University of Southern California Dissertations and Theses

Surface modification of nanomaterials and developments of nanobiosensors

(USC Thesis Other)

Surface modification of nanomaterials and developments of nanobiosensors

PDF

Download a page range

Download transcript

Copy asset link

Request this asset

Transcript (if available)

Content

SURFACE MODIFICATION OF NANOMATERIALS
AND DEVELOPMENT OF NANOBIOSENSORS

by
Marco Curreli

A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(CHEMISTRY)

May 2010

Copyright 2010 Marco Curreli

ii

EPIGRAPH

“There Is Always Something To Learn”

iii

DEDICATION

This Dissertation Is Dedicated To
My Father, Bruno Curreli

iv

ACKNOWLEDGMENTS
I would like to first thank my mentor and advisor, Professor Mark E. Thompson.
Professor Thompson guided me throughout my doctorate studies with unflagging support
and encouragement. Moreover, I had the special chance to actively participate in drafting
several NIH grant proposals and patent applications – a golden opportunity that only few
graduate students are lucky enough to have. My exposure to Professor Thompson’s
Research Group lab environment and the cutting edge technology in our field was
absolutely world class. A very special thank you indeed for the steady financial support
over the years.
My project collaborators, Professors Chongwu Zhou, Richard J. Cote, Ram Datar,
and Emil Kartlov, were of invaluable assistance and peerless company. Their excellent
collaborations and lively discussions broadened my scientific knowledge and expanded
my repertoire of techniques. I would also like to thank Professors Chongwu Zhou, Amy
Barros, Curt Witting, and Florian Mansfeld for overseeing my Screening and Qualifying
Exams; and Professors Richard Brutchey and Mansfeld for participating in my
Dissertation Committee.
Thanks to my fellow graduate students in the nanobiosensing team, particularly
Fumiaki Ishikawa, Chao Li, Rui Zhang, and HsiaoKang Chang. Your contribution has
been enormous, and I feel very privileged to also have earned your friendship. Thanks
v

also to my hardworking undergraduate students whom I had the chance to supervise,
Marie Cuevas and Kunal Patel.
I also would like to thank every member of my family, especially my father,
Bruno Curreli, to whom I dedicate this dissertation, and my grandfather Ferrucio Mosca
and my uncle Adriano Bocalon. Thank you for inspiring me and blessing me with so
many wonderful childhood memories.
A special thank you to former and current Thompson Research Group members
who have made my daily life at the lab always something to look forward to. In
particular, I have enjoyed spending time (and have learnt something in the meantime)
with Alberto Bossi, Arnold Tamayo, Azad Hassan, Biwu Ma, Carsten Borek, Cody
Schlenker, Dolores Perez, Elizabeth I. Mayo, James Ly, James Ly, Kenneth Hanson,
Peter Djurovich, Rui Zhang, Simona Garon, Slawa Diev. Thanks to the USC staff that
help me along the way, in particular: Judy Hom for her valuable and always cheerful
administrative help; the guys at the USC Machine Shop, Don, Mike, and Ramon; the
glass blowers, Jim and Phil; the computer consultants Jaime, Bruno, and Frank; the
remarkably handy and resourceful Ross Lewis.
Thanks to my undergraduate advisor Professor Yang Ba at Cal State LA and my
other college mentors including Professors Wayne Tikkanan, Matthias Selke, Amir,
Carlos Gutierrez, Cohen, Subramaniam, and Daneshvary. They inspired me to continue
my journey in the field beyond college and into graduate school.
vi

