOx.
It was found by transmission electron microscopy, X-ray photoelectric spectroscopy (XPS)
and electrochemical analysis that GOx aggregates the Ir NPs in the ink, blocking NP
interconnectivity and preventing the Ir NPs from being completely oxidized. The aggregation of
the Ir NPs severely affects the IrOx matrix, resulting in diffusion controlled transport. Therefore,
too much GOx in the sensing film affects the ability for electrons to be transferred directly through
the IrOx NP matrix (increasing the amount of O2-mediation that takes place). The optimal amount
of GOx in the sensing films was found to be a mass ratio of GOx:Ir of 1, which allowed for the
largest directly-mediated glucose signal.
It was shown that the water/ethanol composition of the Ir ink is an important parameter
influencing GOx activity, as high ethanol contents in the ink resulted in the denaturation of GOx,
leading to low glucose signals. When the water content of the inks was increased, the O2-mediated
glucose signals (a result of poor contact between GO and the IrOx matrix) were irreproducible and
the films were not stable on the Au substrate. As such, an optimal water concentration in the inks
was determined to be 20 vol. %, which gave good glucose signals, reproducibility and good direct
mediation of electrons through the IrOx matrix.
112
The addition of too much Nafion® to the sensing films was found to severely affect direct
mediation and O2-mediation, presumably by blocking IrOx interconnectivity (as was seen in
Section 4.4.2). Small amounts of Nafion® (1 wt. %), however, were found to be optimal for direct-
mediation as well as O2-mediation.
The understanding of how the glucose sensing film components affect the regeneration
pathway allows for the modification of the film to favor one mechanism vs. the other. One of the
problems with glucose sensing via O2-mediation is that, in practise, O2 levels in the blood can
fluctuate, resulting in poor sensing accuracy (Section 2.1.3). As such, the findings in this chapter
were used to fabricate a sensor that senses glucose independently of the partial pressure of oxygen
in the sample solution, thereby increasing the accuracy of the sensor, especially for implantation
applications (Section 2.1.1). The theory as well as the techniques used to optimize an O2-
independent response, will therefore be discussed in Chapter 6.
113
CHAPTER SIX: TOWARDS A RELIABLE AND HIGH SENSITIVITY O2-
INDEPENDENT GLUCOSE SENSOR BASED ON IRIDIUM OXIDE
NANOPARTICLES*
*Sections 6.1, 6.3.1 and 6.3.2 were published in Campbell, H.B.; Elzanowska, H.; Birss, V. I.
Biosensors and Bioelectronics 2013, 42, 7.
*Section 6.2 was published in Jhas, A.; Elzanowska, H.; Sebastian, B.; Birss, V. Electrochimica Acta
2010, 55, 7.
6.1 Introduction
In Chapter 4, it was seen that an Ir nanoparticle (NP) film with a uniform morphology can
be fabricated from an ethanolic Ir sol, and the Ir can readily be electrochemically oxidized to Ir
oxide (IrOx) NPs. These IrOx NPs exhibited rapid electron transfer kinetics, making them an
excellent electronic “wiring” matrix for the regeneration of glucose oxidase (GOx). In Chapter 5,
an IrOx NP/Nafion®/GOx sensing film that produced a glucose signal in both aerated and
deaerated solutions was fabricated. In the deaerated solution, the GOx molecules are regenerated
by transferring electrons to the interconnected IrOx electron transfer matrix (direct-mediation). In
the aerated solution, the GOx molecules are regenerated via direct mediation, as well as via the
O2-mediation route (dual regeneration, Scheme 5.1). The effect of the film components on sensing
film morphology, IrOx redox kinetics and glucose sensing were studied to determine what ratios
produced the highest and most reproducible glucose signals in both aerated and deaerated
solutions.
A common problem encountered with biosensors that detect enzymatically-produced H2O2
is their dependency on O2, which can cause measurement error due to O2 concentration fluctuations
in blood in practice. To overcome this obstacle, a significant amount of research has been focused
on the development of artificial (non-physiological) mediators (M), such as ferrocene
derivatives252,253, which can replace O2, leading to second-generation glucose sensors (Section
114
2.3)239,254, or the use of a wired, or fourth-generation (Section 2.3) enzyme-mediator
conjugates237,239,254.
One of the complications related to fabricating an ideal fourth-generation enzymatic
glucose sensor is that, in the presence of O2, the sensor can behave as both first and fourth-
generation, due to the transfer of electrons from the enzyme to either O2 or directly to the electrode,
respectively. This dual mechanism of glucose sensor functioning results in challenges associated
with the unwanted production and detection of H2O2, as was previously recognized in the early
stages of third-generation sensor development209,210,255, as well as in recent work10,40.
Amine et al.255, reported dual mediation in sensors composed of GOx, together with
phenazine methosulfate and ferricyanide mediators. It was found that, at low glucose
concentrations, O2 behaves as the main electron mediator. However, as the concentration of
glucose is increased, the non-linear increase in the current was attributed to a decrease in the local
O2 concentration at the surface of the electrode due to its reaction with glucose. This was explained
by the competition of the scarce O2 and the abundant mediators for the enzyme active site. It was
therefore assumed that the reaction with O2 was faster than with the mediator because the O2
pathway dominated at low glucose concentrations. A dual-mechanism (dual-mediation) of active
site regeneration was also observed by the same group for xanthine oxidase. In more recent work,
carbon nanotube based biosensors, containing Nafion® and Pd256 or Au115 NPs have been found to
exhibit small Michaelis-Menten (MM) K’m values. This indicates that the O2 pathway, in contrast
to Amine et al.’s255 sensor, is less efficient than the enzyme reaction with other mediators.
Specifically, it is shown in this chapter that it is possible to determine which GOx
regeneration (oxidation) pathway, aerobic (O2-mediated) or anaerobic (electron transfer directly
from the IrOx NPs), is dominant from the magnitude of the K’m value. To shift the sensing films
115
from being regenerated via dual-regeneration, to being O2-independent (direct-mediation only),
several approaches for fabricating uniform and reproducible film morphologies were examined. It
is shown that the preference of the sensor to regenerate itself via electron transfer from O2 or the
IrOx NPs is controlled by the sensing film thickness in both aerobic and anaerobic environments.
While sonication of the ink prior to film deposition did not result in more reproducibility between
films, it did increase the film sensitivity to glucose.
6.2 Results and discussion
6.2.1 Indicators of direct electron transfer and regeneration of GOx
6.2.1.1 Effect of O2 on glucose signal
When the Ir oxide nanoparticles (IrOx NPs) are uniformly distributed, well-interconnected,
and have good electronic connectivity to all glucose oxidase (GOx) molecules in the film,
regeneration of the GOx active site should occur directly through the IrOx NPs (Scheme 5.1),
independent of the presence of O2. In this chapter, it will be shown that, with the optimal sensing
film fabrication procedures, the IrOx-based matrix is capable of detecting glucose almost
independently of O2. As a method to prove that the glucose signal will not fluctuate substantially
with changing concentrations of O2, the glucose signal was first monitored in a completely
deaerated solution, and then half way through the experiment, O2 was bubbled into the solution.
As is seen in Figure 6.1a, with the addition of O2 after the addition of ca. 25 mM glucose, the
glucose signal does not jump, but rather remains similar to what would have been expected had
O2 not been introduced to the solution.
116
Figure 6.1. Chronoamperometry plot at 1.2 V for (a) a (nearly) O2-independent glucose sensor
and (b) an O2-dependent glucose sensing film. Glucose additions (50 µL of 2M glucose) were
made to a stirred, pH 7, 0.1M phosphate buffer solution (25 mL) under Ar-saturated conditions for
the first 7 min. The solution was then fully aerated and no notable change in the sensor response
to glucose was seen.
When a similar experiment was performed with an O2-dependant IrOx-based film (Fig
6.1b), the enhancement in glucose signal with the addition of O2 to the solution was significant, as
in this “non-optimized film”, many GOx molecules were not in good contact with the IrOx NPs.
As such, the flavin adenine dinucleotide (FAD) active site could not be regenerated until a mobile
mediator, such as O2, was introduced into the environment. With the introduction of O2, all of the
previously inactive GOx molecules (those in the FADH2 state) could be regenerated, resulting in
more active enzyme, thereby increasing the glucose signal.
This observation highlights why it is important to fabricate a sensor that is O2-independent.
When there is an unknown number of GOx molecules that will only react with glucose if sufficient
O2 is present, it would be difficult to calibrate the glucose signal. The rest of this chapter is focused
on understanding how direct electron transfer through the IrOx matrix occurs, and how we can
tailor the sensing films so as to decrease the O2 sensitivity of this IrOx-based glucose sensor by
monitoring direct electron transfer through the IrOx matrix, as was demonstrated in Fig. 6.1b.
0 5 10 15 200
20
40
60
80
i / A
Time / min
(b)
Bubbled
O2
Deaerated
117
6.2.2 K’m and imax values as an indicator of GOx regeneration mechanism
Figure 6.2 shows a set of chronoamperometric data collected at an Ir oxide
(IrOx)/Nafion®/glucose oxidase (GOx) electrode that highly favors O2-mediated regeneration over
direct electron transfer via the IrOx NP matrix. Data were collected at a constant potential of 1.2
V (vs. RHE) in a stirred pH 7, 0.1 M phosphate buffer solution (25 mL), under both aerated and
deaerated conditions. Based on the results in Section 5.4.3, the Ir sol was composed of 20% H2O
and 80% EtOH, and a GOx:Ir mass ratio of 1 (Section 5.5). Glucose additions (20 μL glucose
aliquot of a 1.0 M β-D-glucose solution) were performed first in a deaerated (Ar-saturated) solution
and then in a fresh, aerated (O2-saturated) phosphate buffer solution (PBS), both well-stirred at all
times in order to properly mix the glucose additions into the bulk of the solution. Notably, the film,
estimated to be ca. 2.5 µm in thickness, was not allowed to dry between glucose additions or when
Ar was replaced by O2 to prevent any changes in the film structure.
In Figure 6.2, upon the addition of each glucose aliquot, a rapid increase in the current is
observed (Fig. 6.2 inset) until a steady-state for that concentration is reached. Once a steady-state
current has been achieved, a second aliquot is then added to the solution. All of the currents are
solution stirring rate independent. As the glucose concentration was increased (to > 15 mM) under
aerated conditions, the currents reached a plateau at the system steady-state, where the rate of
formation of the enzyme-substrate complex is equal to its breakdown (Fig. 6.2). This response is
expected, when glucose oxidation is governed by Michaelis-Menten (MM) kinetics. In the absence
of O2, the currents in Figure 6.2 plateau at a lower value of ca. 20 mA/cm2.
118
Figure 6.2. Chronoamperometric data obtained from a 2.5 µm thick IrOx/Nafion®/GOx film,
tested for its glucose response in an anaerobic and aerobic environment at 1.2 V. The current was
measured after the addition of 20 µL aliquots of 1.0 M glucose to the stirred pH 7, 0.1 M phosphate
buffer solution (25 mL). Inset shows the glucose response after the addition of the first glucose
aliquot in both aerobic and anaerobic environments.
Importantly, the IrOx/Nafion®/GOx electrode is seen in Figure 6.2 to respond to glucose
under both aerobic and anaerobic conditions, thus exhibiting the desired dual regeneration
response10,40,209,210,255, as was discussed in Chapter 4. Superficially, the plots in Figure 6.2 appear
to be very similar, with the current initially rising and then approaching a plateau as the glucose
concentration increases. However, a closer look at Figure 6.2 reveals that the response of the
IrOx/Nafion®/GOx film under deaerated conditions actually differs quite significantly from that in
O2-saturated solutions. Initially, the current response to glucose under deaerated conditions
increases very rapidly until a glucose concentration of ca. 2 mM (ca. 300 s) is reached and then
the current reaches a constant value. In contrast, under aerated conditions, Figure 6.2 shows that
the rise in current still occurs at higher glucose concentrations and a plateau is not reached, even
0 600 12000
20
40
60
imax
140 160 180 200
4
6
8
10
i / A
cm
-2
time / s
O2
Ar
i / A
cm
-2
time / s
O2
Ar
119
at 15 mM glucose. This translates to very different K’m values for the Ir/Nafion®/GOx films of ca.
3 mM and ca. 25 mM in a deaerated and aerated solutions, respectively (Table 6.1).
In aqueous solutions, the free enzyme K’m value has been reported to be 20-30 mM in the
presence of O2238. Table 1 shows that, when the dominating regeneration pathway is O2-mediation,
the K’m values for IrOx/Nafion®/GOx films in O2-saturated phosphate buffer solution are roughly
in this range (23 to 32 mM), confirming that a similar mechanism is likely at play whether GOx is
in solution or resides in the IrOx-based film. However, in the Ar-saturated environment (IrOx-
mediated glucose oxidation), K’m is much smaller, being 2.5 to 3.5 mM. In comparison, the K’m
values reported in Chapter 5 were for films that did not have such a high O2-dependancy, and as
such, those K’m values were smaller for the glucose response in the aerated solutions.
Table 6.1. Comparison of the Michaelis-Menten parameters (K’m and imax) for three
IrOx/Nafion®/GOx films (ca. 2.5 µm thick) with Nafion® in aerated and deaerated glucose-
containing 0.1 M, pH 7 phosphate buffer solutions.
Gas
Environment
MM parametersa Trial 1 Trial 2 Trial 3 Average
values
Ar imax 19 14 18 16 ± 2.8
Ar K’m 4.6 2.7 2.7 3.3 ± 1.1
O2 imax 161 161 115 146 ± 27
O2 K’m 24 32 23 29 ± 6.2
imax(O2/Ar)b 8.5 11.5 6.4 8.8 ± 2.1
a. The units for imax are µA cm-2 and the units for K’m are mM.
b. Ratio of average limiting currents (imax) in O2 vs. Ar saturated environments.
In many cases10,40,209,210,255, the current response increases slowly with increasing glucose
concentration, owing to the competition of O2 and the mediator for the enzyme cofactor and large
K’m values are then observed. Mano et al.238, however, reported that the MM plots in aerated and
deaerated conditions were similar in appearance and that the K’m values were small (< 10 mM),
indicating the efficient mediation of the GOx reaction by the electrode matrix (e.g., PVP-[Os(N,N′-
alkylanated-2,2′-biimidazole)3]2+/3+ complexes). The very low K’m values obtained in the present
120
study in the Ar- vs. O2-saturated environments (Table 6.1) are therefore very strong evidence that
the regeneration of the GOx enzyme can occur via the IrOx sites.
The conventional equation for K’m (Equation 2.4) is obtained by assuming that the enzymatic
reaction is at steady-state, i.e., that the rate of formation of the enzyme-substrate complex (ES) is
equal to the rate of product release. Indeed, conventional MM kinetics do not take into account the
regeneration of the redox enzyme active site before the enzyme can bind a second substrate. As
enzyme regeneration is key to the performance of an electrochemical glucose sensor, the steady-
state representation in Reaction 2.14 has been modified. As shown in Reaction 6.1, another rate
constant, k3, which represents the regeneration of the FAD site within GOx has been added for the
regeneration of the active site. This rate constant (k3) will be included in the measured K’m value,
causing K’m to be smaller as k3 becomes larger.
𝐸𝑂𝑋 + 𝑆 𝑘1→
𝑘−1←
𝐸𝑂𝑋𝑆 𝑘2→ 𝐸𝑅𝐸𝐷 + 𝑃
𝑘3→ 𝐸𝑂𝑋 (𝑅𝑒𝑎𝑐𝑡𝑖𝑜𝑛 6.1)
In the anaerobic pathway, k3 represents the rate of electron transfer from the reduced FAD
site in GOx to an IrOx NP and the removal of protons. In the aerobic pathway, k3 represents several
additional processes, such as the rate of the binding of O2 to the FAD site, the formation of H2O2,
and the H2O2 oxidation reaction. This concept has been addressed previously by other groups209,210
using poly(pyrrole) as a direct mediator for GOx. Since k3 will therefore be different under
anaerobic vs. aerobic conditions, especially as it is likely that these two processes will have
different rate determining steps, it is logical that there should be two different K’m values for the
sensor in the two environments. Regeneration via the anaerobic IrOx pathway involves fewer
steps, as opposed to the aerobic pathway, which involves multiple steps and the complications
related to O2 reduction, peroxide formation, and its subsequent oxidation. The K’m parameter
121
should therefore be smaller in the anaerobic case than for the O2 regeneration pathway, consistent
with what is observed in this thesis work (Table 6.1, Tables 5.1 and 5.2).
The imax values observed for the O2-mediation dominated IrOx/Nafion®/GOx films shown
in Figure 6.2 (and Table 6.1) in O2-saturated environments were generally larger than those
obtained under the deaerated conditions, with an average imax ratio (in O2 vs. Ar) of 9. This must
indicate that the majority of the GOx molecules are not well-connected to the IrOx NPs, as required
for GOx regeneration by direct electron transfer to IrOx. However, in aerobic environments, the
GOx molecules that are distant from the IrOx NPs can still be regenerated by O2 and thus higher
imax values are seen under aerated conditions. The maximum current response arises from the
product of the rate determining rate constant (k2) and the total concentration of enzyme. Therefore,
when there is more enzyme accessible for regeneration (by O2), the total current increases
proportionally.
The parameters that are required for it to be said that a sensing film exhibits complete O2-
independence include imax (O2:Ar) and K’m (O2:Ar) ratios of 1. The glucose signal should be equal
in both aerated and deaerated environments, as GOx regeneration should only occur through the
IrOx NPs, and the number of active IrOx NPs in the sensing film remains constant. K’m values
should be equal as well, as all of the rate constants in Equation 6.1 should be the same if
regeneration occurs solely through the IrOx NPs.
6.3 Film fabrication methods to enhance O2-independence
6.3.1 Effect of film thickness on O2-independence and sensitivity
One of the ultimate goals of this work is to miniaturize the sensor by decreasing its size by
ca. 100 times. This smaller sensor is expected to be used in a patch containing an array of sensors
in order to continuously monitor blood glucose levels once per hour, for a full 24 hours. Therefore,
122
the Ir oxide (IrOx)-based layer must be very sensitive in order to generate measurable currents
from the much smaller sensor area. For this reason, the effect of the sensing layer thickness was
examined in this work, as varying this parameter is a simple approach to modifying the signal. All
electrodes were prepared from the standard ink containing a 4:1:1 volume ratio of Ir
sol:Nafion®:glucose oxidase (GOx) (20% H2O and 80% EtOH). Films were fabricated by
depositing various aliquot volumes of the ink (1 to 6 µL) onto a 0.20 cm2 area of a Au-coated glass
slide, giving a layer thickness ranging from 0.7 to 4.0 µm.
Figure 6.3 shows that, as the ink volume and thus the sensing film thickness increases, the
IrOx peak currents increase accordingly. In fact, a linear relationship is seen between the IrOx
anodic CV peak current densities (A1) and the amount of IrOx on the electrode surface, at least to
a film thickness of ca. 3 µm, as shown in Figure 6.3 (inset).
Figure 6.3. Cyclic voltammetry (0.01 V s-1) showing effect of the IrOx/Nafion®/GOx film
thickness (dictated by the size of the aliquot of ink deposited on the Au surface) on IrOx CV
response in deaerated, quiescent, pH 7, 0.1 M phosphate buffer solution. Estimated film thickness
is 0.66 µm (1), 1.30 µm (2), 2.7 µm (3) and 4.0 µm (4). Inset: IrOx anodic peak (A1) current density
vs. the film thickness.
0.0 0.6 1.2-2
0
2
0.0 1.0 2.0 3.0 4.0
2.0
3.0
A1 p
ea
k i / m
A c
m-2
Film Thickness / m
C1
i /
mA
cm
-2
E / V vs. RHE
A1
123
These films were then tested for their response to glucose in both aerobic and anaerobic
environments. It is shown in Figure 6.3 for deaerated conditions that, as the aliquot volume
increases and as the IrOx CV peak current densities increase (Fig. 6.3), the maximum current
response (imax) during glucose sensing also increases in a linear fashion. This demonstrates that the
direct regeneration of GOx by IrOx nanoparticles (NPs) is not hampered in the thicker films, which
is a very positive outcome. A higher sensitivity can clearly be obtained by increasing the film
thickness, with a current density of ca. 225 µA cm-2 achieved for the thickest film investigated
here. Importantly, the anaerobic K’m values remain relatively small, in the range of 3–7 mM, for
all film thicknesses, similar to what was reported in Table 6.1 for IrOx/Nafion®/GOx films.
Figure 6.4a shows that, in the aerobic environment, the glucose response also increases
with sensing layer thickness, as expected. Figure 6.4a also shows that there is substantial scatter
in the glucose response under these conditions, more so than in Figure 6.4b. This is presumed to
be related to the visually observed non-uniformity of the IrOx/Nafion®/GOx coatings, especially
for the thicker films. As a result of the relatively low ink viscosity, it spreads poorly on the Au
substrate. The thicker films dry more slowly, which probably allows the film components more
time to aggregate. This will cause the connectivity of GOx to the IrOx nanoparticles (NPs), of IrOx
NPs to each other, and of IrOx NPs to the Au substrate, to be variable, thus causing some
irreproducibility in the direct regeneration of GOx in the absence of O2, consistent with Figure
6.4b.
124
Figure 6.4. Current density response (imax) of IrOx/Nafion®/GOx films of varying thickness to
glucose, plotted against the IrOx CV peak current (peak A1 in Fig. 6.3) in (a) aerated and (b)
deaerated conditions.
In aerated conditions, even aggregated GOx that is distant from the IrOx NPs will still be
able to regenerate by the direct reaction with O2. However, the resulting rough surface morphology
will interfere with the collection of H2O2, formed during O2 regeneration of GOx (Reaction 2.2).
Any H2O2 produced can only be registered at IrOx NPs that are fully connected to the underlying
Au substrate, as the Au electrode itself is fully blocked by the sensing layer materials. As the
solution is stirred during glucose analysis, and due to the rough surface morphology, the amount
of H2O2 that is lost into solution vs. being trapped in the sensing layer and registered is expected
to be quite variable (Scheme 6.1). Thin films may lose more H2O2 to the bulk solution, while
thicker films are more likely to trap H2O2 within the film, resulting in a greater number of moles
of H2O2 reacting at the IrOx NPs vs. being lost to the bulk solution. The irreproducible film
morphologies result in varying degrees of thin and thick portions of the film. This model could
therefore explain the scatter in the data in Figure 6.4a. The response is therefore more reproducible
for the IrOx regeneration pathway (Fig. 6.4b) because H2O2 is not a by-product and thus does not
need to be collected in the sensing layer and subsequently oxidized.
0 1 2 3 4 5
0
200
400
i max /
A
cm
-2
i (IrOx) / A cm-2
(a)
0 1 2 3 4 50
100
200
i max /
A
cm
-2
i (IrOx) / A cm-2
(b)
125
Scheme 6.1 Shows the effect of film thickness on the collection and oxidation of H2O2 at the
sensing film on a Au substrate.
6.3.2 Effect of sonication of ink on O2-independence
To attempt to minimize the issue of non-uniform morphology formation with drying (Fig.
5.7), sonication (for 2 hours) was employed to more thoroughly mix the ethanolic Ir oxide (IrOx)
and Nafion® solutions before the addition of the aqueous glucose oxidase (GOx) solution to the
mixture, which is immediately followed by the deposition of the mixture on the Au surface. All of
these trials (non-sonicated and sonicated Ir sol/Nafion® inks) were carried out using the same
aliquot volume (4 µL) and the same 4:1:1 volume ratio of Ir sol:Nafion®:GOx (20% H2O and 80%
EtOH). Figure 6.5a shows that the CVs for the three films formed from the sonicated Ir/Nafion®
ink overlay each other very well, giving essentially the same IrOx charge density at all sweep rates.
This observation is in agreement with the visual observation that thorough mixing of the ink by
sonication, prior to film deposition, results in a more uniform surface layer, which would be
expected to have good IrOx NP interconnectivity. Films that were formed from inks that were not
sonicated, however, did not give as reproducible IrOx CVs (Fig. 6.5b), thus indicating that
sonication does indeed improve the interconnectivity of the IrOx NPs and their contact to the
underlying Au substrate.
126
Figure 6.5. (a) and (b) CVs (0.1 V s-1) of 2 µm thick IrOx/Nafion®/GOx films in deaerated, stirred,
0.1 M, pH 7 phosphate buffer solution and (c) and (d) corresponding Michaelis-Menten plots in
aerated (●) and deaerated (■) solutions. The inks were either sonicated for 2 hours (a) and (c) or
non-sonicated (b) and (d) prior to film formation. In the Michaelis-Menten plots, each point
represents the addition of a 20 µL aliquot of 1.0 M glucose to the phosphate buffer solution (25
mL).
Films formed from Ir/Nafion® inks that were either sonicated or non-sonicated before
deposition on the Au substrate were then tested for their response to glucose in aerobic and
anaerobic environments. Figs. 6.5(c and d) show that there is little difference in the reproducibility
of the response as a function of ink sonication, thus suggesting that the problem may be related
more to GOx aggregation during drying, rather than with the distribution/drying of the IrOx
component, as seen from Figure 6.5a. However, an important benefit of sonicating the inks prior
0.0 0.6 1.2-2
0
2
i /
mA
cm
-2
E / V vs. RHE
(a)
0.0 0.6 1.2-2
0
2
4
i /
mA
cm
-2
E / V vs. RHE
(b)
0 5 10 150
40
80
120
160
i / A
cm
-2
[Glucose] / mM
O2
Ar
(c)
0 5 10 150
20
40
60
i / A
cm
-2
[Glucose] / mM
O2
Ar
(d)
127
to deposition/drying (Fig. 6.5c) is that the imax values are always higher (by approximately a factor
of 2.5) than without sonication (Fig. 6.5d), thus leading to improved sensor sensitivity. The film
sensitivity is ca. 4 µA cm-2 per mM glucose, whereas the sensitivity increases to ca. 10 µA cm-2
per mM glucose after sonication of the ink.
Noticeably, the O2-dependence of the glucose signal also decreases with the sonication of
the film (from an imax of ca. 5 to 4), indicating that the improved reproducibility of the IrOx NP
film morphology results in better electronic connectivity between GOx and the IrOx NP matrix.
Overall, while sonication may result in increased activity and sensitivity of the immobilized
enzyme, it does not enhance the reproducibility of the glucose signal. Work is currently underway
in our group to improve the ink characteristics and the deposition methods used in order to
overcome this problem.
6.3.3 Spin-coat deposition of sensing films
Spin-coating is a common procedure used to deposit uniform thin films on a flat substrate
from a liquid precursor257,258. A 10 μl aliquot of an Ir/Nafion®/glucose oxidase (GOx) ink was
aliquot-deposited on the center of a sputtered Au substrate, and the Au was rotated using the
following protocol developed by Li et al. 259, for Pt/carbon/Nafion® catalyst deposition: 100 rpm
during the first 5 s, 1000 and 3000 rpm in the next 10 and 5 s, respectively, and a final speed of
3000 rpm was maintained for an additional 20 s (using a Photo-Resist Spinner (Laurell
Technologies Corp.)). With rotation, the ink spread over the Au via centrifugal force, with excess
ink being spun off the edge of the Au, depositing a thin Ir/Nafion®/GOx film. The resulting film
thickness depends on the rotational speed, as well as the viscosity of the solution and solvent. As
these were all kept constant, the film thickness should be the same for all three samples257.
128
As can be seen in Figure 6.6a, the spin coating process did not produce a uniform
Ir/Nafion®/GOx film. Coffee stains (similar to those observed with non-uniform drying in Figures
4.6b and 4.9 were evident around the edges of the film, radiating outward from the initial
deposition position of the Ir/Nafion®/GOx aliquot. The film is, however, very thin (as typically the
Ir is visible by the naked eye), and there are no large Ir nanoparticle (NP) aggregates, as seen in
Figure 6.6a. Three Ir/Nafion®/GOx films were deposited using an identical procedure, and the Ir
was then electrochemically oxidized to IrOx. The CVs (Fig. 6.6b) indicate that the amount of
interconnected Ir oxide (IrOx) NPs in the sensing films is not reproducible, likely due to varying
amounts of the ink being propelled off the Au substrate.
Figure 6.6. (a) Optical image of a spin-coated Ir/Nafion®/GOx film on a sputtered Au substrate,
(b) CV (100 mV s-1) of three spin coated IrOx/Nafion®/GOx films in stirred, deaerated, pH 7, 0.1
M PBS, and (c) Average Michaelis-Menten response to glucose for films in (b).
0.0 0.4 0.8 1.2
-0.4
0.0
0.4
0.8
i /
mA
cm
-2
E / V vs. RHE
(b)
0 4 8 12 160
10
20
30
40
Ar
i / A
cm
-2
[Glucose] / mM
O2
(c)
129
The glucose signal for the spin-coated films was found to be highly O2-dependant, with an
imax ratio of ca. 10. The glucose signal reached an imax of ca. 31 μA cm-2 in aerated solutions, while
the glucose signal via direct mediation reached an imax of only ca. 3.8 μA cm-2. The reproducibility
of the glucose signal in the aerated solution was very poor with glucose signals having a standard
deviation as high as ca. 14 μA cm-2. Likely, this large error is due to variable amounts of GOx
being spun off of the surface, or poor connectivity between GOx and the IrOx NPs. The K’m values
were also very different, being 5.6 and 1.8 mM in aerated and deaerated solutions, respectively,
confirming that the GOx regeneration pathways are different. Although there was no visible
aggregation of the IrOx NPs in the film, it is likely that this is due to the thinness of the film and
thus the distribution of the NPs and GOx is not uniform, leading to a high O2-dependence. The
irreproducibility may also be attributed to the film thickness, as H2O2 may be lost to the bulk
solution prior to reaction at the IrOx NP matrix, as was observed in Section 6.3.1 (Scheme 6.1).
Overall, the spin-coating procedure used in this work is therefore not ideal for depositing
Ir/Nafion®/GOx sensing films on the sputtered Au substrate.
6.4 Summary
A stable, highly porous, and biocompatible glucose sensor, based on a composite thin film
of Ir oxide nanoparticles (IrOx NPs), Nafion® and glucose oxidase (GOx) deposited on a Au
substrate, has been developed in this work. After reaction with glucose, the reduced enzyme can
be regenerated by O2, or by direct electron transfer to the interconnected, redox-active, IrOx NPs.
The fabrication of these sensing films can be manipulated so as to highly favor one regeneration
mechanism. In this chapter, the benefits of glucose sensing that is independent of the partial
pressure of O2 in the sample solution was the driving factor toward the fabrication of an O2-
independent glucose sensor.
130
To highlight the difference between a highly O2-dependent glucose sensing film and a film
with relatively low O2-dependence, the glucose response to the bubbling of O2 into the solution
midway through a glucose test in deaerated solution was performed. The current signal exhibited
for the film with high O2-dependence increased significantly, while the signal was not substantially
affected for the film that exhibited good O2-independence.
It was determined that small K’m values are indicative of rapid regeneration of the GOx
active site via direct electron mediation through the IrOx matrix. In contrast, larger K’m values
(seen in aerated solutions) result from the slower rate of GOx regeneration by O2 due to the
increased number of reaction steps that must take place before the electron is collected at the Au.
Accordingly, the goal of this Chapter was to fabricate films that exhibit small, identical K’m values
in both aerobic and anaerobic solutions. Various techniques were utilized in an attempt to fabricate
sensing films with reproducible, uniform morphologies and good electronic connectivity between
GOx and the interconnected IrOx NP matrix. These techniques included manipulating the film
thickness, sonicating the ink prior to film deposition, and using the spin coat procedure to deposit
uniform films.
It was found that imax increased linearly as the IrOx/Nafion®/GOx film thickness increased
in the deaerated solutions. The reproducibility of the glucose response, however, was poor in
aerobic solutions, likely due to the variable H2O2 loss associated with non-uniform film
morphology. Sonication of the IrOx/Nafion® ink prior to film deposition resulted in improved IrOx
NP interconnectivity. The glucose signal increased with ink sonication, resulting in a significant
increase in sensitivity (by ca. 2.5 times), as well as a decrease in the O2-dependence of glucose
sensing. Lastly, the spin-coating technique was utilized to deposit the sensing film on the sputtered
Au substrate. This method, however, produced irreproducible sensing films with a high
131
dependence on O2 for glucose sensing. This is likely the result of the thinness of the films and a
poor connectivity between GOx and the IrOx NP matrix.
Unfortunately, the culmination of all of the work done in Chapters 4-6 was not sufficient to
produce a film that exhibited complete O2-independent glucose sensing. Consequently, a new film
deposition method was designed. Chapter 7 will outline the use of diazonium-derived organic
layers to immobilize GOx on a sputtered Au electrode, so that the Ir NP ink can be uniformly
deposited within the pores of the film, with the goal of enhanced reproducibility and an O2-
independent glucose response.
132
CHAPTER SEVEN: DIAZONIUM-IMMOBILIZATION OF GOX IN AN
IRIDIUM OXIDE-BASED GLUCOSE BIOSENSOR
7.1 Introduction
The ultimate goal of this work is to fabricate an Ir oxide (IrOx)-based glucose biosensor
that accurately and reproducibly senses glucose, independent of the O2 partial pressure in the
sample solution. In Chapters 5 and 6, a glucose biosensor was fabricated by depositing an aliquot
of an ink containing IrOx nanoparticles (NPs), Nafion® and aqueous glucose oxidase (GOx) onto
a Au substrate. These sensing films were found to be robust, biocompatible, selective, and
sensitive.
It was found that there are two routes by which GOx can be regenerated after reaction with
glucose (dual regeneration), direct mediation via the IrOx NP matrix, and O2-mediation. By
improving the interconnectivity between the IrOx NPs in the film (by fabricating thick, uniform
film morphologies), a nearly O2-independent sensing film could be developed. It was also found
that O2-independent films should exhibit low K’m values and imax (glucose signal in aerated vs.
deaerated solutions) ratios of 1. However, neither the spin-coating (Section 6.3.3) or aliquot-
deposition (Chapters 5-6) methods were capable of producing films with a sufficiently uniform
IrOx NP and GOx molecule distribution, and although significant headway was made (imax ratios
decreased from ca. 8 to 3.5), full O2-independent sensing was not achieved.
The goal of this chapter is therefore to preserve the excellent sensing characteristics of our
IrOx-based sensors, but to achieve more control of the film morphology and consequently achieve
complete O2-independence of glucose sensing. Thus, diazonium salt chemistry was used to deposit
an organic film on Au, followed by the covalent attachment of GOx, with the objective being
preventing GOx aggregation. The EtOH-based Ir ink was then to be uniformly disseminated
133
around the immobilized GOx, thereby minimizing the aggregation of the IrOx nanoparticles (NPs),
after electrochemical oxidation of Ir, as was observed when water/ethanol Ir inks dry (Fig. 4.6).
The attachment of nano to micro scale organic films onto metal and carbon surfaces has
been widely investigated, as there are many applications for such films, including for corrosion
protection260, biofuel cells261, and sensor fabrication262,263. An increasingly popular method of
grafting organic layers to surfaces is through the reduction of diazonium salts (Section 2.2.1.2).
Diazonium salts are a group of compounds that contain the functional group R-N2+, where R is
typically an aryl substituent. Diazonium salts are favorable for forming organic films on surfaces,
as they undergo rapid electroreduction as well as spontaneous adsorption264-266, as there are many
salts with a wide array of substituent groups66, and as the resulting organic film is robust267.
Common methods used to generate aryl radicals include cyclic voltammetry (CV)66,67,69,265,268,
spontaneous reduction66,264-266, and potentiometry66,269,270.
In this work, nitrophenyl groups were grafted to a Au or carbon substrate from a diazonium
salt solution (Scheme 7.1a i), and the nitro groups were subsequently electrochemically converted
to amines (Scheme 7.1a ii). The reduction of the nitro groups allowed for the semi-quantification
(rough determination) of the final phenyl-group surface coverage. To reduce steric hindrance
during enzyme immobilization, a linking compound was attached to the amine group. In this work,
the diazonium-derived layer was succinylated, as the succinic anhydride linker readily forms
amide bonds (Scheme 7.1a iii) and GOx was then bound to the succinic anhydride linker via an
amide bond (Scheme 7.1a iv and v). Suspended metallic Ir NPs (in a Nafion®/EtOH ink) were then
dispersed throughout the GOx bound-organic layer via aliquot deposition, resulting in a
biocompatible, interconnected pathway for rapid electron transfer and enzyme regeneration
(Scheme 7.1b).
134
Scheme 7.1. (a) Method of depositing the nitrophenyl diazonium-derived layer on sputtered Au,
the subsequent succinylation of the amino group, and immobilization of glucose oxidase (GOx).
(b) Shows GOx immobilized to the Au substrate via a diazonium-derived succinylated
aminophenyl linking chain. Ir/IrOx NPs are intercalated within the film, surrounding GOx,
allowing for direct electron transfer from the active site of GOx to the underlying Au substrate.
The immobilized GOx film did not give a glucose signal prior to the electrical wiring of
the active site to the Au substrate via the IrOx network. Once IrOx was (assumed to be) intercalated
within the organic/GOx film, a Michaelis-Menten response (Section 2.5) was observed in both
aerated and deaerated environments. It was also found that, with the addition of increasing amounts
of IrOx to the film, the current response increased in a linear fashion. Methods to deposit more
nitrophenyl groups to the Au substrate were also employed (such as increasing the number of CV
135
deposition cycles, using potentiometry, and using substrates with high surface areas) in order to
achieve the highest glucose current response, O2-independence, and fast enzyme regeneration.