Thanks to my high school chemistry teachers in my native town of Cervignano
del Friuli, Udine, Italy, who initiated me into the field, witnessed my first exploits and
adventures in the laboratory, and inspired me to never stop pursuing it: Aligi Vidossi
Graziella Mocellin, Gabriella Dovier, Sebastianis, and Reginella. Also, Franco Spanghero
(Italian literature and history) and Rino Puissa (Mathematics). I recall with a guilty
pleasure getting in trouble with my high school partners-in-crime: Fabrizio Scussolin and
Dario Benedetto.
Finally, I would like to thank some friends who directly and indirectly have made
my achievements possible. In alphabetical order, many thanks to: Catherine Chang,
Chico, Chris B, Claudio Zorloni, Debbie Yang, Domenico Lauriola (Mimmo), Elaina
Redmond, Elio Pezzotta, Elizabeth Rosas, Fabio Lo Surdo, Franco De Dominicis,
Gatterina, Giuseppe Amendola, Gloria Banuelos, Gregg Vorbroker, Kathleen O’Neill,
Kris Fujita, Kyle, Laura Conklin, Lily Hwang, Luca Musacchio, Mari Hernandez, Mario
Sabatini, Maryam Azad, Michele Scarpino, Narciso Mones, Nhon Kim Huynh, Olga
Uchima, Paolo Puddu, Paul Antunas, Phil Singer, Roxana Lissa, Ryan Ozar, Sandy Au,
Signora Mara, Stephanie Matsumoto, Sweet P, Terry Kamaile, Tim Bachman, Vanessa
Lo Savio, Wayne J. Rizzi, Wendy Stork, and Yanka Burgos.
Old buddies in my home town in the North East of Italy, Cervignano. In
alphabetical order, many thanks to: Alessio Beuzer, Carlo Job, Christiano Giolo, Denis
Andratta, Francesco (Bruno) Pastore, Rudi Fabbro, and Simonino.
vii

TABLE OF CONTENTS

Epigraph ii

Dedication iii

Acknowledgments iv

List of Tables ix

List of Figures x

Abstract xxiv

Chapter 1. Introduction to Nanobiosensors 1
1.1 Background 1
1.2 Fundamental Principles of FET-Based Nanobiosensors 3
1.3 Carbon Nanotube- and Nanowire-based FET Sensors 6
1.4 Factors Affecting the Performance of Nanobiosensors 8
1.5 Conclusions 26
1.6 Chapter 1 References 27

Chapter 2. Surface Modification of Carbon Nanotubes 34
2.1 Introduction to Carbon Nanotubes 34
2.2 Coordination of RuCp* to the Carbon Nanotube Sidewalls 37
2.3 Recent Achievements on the Coordination of RuCp* to Nanotubes 63
2.4 Controlled Covalent Surface Modification 76
2.5 Selective Functionalization of Carbon Nanotubes 83
2.6 Conclusions 86
2.7 Chapter 2 References 89

Chapter 3. Surface Modification of Indium Oxide Nanowires 93
3.1 Indium Oxide Nanowires: Synthesis And Electronic Properties 93
3.2 Surface Modification 96
3.3 Selective Functionalization of In
2
O
3
Nanowire-Based FET Devices 105
3.4 Recent Achievements in the Selective Functionalization of In
2
O
3
120
3.5 Conclusions 123
3.6 Chapter 3 References 125

Chapter 4. Facilitating the Real Time Detection of Biomolecules 129
4.1 Improvements in the Work Process 129
4.2 Conclusions 145
4.3 Chapter 4 References 146
viii

Chapter 5. Real Time Detection of Biological Molecules 147
5.1 Introduction to Biomarkers 147
5.2 Detection of Prostate Specific Antigen (PSA) 150
5.3 Detection of the SARS’ Nucleocapsid Protein 160
5.4 Conclusions 178
5.5 Chapter 5 References 180

Chapter 6. Concluding Remarks 184
6.1 Summary and Concluding Remarks 184

Bibliography 192

ix

LIST OF TABLES

Table 2.1. Electronic absorption spectra for several Ru-based sandwich
compounds. Those compounds highlighted in the red box involve
complexes of RuCp* η
6
-coordinated to conjugated arenes such as
naphthalene, anthracene, pyrene, and chrysene. The counter ion of all
these complexes is PF
6
-
. .......................................................................................50
Table 5.1: Cancer biomarkers. Cancer biomarkers are listed for some common
cancers along with the biological source. PSA stands for prostate specific
antigen, CA for cancer antigen, and CEA for carcinoembryonic antigen. .......148
Table 5.2. Serum levels of epithelial ovarian cancer (EOC) biomarkers
determined by a multiplexed analysis in healthy controls and EOC patients
at every stage of tumor. Ranges represent one standard deviation of the
median value. This panel of EOC biomarkers includes cancer antigen 125
(CA-125), leptin, prolactin, osteopontin (OPN), insulin-like growth factor
II (IGF-II), and microphage migratory factor (MIF). ........................................149
x