7.2 Modification of Au with diazonium salt chemistry
7.2.1 Nitrophenyl grafting and subsequent reduction
Electrografting of diazonium salts via cyclic voltammetry (CV) has been shown to reliably
produce organic films66,67,69,265,268, giving irrefutable and reasonably quantitative evidence that
grafting has occurred. As such, CV was chosen as a starting point in this research. Figure 7.1a
displays the characteristic CV reduction peak of a nitrophenyl group when cycling in the potential
window of 0.6 V to -0.2 V vs. a pseudo Ag/Ag2O reference electrode (Scheme 7.1a (i)). During
the reduction scan, two large reduction peaks are seen at 0.11 and -0.093 V (Fig. 7.1a). It is
hypothesized271 that multiple peaks are the result of the nitrophenyl radical grafting to different
Au facets. The more positive peak corresponds to deposition on the Au(111) domains, while the
more negative peak corresponds to deposition on the Au(100) facets272. With subsequent cycling
(Fig 7.1a), the charge passed during reduction decreases, indicating that the majority of the passive,
organic layer was formed in the first cycle. If the lower potential limit was extended cathodically
of the reduction waves, the resulting surface becomes further passivated and negligible charge is
passed during subsequent scans. This electrochemical behaviour was found to be typical to what
others have observed58,66,132,262,273,274.
136
Figure 7.1. Cyclic voltammetry of (a) a gold electrode in deaerated 1.0 mM nitrophenyl diazonium
salt + 0.1 M tetrabutylammonium tetrafluoroborate /acetonitrile solution at a scan rate of 100 mV
s-1. Inset: Quartz crystal microbalance data, showing the decrease in frequency with each cycle.
(b) Two cycles (100 mV s-1), depicting the first and second cycle of the electrochemical conversion
of nitrophenyl to aminophenyl groups in deaerated, 0.1 M KCl /10 vol. % MeOH solution are
shown.
The quartz crystal microbalance (QCMB) technique (Section 2.6.1) was used in situ during
CV to determine the total mass of the nitrophenyl groups deposited on a sputtered Au surface (0.48
cm2) with potential cycling in a 1.0 mM nitrophenyl diazonium salt solution. The QCMB data
showed that, after a total of 3 cycles in the diazonium salt solution (Fig. 7.1a), a substantial
frequency decrease of 70 Hz had been achieved (Fig. 7.1a), corresponding to a mass increase of
1.2 µg cm-2. The mass deposited during each cycle decreased with subsequent cycles, from 800 to
260 to 180 ng cm-2.
The first deposition cycle and the corresponding mass change during the first cycle are
shown in Figure 7.2a. The Faradaic efficiency (calculated from the theoretical mass expected to
have been deposited, according to the charge passed during reduction, vs. the measured mass
change) was also found to decrease after the first cycle. The theoretical mass deposited per electron
passed during the electrografting of nitrobenzene to Au on the first scan (slope of mass gain vs.
charge plot, Fig. 7.2b) was calculated to be 1.3 x 10-3 g C-1 (assuming a one electron process), and
-0.2 0.0 0.2 0.4 0.6-0.25
-0.20
-0.15
-0.10
-0.05
0.00
0.05
-0.2 0.0 0.2 0.4 0.6
800
820
840
860
C3
C2
frq
ue
ncy /
Hz
E / V vs. Ag/Ag2O
C1
C1
C2
i /
mA
E / V vs. Ag/Ag2O
C3
(a)
-1.2 -0.8 -0.4 0.0
-200
-100
0
I /
mA
E / V vs. Ag/Ag2O
1st cycle
2nd cycle
(b)
137
experimentally found to be 3.6 x 10-4 g C-1. This difference indicates that, of the radicals being
produced at the electrode surface, only ca. 1/3 of them are adsorbing (by either physisorption or
chemisorption) to the Au surface. This low (30%) efficiency for the first cycle is similar to those
calculated from the electrochemical deposition of nitrophenyl groups, published by Laforgue et
al.66, Kullapere et al.274, and Haccoun132 (0.32, 0.37, and 0.11, respectively).
The decrease in both the deposited mass and the Faradaic efficiency with further cycling was
expected, as the surface becomes further passivated by the growing organic layer, and radical
reactions must occur farther out in solution. After 3 cycles, the total mass deposited corresponded
to a surface coverage of ca. 10 nmol cm-2. Assuming that the maximum surface coverage of one
monolayer of nitrophenyl groups is 1.2 nmol cm-2 58, approximately 10 densely packed nitrophenyl
layers were deposited on Au, with a minimum film thickness of 7 nm.
Brooksby et al.275, described the growth of the organic film from a diazonium salt solution
as the formation of islands, with subsequent lateral growth of the islands to form a semi-continuous
multilayered film, approximately 20% as dense as a close-packed monolayer. A lack of control of
the structure of organic layers formed by diazonium salt chemistry is well documented88. One
possible reason for this is that it is very difficult to capture all of the radicals generated at the
electrode, as the radicals perform side reactions, resulting in the formation of branched structures.
For this reason, the organic layer formed in Figure 7.1a is non-compact, and is thicker than 7 nm.
To our knowledge, there is little in-depth explanation of the QCMB data in the literature
pertaining to the deposition of diazonium salts on Au surfaces.51,52,56,65-67,69,72,274,276. In these
studies, it was observed that a large amount of mass had deposited during the first cathodic cycle
between the onset and end of the nitrophenyl reduction peak (0.34 V and -0.2 V, in Figure 7.1a,
respectively)66,67,274. However, as was also observed in this work, nitrophenyl deposition continues
138
Figure 7.2. (a) Mass gain overlapping the first cyclic voltammogram of a gold electrode in
deaerated 1.0 mM nitrophenyl diazonium salt + 0.1 M tertbutylammonium
tetrafluoroborate/acetonitrile solution (100 mV s-1), and (b) Corresponding plot of mass change as
a function of charge consumed during the first cycle in (a).
after the surface was passivated and only small currents pass (Fig. 7.2). Figure 7.2a shows that, as the
potential is swept anodically, very little charge is passed in the range of -0.1 to 0.34 V.
-0.2 0.0 0.2 0.4 0.6
-0.2
-0.1
0.0
E / V vs. pseudo Ag/Ag2O
i /
mA
0.0
0.2
0.4
0.6
0.8
CVm
ass /
g c
m-2
Mass
(a)
0.0 0.2 0.4 0.6 0.8 1.00.0
0.2
0.4
0.6
0.8
-0.19 V
mass g
ain
, g c
m-2
Q, mC cm-2
Slope: 0.359 g/mC
0.34 V
-0.09 V(b)
139
Figure 7.2b depicts the mass gained vs. the charge passed during the CV in Figure 7.2a.
Laforgue et al.66, found that, during the first reduction wave, ~160 ng cm-2 was deposited on the
Au substrate. When the potential was scanned 0.44 V further negative of the reduction wave, a
mass of ca. 150 ng cm-2 was deposited, even though the charge passed was small. Upon scan
reversal, an additional ~50 ng cm-2 of nitrophenyl groups was deposited. In fact, a larger mass gain
was observed when the surface was passivated (after the peak) than before. In the present work
(Fig. 7.2a), the scan was reversed at the end of the reduction wave. However, more mass (420 ng
cm-2) was deposited on the reverse scan (at the passivated surface) than during the cathodic peaks
(370 ng cm-2) (Fig. 7.2a). The reason for this deposition at a passivated surface is unknown.
7.2.2 Degree of Au coverage by diazonium-derived layer
Film porosity, or access to the underlying Au substrate by the Ir nanoparticles (NPs), is
imperative for obtaining a glucose signal that is proportional to the concentration of glucose in the
sample. To determine what fraction of the Au was blocked by nitrophenyl groups, a bare Au
electrode, two diazonium-derived nitrophenyl films (after 1 and 5 cycles in the diazonium salt
solution), and the nitrophenyl film (5 cycles) after glucose oxidase (GOx) immobilization, were
electrochemically cycled in 0.5 M H2SO4 within an electrochemical window that includes both the
signature Au oxidation and reduction peaks (ca. 1.6 V and 1.06 V vs. RHE, respectively). Figure
7.3 shows the cyclic voltammetry (CVs) obtained for each of these electrodes, demonstrating the
decrease in active/accessible Au surface sites as the number of CV cycles in the diazonium salt
solution is increased.
140
Figure 7.3. Cyclic voltammogram (0.1 V s-1) for bare Au and three nitrophenyl-modified Au
electrodes (after 1, 3 and 5 cycles in the diazonium salt solution) in deaerated 0.5 M H2SO4.
The charge calculated from the reduction peak of the bare Au electrode (Fig. 7.3) after a
scan at 100 mV s-1 from 0.05 to 2.0 V vs. RHE is 1.61 mC cm-2. After nitrophenyl group grafting,
the electrode was cycled in 0.5 M H2SO4 from 0.05 to 2.0 V vs. RHE (Fig 7.3). It was found that
the nitrophenyl groups deposited after one grafting cycling in the diazonium salt solution blocked
approximately 40% of the Au, with the charge in the Au reduction peak being reduced to only 0.96
mC cm-2. With increased deposition time (5 vs. 1 cycle), the charge in the reduction peak decreased
further to 0.58 mC cm-2 (60% blockage). Interestingly, with the immobilization of GOx, the degree
of Au blockage increased slightly only to 68%, indicating that GOx does not block the Au substrate
from the solution, allowing room for the Ir NPs to intercalate within the film (Scheme 7.1b), as
desired.
Downward et al.90, performed a similar experiment with methylphenyl and carboxyphenyl
films on Au. It was found that only 20-30% of the Au surface was blocked after grafting, indicating
0.0 0.5 1.0 1.5 2.0
-1.5
-1.0
-0.5
0.0
0.5
1.0
i /
mA
cm
-2
E / V vs. RHE
Au
Au after 1 cycle
Au after 5 cycles
Au after 5 cycles + GOx
141
that the nitrophenyl diazonium salt used in the present work gives extremely high Au coverages.
Lui et al.277, found that, when they deposited either 4-carboxyphenyl, 4-meraptobenzoic acid or 4-
meraptopropionic acid groups from diazonium salts on Au (to close to a monolayer coverage),
only a low degree of Au blockage (9-15%) was achieved. Lui et al. 277, hypothesized that these
films were very thin, and as such, do not provide a good electron tunneling barrier to the electron
transfer during Au redox electrochemistry.
As a consequence of multilayer formation, the cathodic charge passed during nitrophenyl
film formation correlates with maximal surface coverage277 and does not indicate the number of
functional groups available for GOx immobilization for three reasons. Firstly, some of the
deposited nitrophenyl groups are not strongly bonded and are thus lost during sonication and
rinsing. Secondly, not all of the nitro groups are converted to amino groups54,76, which are required
for the GOx immobilization process. The charge passed during the electrochemical conversion of
nitro to amino groups (Scheme 7.1a (i-ii) and Fig 7.1b) of ca. 2.12 ± 0.06 nmol cm-2, however, can
be used to estimate the number of deposited nitrophenyl groups that were converted to
aminophenyl groups (Fig 7.1b), giving a 20% conversion rate. The complete conversion of Ph-
NO2 to Ph-NH2 is not achieved, as some of the functional groups are converted to Ph-NHOH or
Ph-NHO274. Thirdly, not all of the amino groups are accessible for binding the succinic anhydride
linking compound, and therefore, subsequently, for binding GOx. Unfortunately, it is difficult to
determine the accessibility of the amino groups for GOx binding. Thus, in this work, surface
coverage will be more accurately (but still not definitively) defined as the number of aminophenyl
groups deposited on the Au substrate (as calculated from the nitro reduction CV peak), as opposed
to the number of nitrophenyl groups.
142
7.3 GOx attachment to diazonium-derived aminophenyl layer
Once the nitrophenyl groups were converted to aminophenyl groups, a succinic anhydride
linker was attached to the organic film via an amide bond (Scheme 7.1a (iii)). The free carboxylic
acid on the succinic anhydride was then activated by immersing the organic film in 0.1 M, pH 5.5
2-(N-morpholino)ethanesulfonic acid (MES) buffer containing 1.0 mM N-hydroxysuccinimide
(NHS) and 10 mM Ethyl-3-[3-dimethylaminopropyl]carbodiimide HCl (EDC) (Scheme 7.1a (iv)).
This activation involves the formation of a semi-stable NHS ester with the succinic anhydride
linker. The ester bond is amine reactive, and, as such, when an amine group from glucose oxidase
(GOx) comes into close enough contact with the structure, an amide bond is formed between the
enzyme and the succinic anhydride linker (Scheme 7.1a, iv)184.
It is difficult to compare the glucose signals (imax) obtained for these films to the aliquot-
deposited IrOx/Nafion®/GOx sensing films, as there are substantially more GOx molecules present
in the aliquot-deposited films. There are ca.1.5 x1014 molecules of GOx in the aliquot-deposited
films, while in the immob-GOx films, there can theoretically only be ca. 5.5 x1011 molecules on
the 0.35 cm2 Au substrate, assuming each GOx molecule covers a 65 nm2 area158 and that they are
immobilized in a close-packed monolayer. The experimental value, as measured with an analytical
balance, was found to be 9.0 x1011 molecules (on a 0.35 cm2 Au substrate), indicating that there
are ca. 1.6 times as many GOx molecules as would be found in a close-packed monolayer. From
Section 7.2, it is known that the diazonium-derived film does not form a defect-free film, and
therefore, likely forms branches, thereby allowing for increased GOx immobilization. It is also
possible that there is some water still present in the GOx film which may increase the film mass,
(even though the film was dried for 60 hours prior to weighing). It is also possible that some GOx
molecules are adsorbed on the Au substrate, and not, in fact immobilized. Assuming that the
143
measured mass is accurate, however, the amount of GOx attached is still ca. 170x less than the
amount of GOx present in an aliquot-deposited film.
7.4 Glucose testing
7.4.1 Glucose testing in the absence of IrOx
Reproducibility and selectivity are important for a glucose sensor, and, as such, it is
imperative that glucose only reacts at the active site of glucose oxidase (GOx), and not at the
underlying Au or at the organic film. Similar to what was done in Section 4.5, blank glucose
addition experiments were performed on a bare Au electrode, and on an aminophenyl-modified
electrode to determine if glucose would react non-specifically. A third blank glucose addition
experiment was performed on a GOx-immobilized electrode (no Ir oxide nanoparticles (IrOx
NPs)) to determine if a glucose signal would be observed with O2 as a mediator. The role of O2 is
to accept electrons from the active site of the enzyme, thereby regenerating the enzyme (Reaction
2.2). This reaction forms H2O2, which would be expected to subsequently oxidize (Reaction 2.3)
at the positively charged, underlying Au substrate (at a potential of 1.2 V vs. RHE), resulting in a
current proportional to the amount of glucose in the sample.
A small signal for the oxidation of glucose is observed (ca. 0.4 µA cm-2) on the bare sputtered
Au electrode (Fig. 7.4a). Some oxidation of glucose at Au is to be expected, as Au has been shown
in the literature to allow some oxidation of glucose. For example, it has been shown by Nikolaeva
et al.278, that glucose oxidation can occur at Au in neutral pH due to the presence of Au-OH
functional groups. It is believed that the H of the C1 of glucose adsorbs to the Au surface, and the
–OH group (on Au) aids in the oxidation of glucose to form gluconic acid. It is likely that surface
–OH functional groups are present on Au, especially as the potential of the Au electrode was held
at 1.2 V in an aerated media before and during the glucose testing experiment shown in Figure 7.4.
144
Figure 7.4. (a) Glucose signal (1.2 V) at bare Au substrate (■), aminophenyl-modified Au (●), and
immobilized GOx on Au (▲). Experiments performed in O2-saturated, stirred, 0.1 M, pH 7 PBS
(25 mL) with 0.8 mM additions of 1.0 M glucose, and (b) H2O2 oxidation signal (1.2 V) at bare
Au, and at immobilized GOx on Au. Experiments performed in O2-saturated, stirred, 0.1 M, pH 7
PBS (25 mL) with 2.5 mM additions of 3.0 M H2O2.
Although a linear response is obtained for the direct oxidation of glucose at the Au substrate, this
is not recommended for achieving an accurate glucose signal as other electroactive sugars or
components in the blood may also react at Au, giving a hypoglycemic reading.
When the Au substrate was modified with Ph-NH2 (Scheme 7.1a (i-ii)), a negligible glucose
signal was observed in comparison to the signal at bare Au (Fig. 7.4a). This smaller signal was
0 4 8 120.0
0.1
0.2
0.3
0.4
i / A
cm
-2
[Glucose] / mM
Au
Au-NH2
Au-immobGOx
(a)
0 5 10 150.0
0.2
0.4
0.6
0.8
1.0 Au
immob-GOx-Au
[H2O
2] / mM
I /
mA
for
Au
(b)
0
5
10
15
20
25
I A
for
imm
ob-G
Ox-A
u
145
expected, as ca. 50% of the Au surface was blocked by the diazonium-derived layer (Fig. 7.3a).
Interestingly, it is observed that, when GOx was immobilized to the modified structure, the current
response increased only slightly (in comparison to the modified film) (Fig. 7.4a). As the
experiment was performed in an aerated environment, it would be expected that O2 would
regenerate the GOx active site after reacting with glucose, thereby forming H2O2 (Reaction 2.2).
H2O2, in turn, should readily oxidize at the Au substrate279, resulting in a large glucose signal.
However, this is not observed.
This small glucose signal (Fig. 7.4a) can likely be attributed to a number of factors, including
the ca. > 50 % blocking of the Au surface by with the deposition of the organic film and the
immobilization of GOx. Secondly, the organic layer is likely tortuous, due to multilayer formation,
and, as such, H2O2 may transfer to the bulk solution before diffusing to and reacting at the Au
substrate. Finally, it is known in the literature that during first-generation glucose sensing, H2O2
oxidation can be inhibited if the electrode is contaminated279 with, for example, adsorbed glucose
oxidation reaction intermediate species280 .
Figure 7.4b shows the H2O2 oxidation signal (at 1.2 V) observed at a bare Au substrate and
at an immobilized GOx film on Au (0.35 cm2 area). It can be seen that the H2O2 oxidation signal
is substantially higher at bare Au (ca. 40x) vs. at the Au surface, blocked by the diazonium-
derived/immobilized GOx film. This would be expected, as it is known from Section 7.2.2 that
approximately 50% of the underlying Au is blocked by the diazonium-derived layer and the
immobilized GOx.
7.4.2 Glucose testing at IrOx/Nafion®/immobilized-GOx films
An aliquot of ethanolic Ir nanoparticle (NP)/Nafion® ink (ca. 130 nmol) was then deposited
on a dry, immobilized-glucose oxidase (immob-GOx) film (Scheme 7.1b), and the Ir NPs were
146
then electrochemically oxidized to IrOx, similar to our previous work (Chapters 4-6). Nafion® was
added to the ethanolic ink, as it was shown to enhance the redox kinetics of the resulting Ir oxide
(IrOx) NP film (Section 4.4.2). Figure 7.5a shows that the cyclic voltammetry (CV) for the immob-
GOx/IrOx/Nafion® film exhibits similar characteristics to an IrOx film CV. (Fig. 4.2a). Both films
exhibit Hupd peaks and the characteristic IrOx A1/C1 redox peaks (Fig. 4.2a). The similarities
between the IrOx NP CV and the immob-GOx/IrOx/Nafion® film CV (Fig. 7.5a) confirm that
there is good electrical contact between the IrOx NPs as well as between the IrOx NPs and the
underlying Au substrate.
In similar research, Dyne et al.281, studied the electrochemical properties of electrode-
monolayer-nanoparticle constructs. They observed that previously passivated electrodes (by thiol-
derived self-assembled monolayers, 2-11 carbons in length) became electrochemically active with
the addition of Au or Pt NPs on top of the organic film. Raman studies confirmed that the metal
NPs were located on top of the monolayer (not intercalated into it), and not in contact with the Au
substrate. It was concluded that the switch from being an electrochemically inactive electrode to
an active one could not be the result of direct physical contact between the NPs and Au, but that
electron transfer occurred between the NPs and the underlying substrate. It is believed (although
not confirmed) that, in our immob-GOx/IrOx/Nafion® films, some IrOx NPs are in direct physical
contact with the Au (Scheme 7.1b), as the film is quite porous. It is also possible, however, that
the IrOx NPs that are not in direct contact with the Au substrate are still capable of electron transfer
directly through the organic layer to the underlying Au substrate via electron tunnelling, as was
observed by Dyne et al281.
147
Figure 7.5. (a) Cyclic voltammetry of an immobilized-GOx/IrOx film (133 nmol Ir) (dashed line),
and a standard IrOx film (133 nmol Ir) (solid line), and (b) Glucose signal (at 1.2 V) at a bare Au
substrate (■), an IrOx film on Au (●), immobilized-GOx film on Au (▲), and an immobilized-
GOx/ IrOx film on Au (▼). Experiments performed in aerated 0.1 M, pH 7 PBS (25 mL) with 20
µL aliquots of 1 M glucose.
With the addition of the IrOx NPs to the modified-GOx film, a Michaelis-Menten (MM)
current response (Section 2.5) was observed with increasing glucose concentration in the aerobic
environment (Fig. 7.5b), with an imax (maximum current) of ca. 3.2 µA cm-2 and K’m of ca. 5.8
0.0 0.4 0.8 1.2-0.75
-0.50
-0.25
0.00
0.25
0.50
A1
i /
mA
cm
-2
E / V vs. RHE
Ir/IrOx
immobilized-GOx/Ir/IrOx/Nafion
HupdC
1
(a)
0 4 8 120
1
2
3
4
i / A
cm
�-2
[Glucose] / mM
Au
Au-IrOx
Au-immobilizedGOx
Au-immobilized-GOx/IrOx
(b)
148
mM. Figure 7.5b shows that the glucose oxidation signals are amplified when the IrOx NPs are
deposited (assumed to be intercalated within the film), as prior to the addition of the NPs, the
glucose oxidation signal is negligible. In order to prove that glucose oxidation (at the immob-
GOx/IrOx film) is due to glucose reaction with the enzyme, and not the spontaneous oxidation of
glucose on the high surface area IrOx NPs, the glucose signal at an IrOx film, in an aerated
environment, was also recorded (Fig. 7.5b). It is seen that IrOx is a much poorer catalyst for
glucose oxidation than is Au metal (Fig. 7.5b). Carbohydrate oxidation has been studied at many
different metals and oxides, including Au, IrOx, RuO2, Ag, Cu, Ni, Pt (to name a few), and it has
been found that glucose reacts differently at each of these metals or metal oxides282.
To determine if the glucose signal in Figure 7.5b is due to H2O2 oxidation at the IrOx sites,
or if direct mediation of electrons from the active site through the interconnected IrOx matrix is
taking place, the experiment was repeated under deaerated conditions (shown in Fig. 7.7). After
the solution was saturated with Ar for 20 minutes, a similar glucose signal to the signal detected
under aerated conditions is observed, with an imax of 2.9 µA cm-2 and K’m of 2.8 mM (Fig. 7.7b).
The small difference in the imax values, i.e., the small imax ratio (in O2 vs. Ar) of 1.1, demonstrates
that electron transfer is occurring predominantly from the immobilized enzyme through the
interconnected IrOx NPs to the underlying Au. This small ratio also indicates that the majority of
the current is passing via this route, even in air, suggesting a nearly O2-independent response.
Scanning electron microscopy (SEM) was performed on an immob-GOx/IrOx/Nafion® film
(Fig. 7.6). The distribution of IrOx and Nafion® is seen to be uniform over the immobilized GOx-
organic layer, contrary to what was seen in the optical images for GOx-containing Ir films in Figure
5.7 (aliquot-deposition method). This uniform distribution of the IrOx NPs is ideal, as it results in
good IrOx NP interconnectivity.
149
Figure 7.6. Scanning electron microscopy image of an immobilized-GOx/IrOx/Nafion® film after
glucose sensing.
The immob-GOx/IrOx/Nafion® film shown in Figure 7.6 contains ca. 130 nmol Ir, which
should be sufficient to completely coat and infiltrate the GOx/diazonium-derived organic layer.
However, it is unknown where the Ir NPs are located in the final sensing film, i.e., if they are
embedded in the film pores or just sitting on the top. In Scheme 7.2, two simplified models for Ir
NP intercalation into the immob-GOx film are suggested, showing either tight or loose stacking
around the GOx molecules. In Scheme 7.2a, 80 Ir NPs are shown to be stacked on a Au surface to
a film thickness of 10 nm (the expected thickness for a film fabricated from an aliquot of 88 nmol
Ir NPs). When 80 Ir NPs are intercalated within the immob-GOx film, they can form close
interactions with the GOx molecules (Scheme 7.2b), forming a tightly packed or compact film,
with a minimum film thickness of 14 nm. If the NPs do not interact so closely with GOx (Scheme
7.2c), it is possible that the Ir NPs may not fully surround the GOx molecules. Thus, there will be
an unnecessary amount of Ir NPs not participating in GOx regeneration, resulting in “wasted” Ir
material.
150
It is likely, that the ca. 1 nm dia. Ir NPs (Fig. 4.1 a and b) will, however, pack in closely to
the GOx molecules (probably with a packing density than as shown in Scheme 7.2b), as it is known
from Figure 5.3 that GOx causes the Ir NPs to aggregate in the vicinity of GOx. It is important to
note, however, that this scheme is very simplified, as the porosity and degree of branching of the
diazonium-salt layer, and the proximity of the GOx molecules to each other, are not taken into
account.
Scheme 7.2 Ir NP intercalation models for the deposition of the Ir ink into the diazonium-derived-
immobilized GOx film, prior to the electrochemical conversion of the Ir NPs to IrOx.
7.5 Further enhancement of glucose sensor response
7.5.1 Optimization of amount of Ir in the sensing film
Once it was verified that Ir oxide nanoparticles (IrOx NPs) are required to achieve good
glucose signals, another goal was to determine what quantity of IrOx NPs is optimum. Too little
IrOx likely would not fully intercalate the film, resulting in only a few interconnected, but very
151
thin, IrOx nanowires (Scheme 7.3, left). An optimal signal should be obtained when the IrOx
nanowires are thicker and well-interconnected (Scheme 7.3, right). It is of interest that the structure
on the left would likely result in sturdier IrOx nanowires, with an electrochemical surface area that
is lower than in the left-hand depiction. On the other hand, too much IrOx may result in wasted
material and potential diffusion control of glucose/gluconolactone to and from the buried,
immobilized GOx enzyme.
Scheme 7.3. Schematic representation of single IrOx nanowire (left) and a well-interconnected
and thicker IrOx nanowire, resulting from a higher IrOx loading. The structure on the right would
be expected to have a lower IrOx surface area (per gram) than that on the left.
The number of moles of Ir deposited on the immobilized GOx film was thus varied from 85
to 180 nmol Ir, corresponding to equivalent dense Ir film thicknesses of ca. 20 to 45 nm,
respectively (electrode surface area = 0.35 cm2). The metallic Ir NPs were then partially, or fully,
converted to IrOx via electrochemical cycling. Note that 1 wt. % Nafion® is present in the film, as
it was shown to increase the IrOx NP interconnectivity in films fabricated from ethanolic Ir inks
(Fig 4.10).
152
From the cyclic voltammetry (CVs) in Figure 7.7a, it can be seen that, with increasing
amounts of Ir deposited in the film, more of the Ir is retained in its metallic form after
electrochemical cycling, as evidenced by the persistence of the Hupd peaks observed between 0
and 0.3 V vs. RHE. For films made with 88 nmol Ir, there is no Ir metal remaining after
electrochemical cycling, as all of it was converted to IrOx, as seen by the large Ir(III)/Ir(IV) oxide
redox peaks at ca. 0.8 V. Interestingly, the charge in the Ir(III)/Ir(IV) oxide redox peaks appears to
be largest for the smallest addition of Ir into the film. This is likely a result of a broken IrOx NP
matrix, where the NPs are interconnected, but the surface area is very high, as the NPs are spread
thin across the film. When more Ir NPs are added to the film, they fill the gaps in the film,
increasing the points of IrOx interconnectivity, but slightly decreasing the overall Ir NP surface
area, thereby decreasing the charge in the Ir(III)/Ir(IV) oxide redox peaks.
153
Figure 7.7. (a) Cyclic voltammetry (100 mV s-1) in deaerated, stirred 0.1 M, pH 7 PBS, and (b and
c) Michaelis-Menten glucose response for immobilized-GOx/IrOx/Nafion® containing increasing
amounts of IrOx NPs (ranging from 88, 133, 176 nmol Ir). Experiments performed in stirred, (b)
deaerated and (c) aerated 0.1 M, pH 7 PBS (25 mL) with 20 µL aliquots of 1.0 M glucose.
It is seen that the glucose signal in the anaerobic environment increases with increased
amounts of Ir within the immobilized GOx film (Fig. 7.7b), with 176 nmol IrOx films giving the
highest imax value of 4.2 µA cm-2. This indicates that, more IrOx NPs in the film (ca. 40 nm film
thickness), results in better IrOx-GOx interactions. The K’m values (Table 7.1) are seen to decrease
with increasing amounts of IrOx, suggesting that, as the total volume of IrOx increased, electron
transfer between the active site and the electronically conductive NPs becomes more facile.
Interestingly, the imax ratio (Table 7.1) also decreases for films containing larger amounts of IrOx
0.0 0.4 0.8 1.2-1.6
-0.8
0.0
0.8
1.6
HupdC
1
88 nmol
133 nmol
176 nmol
i /
mA
cm
-2
E / V vs. RHE
A1
(a)
0 4 8 12 160
1
2
3
4
5
88 nmol Ir
133 nmol Ir
i / A
cm
-2
[Glucose] / mM
176 nmol Ir
(b)
0 4 8 12 160
2
4
6(c)
133 nmol Ir
88 nmol Ir
i / A
cm
-2
[Glucose] / mM
176 nmol Ir
154
NPs, indicating that electron transfer is occurring preferentially, directly through the IrOx NP
matrix, as opposed to via the O2-mediation route. These small imax ratios indicate that the
immobilization of GOx prior to the addition of the ethanolic Ir ink to the film is a successful
method of preventing a non-uniform/ aggregated morphology, which was found to result in a film
with high O2-dependence (Tables 5.1, 5.2 and 6.1).
Table 7.1. Michaelis-Menten values for glucose signals at immobilized-GOx/IrOx/Nafion®
sensing films in aerated and deaerated environments (Fig. 7.7).
mol IrOx
(nmol)
aK’m
(Ar)
aimax
(Ar)
aK’m
)2(O
aimax
)2(O
imax Ratiobvs. Ar) 2(O
88 3.3 1.7 ± 7% 5.4 3.5 ± 5% 2.1
133 2.7 3.1 ± 8% 5.6 3.5 ± 7% 1.1
176 2.5 4.2 ± 14% 4.7 6.0 ± 5% 1.4
a. The units for imax are µA cm-2 and the units for K’m are mM.
b. Ratio of average limiting currents (imax) in O2- vs. Ar- saturated environments.
It is also observed (Fig. 7.7 b and c and Table 7.1) that 133 and 176 nmol Ir additions to the
immob-GOx film resulted in small imax ratios, indicating that electron transfer primarily occurred via
the IrOx electron transfer matrix. In contrast, although the imax values are very good in comparison to
those shown in Tables 5.1, 5.2 and 6.1 for aliquot-deposited films, the low Ir NP-content films exhibit
more O2-dependence. It is proposed here that the reason for the enhanced glucose signal and the near
O2-independent response with an assumed increase in Ir NP intercalation, is that, with more IrOx NPs
in the films, the film is packed tighter resulting in denser Ir nanowires (as shown in Scheme 7.3, right).
When fewer IrOx NPs are intercalated into the film, however, the packing will be less dense and the Ir
wires will be thin (Scheme 7.3, left). The tightly packed and denser Ir nanowires equate to excellent
IrOx NP interconnectivity, with many electron transport pathways and rapid kinetics (Scheme 7.4c).
Thus, films containing larger amounts of IrOx NPs display high glucose signals (Fig. 7.7b and c). Less
Ir intercalation results in less dense packing, and less electron transfer pathways through the IrOx
matrix (Scheme 7.4 a and b), subsequently giving a lower glucose signal (Table 7.1).
155
Scheme 7.4. Direct electron transfer (via IrOx) pathways with increased film IrOx content
(assuming tight IrOx NP packing, Scheme 7.2b).
7.5.2 Potentiostatic vs. cyclic voltammetry method for grafting of nitrophenyl groups on Au
One of the goals of this work was to attain a sensing film that has a very high sensitivity to
glucose. In Chapters 5 and 6, films fabricated via aliquot deposition9,10 gave very high glucose
signals (imax) of up to ca. 50 µA cm-2 and 150 µA cm-2 in deaerated and aerated environments,
respectively. It should be taken into consideration, however, that the imax values cannot easily be
compared between the aliquot-deposited films and the immobilized-glucose oxidase (immob-
GOx) films discussed here, as there are less GOx molecules in the immob-GOx films. Therefore,
it would be beneficial to increase the number of immobilized GOx molecules in the film in order
to further enhance the glucose signal.
In order to determine the best method to deposit more nitrophenyl groups on Au, two
different electrochemical reduction approaches (cycling from 0.6 to -0.2 V, vs. holding at a
constant potential of -0.2 V vs. Ag/Ag2O pseudo reference) were compared. In the first approach,
the potential of the Au electrode was cycled in the diazonium salt solution for one, three or five
cycles before the resulting Ph-NO2 groups were electrochemically reduced to Ph-NH2. It was
156
found that, with an increase in the number of deposition cycles, there was an increase in the Ph-
NH2 surface coverage (Fig. 7.8a and Table 7.2).
Figure 7.8 (a) Cyclic voltammetry (100 mV s-1) for immobilized-GOx/IrOx/Nafion® sensing
films, where the diazonium-derived film was deposited in 1, 3 or 5 cycles (resulting in surface
coverages of 1.4, 2.1 and 3.4 nmol cm-2, respectively), and Michaelis-Menten glucose response in
both deaerated (b) and aerated (c) environments for the films in (a).
The decrease in the Ir oxide (IrOx) A1/C1 redox peak currents in the cyclic voltammetry (CV)
in Figure 7.8a, observed when the Ir nanoparticles (NPs) were (assumed to be) intercalated into
the diazonium-derived/immob-GOx film, indicates that films with a higher aminophenyl surface
coverage (Table 7.2) result in less interconnected IrOx NPs. Also, for sensing films with a higher
aminophenyl surface coverage, it is seen that there is a decrease in the charge in the Hupd peaks
0.0 0.2 0.4 0.6 0.8 1.0 1.2
-4
-2
0
2
i /
mA
cm
-2
E / V vs. RHE
1.4 nmol cm-2 Ph-NH
2
2.1 nmol cm-2
Ph-NH2
3.4 nmol cm-2 Ph-NH
2
(a)
0 4 8 120
1
2
3
4
5
i / A
cm
-2
[Glucose] / mM
1.4 nmol cm-2 Ph-NH
2
2.1 nmol cm-2 Ph-NH
2
3.4 nmol cm-2 Ph-NH
2
(b)
0 4 8 120
2
4
6
(c)
i / A
cm
-2
[Glucose] / mM
1.4 nmol cm-2 Ph-NH
2
2.1 nmol cm-2 Ph-NH
2
3.4 nmol cm-2 Ph-NH
2
157
(Fig. 7.8a), indicating a significant loss in Ir NP interconnectivity. It could be expected that a thick,
branched, diazonium-derived film could prevent good IrOx interconnectivity, whereas a thinner
film may allow for increased IrOx NP intercalation (Scheme 7.2b), leading to better NP
interconnectivity.
Using a thin, porous organic layer (1.4 nmol cm-2) for GOx immobilization (after 1 cycle),
it is likely that there are not many GOx molecules immobilized, as the maximum current signal in
the deaerated environment is quite small (ca. 2.3 µA cm-2) (Fig. 7.8b, Table 7.2). With increased
aminophenyl deposition, however (2.1 nmol cm-2), the glucose signal achieved via direct
mediation (deaerated environment) almost doubled to 4.2 µA cm-2. Additional aminophenyl
deposition (3.4 nmol cm-2) did not further enhance the glucose signal in the deaerated environment
(Fig. 7.8b).
The imax ratios (O2:Ar) indicate that the amount of IrOx present within the thin organic layer
(1.4 nmol cm-2) was sufficient to ensure that all of the immobilized GOx molecules are in contact
with the interconnected IrOx NP matrix, resulting in an O2-independent response (achieving the
major goal of Chapter 6 and 7). As the number of Ph-NH2 groups on the surface was increased,
the imax ratio also increased (from ca. 1 to 2, Table 7.2), indicating that, for thicker films, the
immobilized GOx molecules are not fully surrounded by the IrOx NPs and thus GOx regeneration
became slightly more O2-dependant. Overall, however, the imax ratios are still very small, < 2.0
(Table 7.2), especially in comparison to many of the films formed via aliquot deposition (Chapters
5-6), where highly O2-dependent films exhibited imax ratios of ca. 8 (Table 6.1) and the films that
showed the least amount of O2-dependence exhibited imax ratios as low as 3.5 (Table 5.1 and 5.2).
158
Table 7.2. Surface coverage of aminophenyl groups in the diazonium-derived film, deposited on
a Au substrate via either CV cycling or potentiostatically, and Michaelis-Menten values for the the
resulting immobilized-GOx/IrOx/Nafion® sensing films in aerated and deaerated solutions.
Method of diazonium
salt grafting to Au
Surface
Coverage /
2-nmol cm
mK’
(mM)
(Ar)
maxi
(µA
)2-cm
(Ar)
mK’
(mM)
)2(O
maxi
(µA
)2-cm
)2(O
Ratio maxi
vs. 2(O
Ar)
a1 CV cycle 1.4 4.7 2.3 7.7 2.6 1.1
a3 CV cycles 2.1 2.6 4.2 4.7 6.0 1.4
a5 CV cycles 3.4 2.4 4.0 5.1 7.0 1.8
bPotentiostatic 10 s 1.6 5.1 1.1 5.6 0.90 0.8
bPotentiostatic 30 s 1.9 4.8 2.5 5.5 2.8 1.1
bPotentiostatic 60 s 2.1 5.2 3.0 7.8 3.1 1
bPotentiostatic 120 s 2.2 5.2 3.4 5.8 4.3 1.3
a. The potential of the sputtered Au substrate was cycled in diazonium salt solution from
0.6 to -0.2 V vs. Ag/Ag2O pseudo-reference electrode.
b. The potential of the sputtered Au substrate was held constant at -0.2 V vs. Ag/Ag2O
pseudo-reference electrode.
In the second route, the nitrophenyl organic layer was deposited by holding the Au working
electrode at a constant potential of -0.2 V for periods of 10, 30, 60 or 120 s in the diazonium salt
solution. As predicted, the longer the time the potential was held, the more nitrophenyl groups
were deposited on the Au substrate (Table 2). With increasing time at -0.2 V, the CV began to
change shape and it appeared that the charge in the Ir(III)/Ir(IV) oxide redox peaks decreased
linearly (Fig. 7.9a). Interestingly, there were practically no Hupd peaks observed for any of the
immob-GOx/IrOx/Nafion® sensing films when the diazonium-derived layer was formed at a
constant potential. This was seen even though the surface coverage of the nitrophenyl groups (as
calculated by the charge in the reduction peak of the nitrophenyl to aminophenyl groups) was
159
Figure 7.9. (a) Cyclic voltammetry (100 mV s-1) of immobilized-GOx/IrOx/Nafion® sensing
films, where the diazonium-derived film was deposited at a constant potential of -0.6 V for 10 to
120 s in the diazonium salt solution, and the Michaelis-Menten glucose response in (b) deaerated
and (c) aerated solutions for films in (a).
essentially the same for the film formed by cycling 3 times in the diazonium salt solution vs. the
film formed by holding at -0.2 V for 60 s (2.1 nmol, Table 7.2). This suggests that the two different
electrochemical grafting methods produced different diazonium-derived film morphologies.
For the films formed using constant potential deposition, the glucose signal is again found
to increase with increasing nitrophenyl group surface coverage (ca. 1.1 to 3.4 µA cm-2) in deaerated
solutions (Fig. 7.9b, Table 7.2). This improvement in signal is likely due to an increase in the
0.0 0.4 0.8 1.2
-1.6
0.0
1.6
3.2
I /
mA
cm
-2
E / V vs. RHE
1.6 nmol cm-2 Ph-NH
2
1.9 nmol cm-2 Ph-NH
2
2.1 nmol cm-2 Ph-NH
2
2.2 nmol cm-2 Ph-NH
2
(a)
0 4 8 120
2
4
i / A
cm
-2
[Glucose] / mM
1.6 nmol cm-2 Ph-NH
2
1.9 nmol cm-2 Ph-NH
2
2.1 nmol cm-2 Ph-NH
2
2.2 nmol cm-2 Ph-NH
2
(b)
0 4 8 120
2
4 1.6 nmol cm
-2 Ph-NH
2
1.9 nmol cm-2 Ph-NH
2
2.1 nmol cm-2 Ph-NH
2
2.2 nmol cm-2 Ph-NH
2
i / A
cm
-2
[Glucose] / mM
(c)
160
amount of GOx immobilized at the diazonium-derived film. The glucose signal in the aerated
solution is similar to that in the deaerated environment (Fig 7.9c). The imax ratios are very small,
indicating that the immobilized GOx is in very good contact with the IrOx NPs and that direct
mediation is preferred to the O2-mediation route. The K’m values remain fairly constant for each
of the films, being slightly higher in the aerated environment (Table 7.2). The similarity between
the K’m values in the deaerated solution indicates that the GOx active site is being regenerated via
the IrOx NP network. The increasing imsx values therefore indicate that more GOx is present in the
film with increased grafting time, as desired.
When looking at these results as a whole, the following three parameters must be taken
into consideration: imax, K’m values, and O2-independence. As seen in Table 7.2, cycling the
potential of the Au substrate (3-5 times) in the diazonium salt solutions results in films that give a
high current response in deaerated environments (ca. 4 µA cm-2). Both diazonium-deposition
routes result in films that exhibit relatively low K’m values of ca. 2.5 mM, indicating rapid
regeneration of the active site (Section 6.2.2). The imax ratio, however, for the films formed in three
CV cycles was found to be lower than that for five CV cycles (1.4 vs. 1.8, Table 7.2). The films
fabricated from diazonium-derived layers deposited at a constant potential all exhibited an imax
value of ca. 1, as well. The most effective procedure for depositing the diazonium-derived layer is
therefore cycling from 0.6 to -0.2 vs. a Ag/Ag2O pseudo reference in the diazonium salt solution
for three complete CV cycles.
7.5.3 Increasing glucose response by increasing Au surface roughness
Once the optimal grafting procedure was identified, work was done to determine if the
signal could be further enhanced by manipulating the roughness factor of the underlying Au
substrate and thereby immobilizing more glucose oxidase (GOx) molecules. To study this, Au foil
161
was used, as opposed to the smooth sputtered Au surfaces used in Sections 7.2-7.4. The Au foil
was then either polished until shiny and smooth using 0.3 and 0.05 µm alumna slurry on microcloth
pads, or roughened by anodization.
It is known in the literature that, when Au is subjected to anodic oxidation in an aqueous
solution of a carboxylic acid or carboxylate, a uniform porous Au surface results192. Based on a
procedure outlined by Xu et al.283,284, linear sweep voltammetry was performed on the Au foil in
a 0.3 M solution of oxalic acid, immediately followed by holding the potential constant for varying
amounts of time (Section 3.2.1.1). A porous, roughened surface was achieved by first forming a
passive film, and then repeatedly breaking it down on the nanoscale, forming nanopores at the
breakdown sites192,284. Although the anodizing procedure was not found to be reproducible in terms
of obtaining identical Au surface roughness for each of the samples, it did result in varying degrees
of Au roughness. The real surface areas could then be calculated from the cyclic voltammetry (CV)
charge passed during Au oxidation and reduction.
As expected, the number of nitrophenyl groups that could be deposited increased linearly
with the increasing Au surface area/roughness (as calculated according to Section 3.2.1.4) (Fig
7.10a). This indicates that, although multilayers of nitrophenyl groups may form on the Au surface,
the increasing surface area allows for more Ph-NO2 groups to be deposited directly to the Au
substrate, which, in turn, results in more Ph-NH2 groups available for succinylation (Scheme 7.1a
iii) and subsequent GOx immobilization (Scheme 7.1a iv and v). Figure 7.10b shows that, as the
number of Ph-NH2 groups on the surface of Au was increased, the glucose signal also increases,
roughly linearly, up to an aminophenyl surface coverage of ca. 0.7 *1015 mol. At this point, the
current response begins to decrease substantially, likely as the number of Ph-NH2 groups become
excessive, resulting in a diazonium-derived film that sterically hinders GOx immobilization.
162
Figure 7.10. (a) The surface coverage of aminophenyl groups vs. the real Au surface area, and (b)
the glucose signal produced by immobilized-GOx/IrOx/Nafion® sensing films fabricated using Au
substrates of increasing roughness.
The K'm values for these films (not shown) were larger than those achieved by immob-
GOx/Ir oxide (IrOx)/Nafion® sensing films that were fabricated on sputtered Au substrates vs. the
roughened Au foil (K’m values ranged from 4.4 to 6.5 mM). This indicates that, by increasing the
roughness of the Au substrate, direct electron transfer between GOx and the IrOx nanoparticles
(NPs) is hampered. It is possible that this loss of GOx:IrOx connectivity is due to the tortuosity of
0.0 0.2 0.4 0.6 0.8 1.0 1.20.0
0.2
0.4
0.6
0.8
1.0
1.2
1.4
Su
rfa
ce
co
ve
rag
e o
f P
h-N
H2 /
mo
l *1
0-1
5
Real Au surface area / cm2
(a)
0.0 0.2 0.4 0.6 0.8 1.0 1.2 1.41
2
3
4
5
6
7
8
9
10
i max /
A
cm
-2 (
ge
om
etr
ic)
Surface coverage of Ph-NH2 / mol *10
-15
(b)
163
the anodized films (i.e., the IrOx did not infiltrate the pores, leaving some GOx unable to
regenerate via direct mediation through the IrOx NPs). It is also possible that the diffusion of
glucose through the rough film was impeded, decreasing the rate of glucose oxidation by GOx.
7.6 Glucose sensors based on an immobilized GOx/Ir oxide/Nafion® film on carbon
7.6.1 Deposition of immobilized-GOx/Ir oxide/Nafion® sensing film on carbon paper substrate
The motivation behind using a carbon substrate as opposed to the Au substrate was two-
fold. First, there are many different types of carbon that can be used as a substrate, all of which are
substantially less costly than Au. Second, very high surface area substrates can be achieved by
using carbon. The heterogeneity of many carbon surfaces, however, poses a limitation to the use
of carbon as a substrate for diazonium-derived film deposition. Au also often exhibits a superior
rate of electron transfer in comparison to carbon56.
There are several examples in the literature demonstrating the grafting of diazonium salts
to carbon surface54,56,57,61,62,65,73,88,264-266,268,270,275,285,286. However, the majority of these surfaces
are smooth (glassy carbon) and, as such, the available surface area for immobilization is
comparable to that of the sputtered Au used in this work. Thus, a higher surface area carbon
material, carbon paper (CP) (Section 3.2.1.2) was selected for the sensing film substrate.
A 0.25 cm2 piece of CP was electrochemically cleaned by cycling the potential in 0.5 M
sulfuric acid (0.05 to 1.0 V) until the electrochemical features of carbon (the
hydroquinone/quinone peaks, E1/2 of ca. 0.6 V vs. RHE) remained constant between the cyclic
voltammetry (CV) traces. Nitrophenyl groups were then electrochemically grafted to the surface
(CP-PhNO2) by applying a reductive potential sweep at the CP electrode from 0.6 V to -0.4 V vs.
an Ag/Ag2O pseudo reference electrode (Fig. 7.11a). The CV shown in Figure 7.11a has a
comparable shape to that seen for the deposition of nitrobenzene on a glassy carbon substrate in
164
Figure 7.11. Cyclic voltammetry (100 mV s-1) showing (a) the grafting of the nitrophenyl group
from the diazonium salt solution on a carbon paper substrate, (b) the subsequent conversion of the
nitro to amino groups, and (c) a comparison of the grafting process on carbon paper vs. Au
substrates.
the literature268,287. The CP-PhNO2 surface was then reduced to amino groups (CP-PhNH2) by
cycling from 0.2 to -1.2 V vs. an Ag/Ag2O pseudo reference electrode (Fig. 7.11b) in aqueous
KCl/10 vol. % MeOH solution. Both electro-reduction processes were reproducible within a
reasonable error of 15%.
The surface coverage of the aminophenyl groups on CP was calculated to be ca. 2.2 nmol
cm-2. This value is roughly equal to that of two close-packed layers of aminophenyl groups (with
a monolayer being ca. 1.22 nmol/cm2). This is a substantially larger number of deposited groups
than what was deposited on Au (0.79 nmol/cm2) (Section 7.5.1). This large difference in
-0.4 0.0 0.4
-6
-4
-2
0
I /
mA
cm
-2
E / V vs. RHE
(a)
-1.2 -0.8 -0.4 0.0
-30
-20
-10
0
10
20
i /
mA
cm
-2
E / V vs. RHE
(b)
-0.4 0.0 0.4
-6
-4
-2
0
2
i /
mA
cm
-2
E / V vs. RHE
Carbon Paper
Au(c)
165
aminophenyl surface coverage is likely due to the higher real surface area and porosity of CP in
comparison to the sputtered Au substrate (note that the current densities and surface coverages are
calculated using the geometric surface area).
When comparing the deposition CV for the Au and CP substrates (Fig. 7.11c), it is seen
that the reduction wave begins at a much more negative potential (ca. 0.27 V) on the carbon surface
vs. at Au. There is also only one large reduction peak at the carbon surface, as opposed to the two
peaks observed for the Au substrate. Benedetto et al.271, have suggested that the potential at which
the diazonium salt is reduced is directly related to the work function of the substrate, which fits
with the experimental evidence shown here, as the work function of Au is slightly higher at 5.1 eV
vs. 5.0 eV for carbon288. Gui et al.56, found that the 4-nitrophenyl diazonium salt deposited at
roughly +100 mV on a Au substrate as opposed to glassy carbon (with both surfaces polished to a
smooth mirror-like finish).
After the deposition of the nitrophenyl groups on the CP, the CP-PhNO2 CV becomes more
resistive in nature and small redox peaks (E1/2 = ~ 0.61 V vs. RHE) are seen (Fig 7.12a), which are
known to be due to hydroquinone/quinone redox chemistry289. When the nitro groups are
converted to amines, the CV still appears resistive. However, larger redox peaks are seen that
overlap the hydroquinone/quinone peaks (Fig 7.12a). These peaks are due to the redox response
of the Ph-NHOH and Ph-NO functional groups (that were not converted to Ph-NH2)268. After the
nitro groups have been converted to amines, glucose oxidase (GOx) and Ir oxide nanoparticles
(IrOx NPs) were deposited (respectively), using the same methods as were described previously
for Au (Section 7.3 and 7.4.2).
166
Figure 7.12. Cyclic voltammetry (50 mV s-1) of (a) bare carbon paper (CP), a nitrophenyl-CP film,
and an aminophenyl-CP film in deaerated, stirred, 0.5 M H2SO4 and, (b) CVs immobilized-
GOx/IrOx/Nafion® film (180 nmol IrOx) on CP in deaerated, stirred 0.1 M pH 7 PBS. Inset shows
the IrOx anodic and cathodic peak current vs. the square root of the sweep rate.
The CV (Fig. 7.12b) shows that the Ir NPs that were deposited around the immobilized
GOx molecules have good electronic contact with the carbon substrate, as seen from the large
A1/C1 peaks. The lack of hydrogen underpotential deposition (Hupd) peaks (Section 3.2.1.4) in the
region from 0 – 0.3 V in Fig 7.12b show that all of the deposited Ir NPs were electrochemically
converted to IrOx. The inset of Figure 7.12b shows that the electron transfer kinetics through the
IrOx matrix are diffusion controlled, as the Ir(III)/Ir(IV) oxide redox peak currents are proportional
to the square root of the sweep rate (up to a sweep rate of 200 mV s-1). Uniform IrOx films
fabricated from Ir NP sols on a sputtered Au substrate typically show surface controlled electron
transport kinetics, characterized by a linear ip vs. ʋ plot (Fig. 4.11b). The diffusion control
exhibited here is likely due to the blockage of carbon pores by the deposited organic layer, and or
GOx, preventing protons and electrons from being rapidly transported throughout the sensing film.
7.6.2 Glucose signal from a CP/Ir/Nafion®/immobilized GOx sensing film
When glucose oxidase (GOx) is immobilized to the diazonium-derived layer on the carbon
paper (CP) substrate and the Ir oxide nanoparticle (IrOx NP) matrix is formed around it, a
0.0 0.2 0.4 0.6 0.8 1.0-40
-20
0
20
40
i /
mA
cm
-2
E / V vs. RHE
CP
CP-PhNO2
CP-PhNH2
(a)
0.0 0.4 0.8 1.2
-1
0
1
2
0.2 0.4
-2
-1
0
1
2
i /
mA
cm
-2
/ V s-1
i / m
A c
m-2
E / V vs. RHE
(b)
167
Michaelis-Menten (MM) response to glucose is observed (Fig. 7.13). The maximum current (imax)
obtained for the film in the deaerated environment is ca. 10 µA cm-2 and ca. 30 µA cm-2 in the
aerated environment. The good response in the deaerated environment indicates that the IrOx NPs
are well-connected to the GOx molecules (Section 6.2.1.1). The imax ratio (imax in O2 / imax in Ar)
is relatively high (ca. 3), in comparison to the immob-GOx/IrOx/Nafion® films deposited on Au
(Table 7.2), indicating that the sensing film is significantly O2-dependent, likely due to the
diffusion controlled electron transfer kinetics through the IrOx NP matrix (Fig. 7.12).
Figure 7.13. Michaelis-Menten glucose response in both deaerated and aerated environments for
immobilized-GOx/IrOx/Nafion® sensing films on carbon paper. Experiments were performed in
stirred, pH 7, 0.1 M PBS.
When comparing the surface area of a GOx molecule to the surface area of a single
aminophenyl group, there should be approximately 1 GOx molecule for every 370 aminophenyl
groups (2.2 nmol cm-2 aminophenyl groups were deposited, Fig. 7.11b). As such, it can roughly
be assumed that there are 50 times as many GOx molecules present in the aliquot deposited films
(Chapters 5-6, Scheme 5.1) than there are immobilized to the surface of the CP. The glucose
response achieved for the aliquot deposited films, however, is only on ca. 5 times greater than the
signal achieved with the modified CP substrate.
0 4 8 120
10
20
30
i / A
cm
-2
[Glucose] / mM
Ar
O2
168
As such, it can be concluded that the GOx activity is higher in the immob-
GOx/IrOx/Nafion® film on the CP substrate than when it is trapped within the IrOx/Nafion®
matrix of the aliquot deposited film. However, the immob-GOx/IrOx/Nafion® film on the CP
substrate displayed a relatively poor sensitivity of only 2.23 μA cm−2 per mM glucose and 1.53
μA cm−2 per mM glucose in the aerated and deaerated environments, respectively, in comparison
to the aliquot deposited films that reached sensitivities of 10 mA cm-2 per mM glucose (Figs. 6.5
c and d), possibly due to the high background current exhibited by the CP. Although a
comparatively high GOx activity can be achieved by immobilizing GOx on a CP substrate via a
diazonium-derived film, the signal is not O2-dependent, as desired, and has low sensitivity.
7.7 Summary
Glucose biosensors that are on the market today suffer from O2-dependent glucose sensing
and related poor accuracy. In Chapter 6, an Ir oxide (IrOx)-based glucose biosensor that sensed
glucose, nearly independent of the O2 partial pressure in the sample solution, was developed.
However, it was shown to be difficult to fabricate a sensing film with a uniform morphology, as
glucose oxidase (GOx) and IrOx form aggregates during the drying process. To prevent
aggregation, in the present chapter, a diazonium-derived organic layer was deposited on a sputtered
Au substrate and GOx was immobilized by covalent attachment to the diazonium-derived layer.
An ethanolic Ir sol was then (assumed to be) intercalated into the immobilized-GOx film to provide
an electron transfer matrix for the direct regeneration of GOx.
The quartz crystal microbalance technique and cyclic voltammetry were used to determine
the surface coverage of the diazonium-derived layers, as well as to provide an indication of the
porosity of the film. Blank glucose testing was performed on the sensing film at each step of the
fabrication process to ensure non-selective glucose oxidation did not occur. During these tests, it
169
was observed that the immobilized GOx film did not respond to glucose in the absence of the IrOx
NPs, and, as such, the effect of adding increasing amounts of IrOx nanoparticles (NPs) to the film
was studied. It was found that, with increased IrOx content, the film became more O2-independent
and the signal was very reproducible between sensing films. However, the glucose signal (imax)
was very small in comparison to the signals achieved in Chapters 5 and 6 for the films fabricated
by the aliquot-deposition method.
The number of nitrophenyl groups deposited on the Au substrate was thus increased (to
increase the number of immobilized GOx molecules) by performing electrochemical deposition
for longer periods of time and using higher surface area substrates. It was found that there was an
optimal surface coverage for GOx immobilization. However, if the surface coverage became too
high, the glucose response became O2-dependent. Finally, it was found that, if a high surface area
carbon paper (CP) was used as the underlying substrate, a higher number of nitrophenyl groups
could be deposited. The GOx molecules immobilized to the carbon paper exhibited high enzyme
activity, giving signals more comparable to the signals observed in Chapters 5 and 6 (for aliquot-
deposited films containing substantially more GOx molecules). Despite these positive features,
however, these films still suffer from significant O2-dependency.
170
CHAPTER EIGHT: CONCLUSIONS AND FUTURE WORK
8.1 Conclusions
The global conclusions reached in this thesis work are presented below with the intent of
bringing the conclusions from all chapters together. The main goals of this thesis work were to
fabricate an IrOx-based glucose biosensor that accurately senses glucose concentrations,
independent of the partial pressure of the sample solution. The signal should be reproducible,
sensitive, and specific for glucose, i.e., not due to the reaction of electroactive interfering species.
To do this, the effect of the film components: water/ethanol, glucose oxidase (GOx), Ir oxide
nanoparticles (IrOx NPs) and Nafion® on film morphology, IrOx kinetics and glucose sensing
were studied.
In Chapters 5 and 6, the sensing films were fabricated by mixing an Ir NP sol-based ink
together with an aqueous solution of GOx and an ethanolic solution of Nafion®. The ink was then
deposited on a sputtered Au electrode and dried in air at room temperature. The Ir NPs were then
electrochemically oxidized to IrOx. In chapter 7, GOx was immobilized to the Au substrate via a
diazonium-derived film, and the ethanolic Ir ink was (assumed to be) intercalated within the film.
8.1.1 Conditions that affect the stability of Ir NP sol-based inks
The stabilization of an Ir NP sol-based ink is not only important for the fabrication of glucose
biosensors, but it is also important for other applications, such as for the fabrication of oxygen
evolution catalysts, reference electrodes, or for other medical applications.
Avoiding the addition of water to the ethanolic Ir NP ink (store in a sealed vessel) is
important for Ir NP stability. Films fabricated from water-containing inks dry at an non-
uniform rate, resulting in Ir NP aggregation. If the addition of water cannot be avoided, a
binding agent such as Nafion® should be added to the ink.
171
GOx adsorbs to the Ir NPs, causing them to aggregate, with time, causing them to
precipitate out of solution.
Ir inks show some thermoinstability, with optimal storage conditions being in a sealed
vessel at 4 °C, for no longer than 6 months.
8.1.2 Conditions that give the most reproducible glucose signals and film morphologies
Film morphology reproducibility, and the reproducibility of the sensing film response to
glucose, are among the most important parameters studied in this work. The highest degree of
reproducibility can be achieved when all of the GOx molecules are in contact with the well-
interconnected IrOx network (designated by a large charge in the Hupd and Ir(III)/Ir(IV) oxide
peaks). The conditions that most affect reproducibility involve the water, Nafion® and GOx content
of the glucose sensing ink, the dispersion of the Ir NPs in the ink, and the thickness of the sensing
film.
Ethanolic inks produce Ir NP films with uniform Ir distribution, whereas water-containing
inks produce films that have regions that dry at different rates, resulting in different degrees
of Ir NP aggregation. Water, however, is required in the ink for high GOx activity. When
GOx is incorporated into the water-containing inks, the non-uniformity of the resulting
film morphologies becomes worse, leading to irreproducibility between sensing film
morphologies. The optimal water content of the ink for the fabrication of a glucose sensing
film was found to be 20 vol.%.
The addition small amounts of Nafion® (1 wt. %) to the ink ensures enhanced Ir and Ir
oxide nanoparticle (IrOx NP) interconnectivity in the sensing film. This aids in improving
sensor performance and reproducibility. Too much Nafion®, however, blocks Ir and IrOx
NP interconnectivity, impeding the sensor.
172
Sonication of the Ir/Nafion® ink (for 2 hours) prior to film deposition ensures that the Ir
NPs and Nafion® polymer do not aggregate in the ink, thus resulting in a more uniformly
dispersed ink for film deposition.
Sensing films that are too thin (≤0.7 µm) likely result in the loss of some H2O2 (produced
during the O2-mediation route) to the bulk solution, thereby decreasing the reproducibility.
Thicker films (up to 4.0 µm, with uniform morphologies) are more likely to trap greater
amounts of H2O2, leading to larger, more accurate glucose signals, and are therefore more
desirable.
8.1.3 Conditions that give the overall highest glucose signals
The highest glucose signals achieved in an aerated solution are with poor interconnectivity
between GOx molecules and the IrOx network. In contrast, the highest glucose signals that
can be obtained from deaerated solutions are achieved only when all GOx molecules are
in contact with the interconnected IrOx matrix. The optimal GOx:Ir mass ratio (for aliquot
deposited films) of 1 g:1 g was found give the highest glucose signals in deaerated
envrionments.
Increasing the film thickness equates to an increased number of GOx molecules in the film,
thereby increasing the number of potential reaction sites for glucose, giving a rapid, large
glucose signal.
A small amount of Nafion® (1 wt. %) in the Ir ink ensures enhanced IrOx NP
interconncetivity in the film, enhancing direct electron transfer through the IrOx matrix, to
give a higher glucose signal.
173
Increasing the water-content of the ink leads to higher GOx activity. Films fabricated from
50 vol.% water-containing inks gave the highest glucose signals (but is not optimal as these
films are not reproducible, or O2-independent).
Diazonium salt-derived films used for the immobilization of GOx give relatively high
glucose signals for the amount of GOx that is present in the sensing film (in comparison to
aliquot-deposited films on Au), likely due to the prevention of GOx agglomeration.
The carbon paper substrate has a very high roughness in comparison to the sputtered Au
surface. It was found that by increasing the surface area available for nitrophenyl
deposition (via the electroreduction of nitrophenyl diazonium salts), more GOx molecules
can be immobilized, leading to higher glucose signals.
8.1.4 Conditions required for direct mediation and/or O2-independent sensing
Direct mediation requires good electrical connectivity between GOx and the interconnected
IrOx matrix. If the IrOx NPs are close enough to the redox active flavin adenine dinucleotide site
(buried ca. 1.3 nm158 inside the enzyme), electron tunnelling will occur.
In this thesis work (Chapters 5-6), O2-independent regeneration (characterized by identical
K’m values in aerated and deaerated solutions, as well as an imax of 1) could not be achieved.
However, direct mediation was clearly evident when sensing films were tested for their
response in deaerated environments. In comparison to the signals achieved via the O2-
mediated route, the direct-medation route exhibited smaller, but typically more
reproducible glucose signals.
The major preventative factor to achieving direct mediation is the inability to form a
reproducible, uniform film morphology. Aggregation of Ir nanoparticles is a result of non-
uniform film drying as well as adsorption by GOx molecules. Although the interactions
174
between GOx and the Ir NPs bring them into close contact, GOx likely blocks some of the
interconnectivity between the NPs in the process, as was evidenced by the decrease in the
charge in the Hupd peaks and Ir(III)/Ir(IV) redox peaks with increasing GOx content in
Chapter 6.
The immobilized (via the diazonium-derived film) GOx molecules exhibited O2-
independent glucose sensing. By immobilizing GOx it prevents enzyme aggregation, and
allows for the uniform distribution of the IrOx NPs around the GOx molecules.
To achieve direct mediation and O2-independent sensing, all of the conditions required for
reproducibility (Section 8.1.2) must be met.
8.1.5 Global comparison to other glucose biosensors
The IrOx-based electron transfer matrix, or “wiring”, that categorizes this sensor as being
fourth-generation, is a benefit over other sensing films, as the sensor does not depend on
the use of external mediators, such as ferrocene, ferricyanide, phenoxazine or quinones. By
avoiding the use of external mediators, this increases the accuracy of glucose sensing, as
issues such as mediator leakage or diffusion control do not affect the glucose signal. The
most well-known example of a wired glucose sensor is Heller’s long flexible
poly(vinylpyridine) or poly(vinylimidazole) polymer backbones decorated covalently
linked osmium-complexes which bind GOx 237,290. Other examples include the covalent
attachment of electron relays such as ferrocene chains291, Au NPs 292, and carbon
nanotubes293 to GOx.
The redox kinetics of the IrOx NP matrix in this work are reversible and surface-controlled.
Surface controlled transport kinetics are advantageous for glucose sensing, as there are no
175
rate determining steps in which to slow the glucose sensing process down, giving a rapid
and accurate glucose signal. Other sensors which rely on mobile mediators are susceptible
to diffusion controlled transport kinetics.
Ir NPs were chosen at the matrix as opposed to other metals such as Au and Pt due to the
stability of IrOx at anodic potentials. Ir can be converted into a thick, conductive, hydrous
IrOx layer that cannot be reduced back to metal. Pt and Au, however, form thin oxides that
slowly dissolve under aggressive conditions, resulting in the loss of the sensing matrix.
The sensing films are also capable of O2-independent glucose sensing, due to the faster rate
determining step in the direct mediation pathway, vs. the O2-mediation pathway, as
demonstrated by the “k3” concept discussed in Chapter 6.
The IrOx/Nafion®/GOx sensing film is scalable, especially since it does not require
diffusional mediators that require extra sensing components, such as membranes, to keep
the mediators close to the electrode surface.
The IrOx/Nafion®/GOx sensing film is biocompatible and can be used for in vivo glucose
sensing, whereas many glucose sensors are toxic (those that require diffusional mediators)
and should not be implanted, in case of leakage.
There are many different glucose biosensors in the literature and on the market today. The
main goals for glucose sensing are to fabricate a semi- to non-invasive sensor that is
capable of accurately and reliably determining blood or interstitial fluid glucose levels
continuously over a long period of time. The IrOx/Nafion®/GOx film has been proven to
be readily integrated into a semi-invasive “mosquito” glucose monitoring device8,
developed by Mintchev et al, at the University of Calgary.
176
8.2 Suggestions for Future Work
1. Electrochemical impedance spectroscopy (EIS) would be useful for the characterization of
the immobilized GOx/IrOx/Nafion® films, as well as the films fabricated by aliquot
deposition. This would help to uncover how resistive the diazonium-derived organic layer
was, as well as how resistive the addition of increasing amounts of Nafion® and/or GOx
was to the films, as was discussed in Chapters 4 and 5.
2. Based on Chapters 4 and 5, an investigation into alternative binding compounds to Nafion,
such as chitosan, or poly-vinyl alcohol (PVA), should also be carried out to determine if
these compounds will enhance the IrOx stability on the Au substrate, or the Ir
interconnectivity within the films further than that which was achieved using Nafion®.
3. In Chapter 5, XPS was performed on and IrOx/Nafion®/GOx film containing a mass ratio
of IrOx:GOx of 1. It would be beneficial to perform XPS on films of increasing IrOx:GOx
mass ratios to determine if there is more Ir retained in the films which contained higher
GOx contents. This would provide further evidence of GOx-Ir interactions.
4. In Chapters 5 and 6, when drying the Ir/Nafion®/GOx inks, it may be beneficial to dry them
underneath a stream of room temperature Ar to quicken the drying process. It is possible
that this may allow less time for the NPs to aggregate.
5. In Chapter 6, only the Ir sol and Nafion® film components were mixed together via
sonication prior to film fabrication. It would be beneficial for the fabrication of a uniform
film if a method in which GOx could be uniformly dispersed within the ink could be
developed, without denaturing the enzyme.
6. In Chapter 6, spin coating was found not to be ideal for depositing Ir/Nafion®/GOx sensing
films on Au. It may be beneficial to try different rotation protocols, and to deposit multiple
films, layered on top of each other to enhance film thickness, and fill in ‘gaps’ where the
177
IrOx NPs are not properly connected. It would also be of interest to use a technique such
as ellipsometry to determine film thickness, relative to the films formed via aliquot-
deposition.
7. In Chapter 7, immobilizing GOx to a high surface area carbon paper gave high GOx
activity, but poor O2-dependence and sensitivity. The high activity, however, is very
important and, as such, the process should be studied further. Transmission electron
microscopy imaging should also be performed to determine how the diazonium-derived
layer, the Ir NPs and the GOx molecules are deposited on the carbon paper substrate.
8. In Chapter 7, the quartz crystal microbalance technique could be used to confirm the
amount of GOx immobilized on a typical diazonium-derived film.
9. In Chapter 7, conducting atomic force microscopy could be used to determine where the
IrOx NP matrix could come into direct contact with the underlying Au substrate through
the pores in the diazonium-derived films. This would be beneficial, as it would be possible
to determine the degree of access of the Ir to the Au, and determine if more or less pores
were required for optimal electron transfer kinetics.
10. In chapter 7, thiol-based self-assembled monolayers (SAMs) should be compared to the
diazonium-salt derived films for the immobilization of GOx. As thiol-based SAMs form
reproducible, defect free monolayers that may enhance the reproducibility of the sensing
film morphologies, and hence the glucose signal.
11. The IrOx/Nafion®/GOx sensor should be tested for its long term stability in storage and in
blood-mimicking solutions, so as to determine if this sensing film is a candidate for
implantable glucose sensing devices.
178
REFERENCES
(1) Buttry, D. A. Applications of the Quartz Crystal Microbalance to
Electrochemistry; Electroanal. Chem., : New York, 1991; Vol. 17.
(2) Federation, I. D. 2012; Vol. 2014.
(3) Association, C. D. Canada, 2014; Vol. 2014.
(4) Van Bell, T. D.; Coppieters, K. T.; Von Herrath, M. G. Physiological Reviews.
2011, 9, 40.
(5) Alberti, K. G.; Zimmet, P. Z. Diabet. Med. 1998, 15, 15.
(6) Lindsay, R. S. Brisitsh Journal of Diabetes and Vascular Disease 2009, 9, 5.
(7) Clinic, M. 2013; Vol. 2014.
(8) Wang, G.; Mintchev, M. P. Engineering 2013, 5, 42.
(9) Campbell, H. B.; Elzanowska, H.; Birss, V. I. Biosens. Bioelectron. 2013, 42,
563.
(10) Jhas, A.; Elzanowska, H.; Sebastian, B.; Birss, V. Electrochim. Acta 2010, 55,
7683.
(11) Chambers, J. P.; Arulanandam, B. P.; Matta, L. L.; Weis, A.; Valdes, J. J. Curr.
Issues. Mol. Biol. 2008, 10, 12.
(12) Wilson, J. S. Sensor Technology Handbook; Elsevier: Burlington, MA, 2005; Vol.
1.
(13) Bioanalytic Sensors. The Biomedical Engineering Handbook; 2 ed.; Buck, R. P.,
Ed.; CRC Press LLC: Boca Raton, 2000.
(14) Analytical Applications of Piezoelectric Crystal Biosensors, Biosensor Principles
and Applications; Luong, J. H. T.; Guilbault, G. G., Eds.; Marcel Dekker: New York, 1991.
(15) Fan, X.; White, I. M.; Shopova, S. I.; Zhu, H.; Suter, J. D.; Sun, Y. Analytica
Chimica Acta 2008, 620, 19.
(16) Green, D. W.; Suns, H.; Plappll, B. J. Biol. Chem. 1993, 268, 7.
(17) Comstock, J.; mobihealthnews: 2014; Vol. 2014.
(18) Guo, X. S.; Liang, B.; Jian, J. M.; Zhang, Y. L.; Ye, X. S. Microchim Acta 2014,
181, 519.
(19) Kotzian, P.; Brazdilova, P.; Kalcher, K.; Vytras, K. Anal. Lett. 2005, 38, 15.
(20) Schmidtke, D. W.; Heller, A. Analytical Chemistry 1998, 70, 2149.
(21) Liao, Y.; Chou, T. Electroanalysis 2000, 12, 5.
(22) Tang, J.; Tang, D.; Niessner, R.; Knopp, D. Anal Bioanal Chem 2011, 400, 2041.
(23) Uludag, Y.; Tothill, I. E. Talanta 2010, 82, 6.
(24) Srivastava, M.; Srivastava, S. K.; Nirala, N. R.; Prakash, R. Analytical Methods
2014, 6, 817.
(25) Thennadil, S. N.; Rennert, J. L.; Wenzel, B. J.; Hazen, K. H.; Ruchti, T. L.;
Block, M. B. Diabetes technology and therapeutics. 2001, 3, 357.
(26) Rebrin, K.; Steil, G. M. Diabetes Technol. Ther. 2000, 2, 12.
(27) Dolan, B.; mobihealthnews: 2014; Vol. 2014.
(28) Beiderman, Y.; Blumenberg, R.; Rabani, N.; Teicher, M.; Garcia, J.; Mico, V.;
Zalevsky, Z. Biomedical Optics Express 2011 2.
(29) Lessin, J. E.; The Wall Street Journal: 2013; Vol. 2014.
(30) Reuters, T.; CBC news: 2014; Vol. 2014.
179
(31) Yao, H.; Liao, Y.; Lingley, A. R.; Afanasiev, A.; Lähdesmäki, I.; Otis, B. P.;
Parviz, B. A. Journal of Micromechanics and Microengineering 2012, 22, 1.
(32) Luong, J. H. T.; Male, K. B.; Glennon, J. D. Biotechnology Advances 2008, 26, 9.
(33) Biomedical, N.; Vol. 2014.
(34) Inc., B.; Bayer Inc. : Toronto, Ontario, 2014; Vol. 2014.
(35) Diagnostics, R.; Roche Diagnostics: 2014; Vol. 2014.
(36) LifeScan, I.; LifeScan, Inc: 2012; Vol. 2014.
(37) Dexcom; Dexcom: 2012; Vol. 2014.
(38) Vaddiraju, S.; Burgess, D. J.; Tomazos, I.; Jain, F. C.; Papadimitrakopoulos, F.
Journal of Diabetes Science and Technology 2010, 4, 1540.
(39) Gambardella, A. A.; Feldberg, S. W.; Murray, R. W. J. Am. Chem. Soc. 2012,
134, 5774.
(40) Vaddiraju, S.; Tomazos, I.; Burgess, D. J.; Jain, F. C.; Papadimitrakopoulos, F.
Biosens. Bioelectron. 2010, 25, 1553.
(41) Boyne, M. S.; Silver, D. M.; Kaplan, J.; Saudek, C. D. Diabetes. 2003, 52, 4.
(42) Heller, A.; Feldman, B. Chem. Rev. 2008, 7, 2482.
(43) Wilson, R.; Turner, A. P. F. biosens. Bioelectron. 1992, 7, 165.
(44) Leskovaca, V.; Trivić, S.; G. Wohlfahrt, G.; Kandrač, J.; Peričina, D. The
International Journal of Biochemistry & Cell Biology 2005, 37, 731.
(45) Zafar, M. N.; Beden, N.; Leech, D.; Sygmund, C.; Ludwig, R.; Gorton, L. Anal.
Bioanal. Chem. 2012, 402, 2069.
(46) Nanobioelectrochemistry; Luz, R. A. S.; Iost, R. M. C., F.N., Eds.; Springer-
Verlag: Berlin Heidelberg, 2013.
(47) Umar, A.; Rahman, M. M.; Vaseem, M.; Hahn, Y. B. Electrochem. Commun.
2009, 11, 118.
(48) Wang, J. X.; Sun, X. W.; Wei, A.; Lei, Y.; Cai, X. P.; Li, C. M.; Dong, Z. L.
Appl. Phys. Lett. 2006, 88, 233106.
(49) Li, J.; Lin, X. Biosens. Bioelectron. 2007, 22, 2878.
(50) Wan, D.; Yuan, S. M.; Li, G. L.; Neoh, K. G.; Kang, E. T. Appl. Mater. Interfaces
2010, 2, 3083.
(51) Harper, J. C.; Polsky, R.; Dirk, S. M.; Wheeler, D. R.; Brozik, S. M. Electroanal.
2007, 19, 1268.
(52) Radi, A.; Muñoz-Berbel, X.; Cortina-Puig, M.; Marty, J. Electroanalysis 2009,
21, 696.
(53) Shewchuk, D. M.; McDermott, M. T. Langmuir 2009, 25, 8.
(54) Delamar, M.; Hitmi, R.; Pinson, J.; Saveant, J. M. Journal of the American
Chemical Society 1992, 114, 5883.
(55) Adenier, A.; Combellas, C.; Kanoufi, F.; Pinson, J.; Podvorica, F. I. Chem. Mater.
2006, `8, 2021.
(56) Gui, A. L.; Liu, G.; Chockalingam, M.; Le Saux, G.; Luais, E.; Harper, J. B.;
Gooding, J. J. Electroanal. 2010, 22, 1824.
(57) Baranton, S.; Bélanger, D. The Journal of Physical Chemistry B 2005, 109,
24401.
(58) Jean Pinson, J.; Podvorica, F. Chem. Soc. Rev. 2005, 34, 429.
(59) Allongue, P.; Henry de Villeneuve, C.; Cherouvrier, G.; Cortès, R.; Bernard, M.
C. Journal of Electroanalytical Chemistry 2003, 550–551, 161.
180
(60) Chehimi, M. M.; Lamouri, A.; Picot, M.; Pinson, J. J. Mater. Chem. C 2014, 2,
356.
(61) Chambers, S. D.; McDermott, M. T.; Lucy, C. A. Analyst 2009, 134, 2273.
(62) Kariuki, J. K.; McDermott, M. T. Langmuir 2001, 17, 5947.
(63) Boukerma, K.; Chehimi, M. M.; Pinson, J.; Blomfield, C. Langmuir 2003, 19,
6333.
(64) Hurley, B. L.; McCreery, R. L. J. Electrochem. Soc. 2004, 151, B252.
(65) Laurentius, L.; Stoyanov, S. R.; Gusarov, S.; Kovalenko, A.; Du, R.; Lopinski, G.
P.; McDermott, M. T. ACS Nano 2011, 5, 4219.
(66) Laforgue, A.; Addou, T.; Belanger, D. Langmuir 2005, 21, 6855.
(67) Kullapere, M.; Kozlova, J.; Matisen, L.; Sammelselg, V.; Menezes, H. A.; Maia,
G.; Schiffrin, D. J.; Tammeveski, K. J. Electroanal. Chem 2010, 641, 90.
(68) Lehr, J. L.; Williamson, B. E.; Flavel, B. S.; Downard, A. J. Langmuir 2009, 25,
13503.
(69) Kibena, E.; Marandi, M.; Maeorg, U.; Venarusso, L. B.; Maia, G.; Matisen, L.;
Kasikov, A.; Sammerlselg, V.; Tammeveski, K. ChemPhysChem 2013, 14, 1043.
(70) Lehr, J.; Williamson, B. E.; Flavel, B. S.; Downard, A. J. Langmuir 2009, 25,
13503.
(71) Combellas, C.; Kanoufi, F.; Pinson, J.; Podvorica, F. I. J. Am. Chem. Soc. 2008,
130, 8576.
(72) Liu, G.; Iyengar, S. G.; Gooding, J. J. Electroanal. 2013, 25, 881.
(73) Radi, A.; Muñoz-Berbel, X.; Cortina-Puig, M.; Marty, J. Electroanalysis 2009,
21, 1624.
(74) Polsky, P.; Harper, J. C.; Dirk, S. M.; Arango, D. C.; Wheeler, D. R.; Brozki, S.
M. Langmuir 2007, 23, 364.
(75) WHITESIDES, G. M.; KRIEBEL, J. K.; LOVE, J. C. Sci. Prog. 2005, 88, 17.
(76) Allongue, P.; M. Delamar, M.; Desbat, B.; Fagebaume, O.; Hitmi, R.; Pinson, J.;
Saveant, J. M. J. Am. Chem. Soc. 1997, 119, 7.
(77) Cherevko, S.; Chung, C. Sens. Actuators, B 2009, 142, 8.
(78) Niu, X.; Lan, M.; Chen, C.; Zhao, H. Talanta 2012, 99, 6.
(79) Ci, S.; Huang, T.; Wen, Z.; Cui, S.; Mao, S.; Steeber, D. A.; Chen, J. Biosens.
Bioelectron. 2014, 54, 7.
(80) Hameed, R. M. A. Biosensors and Bioelectronics 2013, 47, 10.
(81) Toghill, K. E.; Compton, R. G. Int. J. Electrochem. Sci., 2010, 5, 56.
(82) Solanki, P. R.; Kaushik, A.; Agrawal, V. V.; Malhotra, B. D. NPG Asia Mater.
2011, 3, 8.
(83) da Silva, C. P.; Franzoi, A. C.; Fernandes, S. C.; Dupont, J.; Vieira, I. C. Enzyme
Microb Technol. 2013, 52, 6.
(84) Shen, J.; Dudik, L.; Liu, C. Sensors and Actuators B: Chemical 2007, 125, 8.
(85) Jiang, X., Massey University, 2009.
(86) Ren, X.; Meng, X.; Chen, D.; Tang, F.; Jiao, J. Biosensors and Bioelectronics
2005, 21, 5.
(87) Wu, B.; Hu, D.; Kuang, Y.; Yu, Y.; Zhang, X.; Chen, J. Chemical
Communications 2011, 47, 5253.
(88) Campbell, H. B.; H., E.; Birss, V. I. Biosensors and Bioelectronics 2013, 42, 7.
(89) Chen, A.; La Russa, D. J.; Miller, B. Langmuir 2004, 20, 8.
181
(90) Jhas, A.; Elzanowska, H.; Sebastian, B.; Birss, V. Electrochimica Acta 2010, 55,
7.
(91) Choi, H. N.; Kim, M. A.; Lee, W.-Y. Anal. Chim. Acta. 2005, 537, 179.
(92) Zhao, Z.; Lei, W.; Zhang, X.; Wang, B.; Jiang, H. Sensors 2010, 10, 1216.
(93) Ispas, C.; Njagi, J.; Cates, M.; Andreescu, S. J. Electrochem. Soc. 2008, 155,
F169.
(94) Liu, B.; Cao, Y.; Chen, D.; Kong, J.; Deng, J. Anal. Chim. Acta 2003, 478, 59.
(95) Rahman, M. M.; Ahammad, A. J. S.; Jin, J.-H.; Ahn, S. J.; Lee, J.-J. Sensors
2010, 10.
(96) Zang, J.; Li, C. M.; Cui, X.; Wang, J.; Sun, X.; Dong, H.; Sun, C. Q.
Electroanalysis 2007, 19, 1008.
(97) Tang, H.; Chen, J.; Yao, S.; Nie, L.; Deng, G.; Kuangb, Y. Anal. Biochem. 2004,
331, 89.
(98) Deng, C.; Li, M.; Xie, Q.; Liu, M.; Tan, Y.; Xu, X.; Yao, S. Anal. Chim. Acta
2006, 557, 85.
(99) Dai, M. Z.; Maxwell, S.; Vogt, B. D.; La Belle, J. T. Analytica Chimica Acta
2012, 738, 27.
(100) Wang, J.; Musameh, M. Analytical Chemistry 2003, 75, 2075.
(101) Courjean, O.; Gao, F.; Mano, N. Angew. Chem. Int. Ed. 2009, 48, 5897.
(102) Zafar, M. N.; Safina, G.; Ludwig, R.; Gorton, L. Analytical Biochemistry 2012,
425, 36.
(103) Hunt, L. B.; Lever, F. M. Platinum Metals Review 1969, 13, 13.
(104) Conway, B. E. J. Electrochem. Soc. 1991, 138, 1539.
(105) Lervik, I. A.; Tsypkin, M.; Owe, L.; Sunde, S. Journal of Electroanalytical
Chemistry 2010, 645, 8.
(106) Backholm, J.; Niklasson, G. A. Solar Energy Materials and Solar Cells 2008, 92,
1388.
(107) Y., T.; B., N.; A., K. Japanese Journal of Applied Physics 1987, 26, 1547.
(108) Weiland, J. D.; Anderson, D. J. IEEE Transactions on Biomedical Engineering
2000, 47, 8.
(109) Di Mario, C.; Grube, E.; Nisanci, Y.; Reifart, N.; Colombo, A.; Rodermann, J.;
Muller, R.; Umman, S.; Liistro, F.; Montorfano, M.; Alt, E. Int. J. Cardiol. 2004, 95, 329.
(110) Pickup, P. G.; Birss, V. I. J. Electroanal. Chem 1987, 220, 18.
(111) Pickup, P. G.; Birss, V. I. J. Electrochem. Soc. 1988, 135, 8.
(112) Birss, V. I.; Andreas, H.; Serebrennikova, I.; Elzanowska, H. Electrochem. Sol.
State Lett. 1999, 2, 4.
(113) Irhayem, E.; Elzanowska, E.; Jhas, A.; Skrzynecka, B.; Birss, V. Journal of
Electroanalytical Chemistry 2002, 538-539, 12.
(114) Elzanowska, E.; Abu-Irhayem, E.; Skrzynecka, B.; Birss, V. I. Electroanal. 2004,
16, 13.
(115) Pingarrón, J. M.; Yáñez-Sedeño, P.; González-Cortės, A. Echem. Acta. 2008, 53,
5848.
(116) Robblee, L. S.; Lefko, J. L.; Brummer, S. B. J. Electrochem. Soc. 2003, 130, 731.
(117) Bak, M.; Girvin, J. P.; Hambrecht, F., T,; Kufta, C. V.; Loeb, G. E.; Schmidt, E.
M. Med. Biol. Eng. Comput. 1990, 28, 257.
182
(118) Göbbels, K.; Kuenzel, T.; van Ooyen, A.; Baumgartner, W.; Schnakenberg, U.;
Bräunig, P. Biomaterials 2010, 31, 1055.
(119) Abu-Irhayem, E.; Elzanowska, E.; Jhas, A. S.; Skrzynecka, B.; Birss, V. I. J.
Electroanal. Chem 2002, 538-539, 153.
(120) Escalante-Garcia, I. L.; Duron-Torres, S. M.; Cruz, T. C.; Arriaga-Hurtado, L. G.
Journal of New Materials for Electrochemical Systems 2010, 13, 7.
(121) Nakagawa, T.; Beasley, C. A.; Murray, R. W. J. Phys. Chem. C 2009, 113, 12958.
(122) Rekstena, A.; Moradib, F.; Selanda, F.; Sundea, S. ECS Trans 2014, 58, 12.
(123) Sanchez Casalongue, H. G.; Ng, M. L.; Kaya, S.; Friebel, D.; Ogasawara, H.;
Nilsson, A. Angewandte Chemie International Edition 2014, n/a.
(124) Zhao, C.; E, Y.; Fan, L. Microchim Acta 2012, 178, 107.
(125) Smith, R. D. L.; Sporinova, B.; Fagan, R. D.; Trudel, S.; Berlinguette, C. P.
Chem. Mater. 2014, 26, 1654.
(126) Chang, C. H.; Yuen, T. S.; Nagao, Y.; Yugami, H. J. Power Sources 2010, 195 4.
(127) Fonseca, G. S.; Umpierre, A. P.; Fichtner, P. F. P.; Teixeira, S. R.; Dupont, J.
Chemistry – A European Journal 2003, 9, 3263.
(128) Hayek, K.; Goller, H.; Penner, S.; Rupprechter, G.; Zimmermann, C. Catalysis
Letters 2004, 92, 1.
(129) Motoyama, Y.; Taguchi, M.; Desmira, N.; Yoon, S.-H.; Mochida, I.; Nagashima,
H. Chemistry – An Asian Journal 2014, 9, 71.
(130) Gray, H. B. Nat Chem 2009, 1, 7.
(131) Kanan, M. W.; Nocera, D. G. Science 2008, 321, 1072.
(132) Zhang, Y. N.; Zhang, H. M.; Zhang, Y.; Ma, Y. W.; Zhong, H. X.; Ma, H. P.
Chemical Communications 2009, 3.
(133) Mayorga-Martinez, C. C.; Pino, F.; Kurbanoglu, S.; Rivas, L.; Ozkan, S. A.;
Merkoci, A. Journal of Materials Chemistry B 2014, 2, 7.
(134) Sau, T. K.; Rogach, A. L.; Jäckel, F.; Klar, T. A.; Feldmann, J. Advanced
Materials 2010, 22, 1805.
(135) Anderson, J. R.; Howe, R. F. Nature 1977, 268, 129.
(136) Baida, H.; Diao, P. Rare Met. 2012, 31, 523.
(137) Lee, W. H.; Kim, H. Catalysis Comm. 2011, 12.
(138) Zhao, C.; Yu, H.; Li, Y.; Li, X.; Ding, L.; Fan, L. Journal of Electroanalytical
Chemistry. 2013, 688.
(139) Weiland, J. D.; Anderson, D. J. IEEE Trans. Biomed. Eng. 2000, 47, 911.
(140) Meyer, R. D.; Cogan, S. F.; Nguyen, T. H.; Rauh, R. D. IEEE Trans. Neural
Systems Rehab. Eng.
2001, 9, 2.
(141) Cammilli, L.; Alcidi, L.; Papeschi, G.; Wiechmann, V.; Padeletti, L.; Grassi, G.
Pacing Clin. Electrophysiol. 1978, 1, 448.
(142) Niebauer, M. J.; Wilkoff, B.; Yamanouchi, Y.; Mazgalev, T.; Mowrey, K.; Tchou,
P. Circulation 1997, 96, 3732.
(143) Paasche, G.; Tasche, C.; Stöver, T.; Lesinski-Schiedat, A.; Lenarz, T. Otol.
Neurotol. 2009, 30, 592.
(144) Gierke, T. D.; Munn, G. E.; Wilson, F. C. J. Polym. Sci., Part B: Polym. Phys.
1981, 19, 1687.
(145) Li, J.; Wilmsmeyer, K. G.; Hou, H.; Madsen, L. A. Soft Matter 2009, 5, 7.
183
(146) Meredith, S.; Xu, S.; Meredith, M. T.; Minteer, S. D. J Vis Exp. 2012, 65.
(147) Harrison, D. J.; Turner, R. F. B.; Baltes, H. P. Analytical Chemistry 1988, 60,
2002.
(148) Klinge, U.; Klosterhalfen, B.; Ottinger, A. P.; Junge, K.; Schumpelick, V.
Biomaterials 2002, 23, 3487.
(149) Laroche, G.; Marois, Y.; Guidoin, R.; King, M. W.; Martin, L.; How, T.;
Douville, Y. Journal of Biomedical Materials Research 1995, 29, 1525.
(150) Mercado, R. C.; Moussy, F. Biosens. Bioelectron. 1998 13, 13.
(151) Zhang, Y.; Hu, Y.; Wilson, G. S.; Moatti-Sirat, D.; Poitout, V.; Reach, G.
Analytical Chemistry 1994, 66, 1183.
(152) T., T.; Yang, Y.; Chuang, M.; Lou, S.; Galik, M.; Flechsig, G.; Wang, J.
Electrochem commun. 2009, 11, 4.
(153) Fortier, G.; Vaillancourt, M.; Belanger, D. Electroanalysis 1992, 4, 9.
(154) Lee , Y. J.; Kim, J. D.; Park, J. Y. In 4th IEEE International Conference; IEEE:
Shenzhen, 2009, p 335.
(155) Moatti-Sirat, D.; Poitout, V.; Thomé, V.; M.N., G.; Zhang, Y.; Hu, Y.; Wilson, G.
S.; Lemonnier, F.; Klein, J. C.; Reach, G. Diabetologia 1994, 37, 7.
(156) Moore, C. M.; Akers, N. L.; Hill, A. D.; Johnson, Z. C.; Minteer, S. D.
Biomacromolecules 2004, 5, 7.
(157) Clark, L. C.; Lyons, C. Ann. N. Y. Acad. Sci. 1962, 102, 16.
(158) Hecht, H. J.; Kalisz, H. M.; Hendle, J.; Schmid, R. D.; Schomburg, D. J. Mol.
Biol. 1993, 229, 153.
(159) Andreas, H.; Elzanowska, E.; Serebrennikova, I.; Birss, V. I. J. Electrochem. Soc.
2000, 147, 4598.
(160) Abu-Irhayem, E.; Elzanowska, H.; birss, V. I. In 197th ECS Meeting Toronto,
Canada, 2000.
(161) Abu-Irhayem, E.; Amit, J.; Birss, V. J. Electroanal. Chem 2002, 538.539, 153.
(162) Jhas, A.; Elzanowska, H.; Sebastian, B.; Birss, V. Electrochim. Acta 2010, 55, 7.
(163) Abu-Irhayem, E.; Birss, V. I. In 201st ECS meeting Philadelphia, PA, 2002.
(164) Jhas, A. S., University of Calgary, 2007.
(165) Abu-Irhayem, E., University of Calgary 2004.
(166) Kim, B. K.; Kim, J. Y.; Kim, D.-H.; Choi, H. N.; Lee, W.-Y. Bull. Korean Chem.
Soc. 2013, 34, 1065.
(167) Berg, J. M.; Tymoczko, J. L.; Stryer, L. Biochemistry; 5 ed.; W.H. Freeman: New
York, NY, 2002.
(168) Atkins, G. L.; Nimmo, I. A. Biochemical Journal 1975, 149, 775.
(169) Chang, C. H.; Yuen, T. S.; Nagao, Y.; Yugami, H. J. Power Sources 2010, 195,
5938.
(170) Tang, D.; Li, Q.; Tang, J.; Su, B.; Chen, G. Analytica Chimica Acta 2011, 686, 6.
(171) Buchatip, S.; Ananthanawat, C.; Sithigorngul, P.; Sangvanich, P.; Rengpipat, S.;
Hoven, V. P. Sens. Actuators, B 2010, 145, 259.
(172) Chen, J. C.; Sadhasivam, S.; Lin, F. H. Process Biochem. 2011, 46, 543.
(173) Sauerbrey, G. Zeitschrift für Physik 1959, 155, 206.
(174) Bard, A. J.; Faulkner, L. R. Electrochemical methods, fundamentals and
applications; John Wiley & Sons, Inc.,
: New York, NY, 1980.
184
(175) Electron microscopy: principles and fundamentals; Amelinckx, S.; van Dyck, D.;
van Landuyt, J.; van Tendello, G., Eds.; VCH Verlagsgescllschaft mbH: Weinheim, Germany,
1997.
(176) Ohring, M. Materials science of thin films; Academic Press: Burlington: MA,
USA, 2001.
(177) Christie, A. B. In Chapter 5. X-Ray photoelectron spectroscopy; Walls, J. M., Ed.;
Cambridge University Press: New York, NY, 1989.
(178) Vij, D. R.; Lee, H. L.; Flynn, N. Handbook of applied solid state spectroscopy;
Springer USA, 2006.
(179) Atanasoskaa, L.; Guptaa, P.; Denga, C.; Warnera, R.; Larsona, S.; Thompson, J.
ECS Trans 2009, 16, 12.
(180) William, D.; Casllister, J. Materials Science and Engineering: An Introduction;
6th ed. ed.; John Wiley & Sons, Inc.: New York, NY., 2004.
(181) Wang, J.; Carlisle, J. A. Diamond Relat. Mater. 2006, 15, 279.
(182) Lyskawa, J.; Belanger, D. Chem. Mater. 2006, 18, 4755.
(183) Staros, J. V.; Wright, R. W.; Swingle, D. M. Anal. Biochem. 1986, 156, 220.
(184) Hermanson, G. T. Bioconjugate Techniques; 3rd. ed. ed.; Academic Press: San
Diega, CA., 2013.
(185) Jeong, H. a. J. K., J. Electrochim. Acta 2012, 80, 383.
(186) Store©, F. C.; Fuel Cell Store©: College Station, Texas, USA, 2014; Vol. 2014.
(187) Pickup, P. G.; Birss, V. I. J. Electroanal. Chem 1987, 220, 83.
(188) Angerstein-Kozlowska, H.; Conway, B. E.; Hamelin, A.; Stoicoviciu, I. J.
Electroanal. Chem 1987, 118, 429.
(189) Burke, L. D.; P.F. Nugent, P. F. Gold Bulletin 1997, 30, 43.
(190) Hernandez, A.; Ruiz, M. T. Bioinformatics 1998, 14, 2.
(191) 2006.
(192) 2008.
(193) Zhao, S.; Zhang, K.; Bai, Y.; Yang, W.; Sun, C. Bioelectrochemistry 2006, 69.
(194) Yang, M.; Yang, Y.; Liu, Y.; Shen, G.; Yu, R. Biosens. Bioelectron. 2006, 21.
(195) Salimi, A.; Sharifi, E.; Noorbakhsh, A.; Soltanian, S. Biosens. Bioelectron. 2007,
22.
(196) Yin, Y.; Lu, Y.; Wu, P.; Cai, C. Sensors 2005, 5.
(197) Rahman, M. A.; Kuma, P.; Park, D.; Shim, Y. Sensors 2008, 8.
(198) Andreas, H.; Elzanowska, H.; Serebrennikova, I.; Birss, V. I. J. Electrochem. Soc.
2000, 142, 7.
(199) Iwuoha, E. I.; Malcolm, M. R. Analytical Proceedings Including Analytical
Communications 1994, 31.
(200) Saini, S.; Hall, G. F.; Downs, M. E. A.; Turner, A. P. F. Anal. Chim. Acta 1991,
249, 15.
(201) Kapp, A.; Beissenhirtz, M. K.; Geyer, F.; Scheller, F.; Viezzoli, M. S.; Lisdat, F.
Electroanalysis 2006, 18, 1909.
(202) Schaefer, T. G., Massachuesetts institute of technology, 2009.
(203) Ilyas, S. U.; Pendyala, R.; Marneni, N. Applied Mechanics and Materials 2013,
372, 143.
(204) Zhang, Y. N.; Zhang, H. M.; Zhang, Y.; Ma, Y. W.; Zhong, H. X.; Ma, H. P.
Chem. Commun. 2009, 6589.
185
(205) Selvaraja, V.; Alagara, M.; Kumarb, K. S. Appl. Catal., B 2007, 75, 129.
(206) Pinho, S. P.; Macedo, E. A. J. Chem. Eng. Data. 2005, 50, 3.
(207) Colloidal Dispersions; Overbeek, J. T. G., Ed.; Royal Society of Chemistry:
London, 1981.
(208) El Sawy, E. N.; Birss, V. I. J. mater. Chem. 2009, 19, 8244.
(209) Koopal, C. G. J.; Bos, A. A. C. M.; Nolte, R. J. M. Sens. Actuators, B 1994, 18-
19, 166.
(210) Koopal, C. G. J.; Nolte, R. J. M. Bioelectrochem. Bioenerg. 1994, 33, 45.
(211) Augustynski, J.; Koudelka, M.; Sanchez, J.; Conway, B. E. J. Electroanal. Chem.
Interfacial Electrochem. 1984, 160, 233.
(212) Peuckert, M. Surf. Sci. 1984, 144, 451.
(213) Atanasoska, L. J.; Atanasoski, R.; Trasatti, S. Vacuum 1990, 40, 91.
(214) El Sawy, E. N.; Birss, V. I. J.Mater.Chem. 2009, 19, 8244.
(215) Li, M.; Wang, Y. B.; Zhang, X.; Li, Q. H.; Liu, Q.; Cheng, Y.; Zheng, Y. F.; Xi,
T. F.; Wei, S. C. Mater. Sci. Eng., C 2013, 33, 15.
(216) Nong, H. N.; Gan, L.; Willinger, E.; Teschner, D.; Strasser, P. Chem. Sci. 2014, 5,
2955.
(217) Kaduk, J.; Poly Crystallography, Inc.: Naperville IL 2014.
(218) Meyer, M.; Wohlfahrt, G.; Knäblein, J.; Schomburg, D. J. Comput. Aided Mater.
Des. 1998, 12, 425.
(219) Bradley, J. S.; Schmid, G.; Talapin, D. V.; Shevchenko, E. V.; Weller, H. In
Nanoparticles; Wiley-VCH Verlag GmbH & Co. KGaA: 2005, p 185.
(220) O'Hare, K. D.; Spedding, P. L. Chem. Eng. J. 1992, 48, 1.
(221) O'Hare, K. D.; Spedding, P. L.; Grimshaw, J. Developments in Chemical
Engineering and Mineral Processing 1993, 1, 118.
(222) Armstrong, F. A. Russ. J. Electrochem. 2002, 38, 58.
(223) Angerstein-Kozlowska, H.; Conway, B. E. J. Electroanal. Chem. Interfacial
Electrochem. 1979, 95, 1.
(224) Majumder, M.; Rendall, C. S.; Eukel, J. A.; Wang, J. Y. L.; Behabtu, N.; Pint, C.
L.; Liu, T.; Orbaek, A. W.; Mirri, F.; Nam, J.; Barron, A. R.; Hauge, R. H.; Schmidt, H. K.;
Pasquali, M. J. Phys. Chem. B 2012, 116, 6536.
(225) Karyakin, A. A.; Karyakina, E. E.; Gorton, L. Talanta 1996, 43 1597.
(226) Pasta, M.; La Mantia, F.; Cui, Y. Electrochim. Acta 2010, 55, 5561.
(227) Tominaga, M.; Shimazoe, T.; Nagashima, M.; Taniguchi, I. Electrochem.
Commun. 2005, 7, 189.
(228) Pickup, P. G.; Birss, V. I. J. Electrochem. Soc. 1988, 135, 126.
(229) Sethi, M.; Knecht, M. R. ACS Applied Materials & Interfaces 2009, 1, 1270.
(230) Tellechea, E.; Wilson, K. J.; Bravo, E.; Hamad-Schifferli, K. Langmuir 2012, 28,
5190.
(231) [Hong, G.; Heinz, H.; Naik, R. R.; Farmer, B. L.; Pachter, R. ACS Appl. Mater.
Interfaces 2009, 1, 388.
(232) Ren, X.; Meng, X.; Chen, D.; Tang, F.; Jiao, J. Biosens. Bioelectron. 2005, 21,
433.
(233) Pereira, A. R.; Iost, R. M.; Martins, M. V. A.; Yokomizo, C. H.; da Silva, W. C.;
Nantes, I. L.; Crespilho, F. N. PCCP 2011, 13, 12155.
186
(234) Graeupner, J.; Hintermair, U.; Huang, D. L.; Thomsen, J. M.; Takase, M.;
Campos, J.; Hashmi, S. M.; Elimelech, M.; Brudvig, G. W.; Crabtree, R. H. Organometallics
2013, 32, 5384.
(235) Gallaway, J. W.; Barton, S. A. C. J. Electroanal. Chem 2009, 626, 149.
(236) Degani, Y.; Heller, A. J. Phys. Chem. 1987, 91, 1285.
(237) Ohara, T. J.; Rajagopalan, R.; Heller, A. Anal. Chem. 1994, 665.
(238) Mano, N.; Yoo, J. E.; Tarver, J.; Loo, Y. L.; Heller, A. J. Am. Chem. Soc. 2007,
129, 7006.
(239) Wang, J. Chem. Rev. 2008, 108, 814.
(240) Sirotkin, A.; Zazybin, A. G.; Osipova, O. L.; Solomonov, B. N.; Faizullin, D. A.;
Fedotov, V. D. Moscow Univ. Chem. Bull. 2000, 41, 114.
(241) Mozhaev, V. V.; Khmelnitsky, Y. L.; Sergeeva, M. a. V.; Belova, A. B.;
Klyachko, N. L.; Levashov, A. V.; Martinek, K. Eur. J. Biochem. 1989, 184, 597.
(242) Griebenow, K.; Klibanov, A. M. J. Am. Chem. Soc. 1996, 118, 11695.
(243) Chin, J. T.; Wheeler, S. L.; Klibanov, A. M. Biotechnol. Bioeng. 1994, 44, 140.
(244) Griebenow, K.; Klibanov, A. M. Journal of the American Chemical Society 1996,
118, 11695.
(245) Iwuoha, E. I.; Smyth, M. R. Analyst 1994, 119.
(246) Karyakin, A. A.; Kotel'nikova, E. A.; Lukachova, L. V.; Karyakina, E. E.; Wang,
J. Analytical Chemistry 2002, 74, 1597.
(247) Vasileva, N.; Godjevargova, T. S. Mater. Sci. Eng., E, 25, 17.
(248) Shchukarev, S. A.; Tolmacheva, T. A. Zhurnal Strukturnoi Khimii 1968, 9, 21.
(249) Zhang, M.; Karra, S.; Gorski, W. Analytical Chemistry 2013, 85, 6026.
(250) Deng, Z. J.; Mortimer, G.; Schiller, T.; Musumeci, A.; Martin, D.; Minchin, R. F.
Nanotechnology 2009, 20, 1.
(251) Tenzer, S.; Docter, D.; Kuharev, J.; Musyanovych, A.; Fetz, V.; Hecht, R.;
Schlenk, F.; Fischer, D.; Kiouptsi, K.; Reinhardt, C.; Landfester, K.; Schild, H.; Maskos, M.;
Knauer, S. K.; Stauber, R. H. Nat. Nano 2013, 8, 772.
(252) Cass, A.; Davis, G.; Francis, E.; Hill, H. A.; Aston, W.; Higgins, J.; Plotkin, E.;
Scott, L.; Turner, A. P. Anal. Chem. 1984, 56, 667.
(253) Frew, J. E.; Hill, H. A. Anal. Chem. 1987, 59, 933A.
(254) Wang, J. Electroanal. 2001, 13, 983.
(255) Amine, A.; Kauffmann, J. M.; Patriarche, G. J.; Christian, G. D. Talanta 1993, 40,
1157.
(256) Foxx, D.; Kalu, E. E. Electrochem. Commun. 2007, 9, 584.
(257) Scriven, L. E. MRS Proceedings 1988, 121, 717.
(258) Foo, K. L.; Kashif1, M.; Hashim, U.; Ali, M. E. Current Nanoscience 2013, 9, 1.
(259) Li, X.; Feng, F.; Zhang, K.; Ye, S.; Kwok, D. Y.; Birss, V. Langmuir 2012, 28.
(260) Chaussé, A.; Chehimi, M. M.; Karsi, N.; Pinson, J.; Podvorica, P.; Vautrin-Ul, C.
Chem. Mater. 2001, 14, 392.
(261) Gutiérrez-Sánchez, C.; Pita, M.; Vaz-Dominguez, C.; Shleev, S.; De Lacey, A. L.
J. Am. Chem. Soc. 2012, 134, 17212.
(262) Radi, A.-E.; Lates, V.; Marty, J.-L. Electroanal. 2008, 20, 2557.
(263) Prieto-Simón, B.; Saint, C.; Voelcker, N. H. Anal. Chem. 2014, 86, 1422.
(264) Lehr, J.; Williamson, B. E.; Downard, A. J. J. Phys. Chem. 2011, 115, 6629.
(265) Smith, R. D. L.; Pickup, P. G. Electrochim. Acta 2009, 54, 2305.
187
(266) Toupin, M.; Belanger, D. Langmuir 2008, 24, 1910.
(267) Gooding, J. J. Electroanal. 2008, 20, 573.
(268) Abiman, P.; Wildgoose, G. G.; Compton, R. G. Int. J. Electrochem. Sci., 2008, 2,
104.
(269) March, G.; Reisberg, S.; Piro, B.; Pham, M. C.; Fave, C.; Noel, V. Anal. Chem.
2010, 82, 3523.
(270) Le Floch, F.; Bidan, G.; Pilan, L.; Ungureanu, E.; Simonato, J. Molecules crystals
and liquid crystals 2008, 486, 271.
(271) Benedetto, A.; Balog, M.; Viel, P.; Le Derf, F.; Salle, M.; Palacin, S. Electrochim.
Acta 2008, 53, 7117.
(272) Aryl Diazonium Salts: New Coupling Agents and Surface Science; Chehimi, M.
M., Ed.; Wiley-VCH Verlag & Co. KGaA: Weinheim, Germany.
(273) Kongsfelt, M.; Vinther, J.; Malmos, K.; Ceccato, M.; Torbensen, K.; Knudsen, C.
S.; Gothelf, K. V.; Pedersen, S. U.; Daasbjerg, K. J. Am. Chem. Soc. 2011, 133, 3788.
(274) Kullapere, M.; Marandi, M.; Matisen, L.; Mirkhalaf, F.; Carvalho, A.; Maia, G.;
Sammelselg, V.; Tammeveski, K. J. Solid State Electrochem. 2012, 16, 569.
(275) Du, W.; Wang, Q.; Saxner, D.; Deskins, N. A.; Su, D.; Krzanowski, J. E.;
Frenkel, A. I.; Teng, X. J. Am. Chem. Soc. 2011, 133, 15172.
(276) Shewchuk, D. M.; McDermott, M. T. Langmuir 2009, 25, 4556.
(277) Liu, G.; Paddon-Row, M. N.; Gooding, J. J. Electrochem. Commun. 2007, 9,
2218.
(278) Ganske, G.; Topalov, G.; Slavcheva, E.; Mokwa, W.; Schnakenberg, U.
Transducers 2009, 2106.
(279) Gerlache, M.; Sentruk, Z.; Quarin, G.; Kauffmann, J. Electroanalysis 1997, 9,
1088.
(280) Yan, X.; Ge, X.; Cui, S. Nanoscale Res Lett. 2011, 6, 313.
(281) Dyne, J.; Lin, Y.-S.; Lai, L. M. H.; Ginges, J. Z.; Luais, E.; Peterson, J. R.; Goon,
I. Y.; Amal, R.; Gooding, J. J. ChemPhysChem 2010, 11, 2807.
(282) Dharuman, V.; Chandrasekara Pillai, K. J. Solid State Electrochem. 2006, 10,
967.
(283) Xu, S.; Yao, Y.; Wang, P.; Yang, Y.; Xia, Y.; Liu, J.; Li, Z.; Huang, W. Int. J.
Electrochem. Sci., 2013, 8, 1863.
(284) Nishio, K.; Masuda, H. Angewandte Chemie International Edition 2011, 50,
1603.
(285) Allongue, P.; M. Delamar, M.; Desbat, B.; Fagebaume, O.; Hitmi, R.; Pinson, J.;
Saveant, J. M. J. Am. Chem. Soc. 1997, 119, 201.
(286) Wan, D.; Yuan, S. J.; Li, G. L.; Neoh, K. G.; Kang, E. T. Appl. Mater. Interfaces
2010 2, 9.
(287) Adenier, A.; Cabet-Deliry, E.; Chausse, A.; Griveau, S.; Mercier, F.; Pinson, J.;
Christine Vautrin-Ul, C. Chem. Mater. 2005, 17, 491.
(288) Michaelson, H. B. Journal of Applied Physics 1977, 48, 4729.
(289) Yadav, A. P. J. Nepal Chem. Soc. 2011, 28, 80.
(290) Pishko, M. V.; Katakis, I.; Lindquist, S.-E.; Ye, L.; Gregg, B. A.; Heller, A.
Angewandte Chemie. 1990, 29, 82.
(291) Badia, A.; Carlini, R.; Fernandez, A.; Battaglini, F.; Mikkelsen, S. R.; English, A.
M. J. Am. Chem. Soc. 1993, 115, 7053.
188
(292) Xiao, Y.; Patolsky, F.; Katz, E.; Hainfeld, J. F.; Willner, I. Science 2003, 299,
1877.
(293) Gooding, J. J.; Wibowo, R.; Liu, J.; Yang, W.; Losic, D.; Orbons, D.; Mearns, F.
J.; Shapter, J. G.; D.B., H. J. Am. Chem. Soc. 2003, 125, 9006.