LIST OF FIGURES
Figure 1.1. Structure of a field effect transistor (FET) nanobiosensor. (a) Cross
sectional view: source and drain electrodes bridge the semiconductor
channel. The gate electrode can be used to modulate the conductivity of
the semiconductor channel. A receptor molecule attached to the surface of
the semiconductor material can specifically recognize and capture a target
molecule from a buffer solution. (b)-(d) Top view, SEM images of typical
nano-FETs manufactured at our nanofabrication facility using aligned
carbon nanotubes (b), carbon nanotube networks (c), and indium oxide
nanowires (d). These nanomaterials are visible as “filament(s)” between S
and D electrodes. ....................................................................................................4
Figure 1.2. Mechanism to modulate the conductance of a p-type nanowire FET.
(a) A p-type nanowire functionalized with a receptor exhibits uniform
conductance prior to any interaction with the target molecule. (b) When a
negatively charged target molecule binds to the receptor, it attracts
positive hole carriers in the region of the wire below the receptor. This
attraction causes carriers to accumulate, resulting in an increase in
conductance. (c) On the other hand, when a positively charged target
molecule is captured, it depletes the carrier density in the nanowire,
causing a decrease in conductance. ........................................................................5
Figure 1.3. Cross-sectional dimensions of nanowires and nanotubes and their
importance in the sensitivity of the sensor device. Both the nanowire (a)
and the nanotube (b) have been configured with a similar linker molecule,
the same capture probe, and bound the same analyte while submerged in
the same buffer solution (identical Debye length). The charged bound
analyte exerts an electric field as deep as the Debye length, attracting or
repelling charge carries in the nanowire or nanotube. The small diameter
of the nanotube allows for the bound analyte influencing the charge
concentration on the entire cross-section of the nanotube, resulting a
strong signal generation. On the other hand, the large diameter of the
nanowire limits region of influence of the analyte to the proximity of the
surface, resulting weak accumulation/depletion of charge carries, and thus
in a weak signal generation. Binding of multiple analytes on the nanowire
surface can certainly yield stronger signals, whereas a single bound
analyte might be sufficient to observing the binding event for a nanotube.
Cartoons (c) and (d) show how a small protein like PSA (34 kDa) is
sufficient to cover the entire nanotube cross-section when it bounds to a
peptide amptamer capture probe with a lock-key mechanism. Covering the
entire cross-section might display buffer ions from the nanotube surface,
helping generating a signal. .................................................................................10
xi

Figure 1.4. Length of the channel device and device performance. Long channels
are desired to decrease the time required to capture an analyte since more
capture probes can be immobilized on a long channel, and thus it would be
easier to capture the analyte from solution. However, long channels are
more resistant. For this reason, the short channels are expected to yield
more sensitive devices since the sensitivity is inversely proportional to the
channel resistance. A compromise has to be reached with regards to the
best channel length. A common channel length is on the order of 2-10 µm. ........14
Figure 1.5. Influence of the bound analyte electric field on the charge carries of
the nanowire. ........................................................................................................16
Figure 1.6. Size of the three common capture probes on the surface on a nanowire
based FET. Classical antibodies are 10-15 nm “tall” and would place a
captured analyte far away from the nanowire (the “sensing” component of
the device). On the other hand, aptamers are much smaller and a capture
analyte will be closer to the nanowire, thus resulting in the production of a
stronger signal. .....................................................................................................17
Figure 1.7. Delivery systems. (a) Microfluidic channels can be used to precisely
deliver a small volume of analyte above the nanowire sensor. (b) View
along the flow direction. The flux inside the microfluidic channel is higher
in the central part of the channel. The analyte has then to diffuse
perpendicularly to the flux in order to reach the sensor surface. (c)
Alternatively, a mixing cell (or solution chamber) can be used for analyte
delivery. A mixing cell usually consists of a cone shaped device with a
narrow bottom aperture. A sensor is placed underneath in contact with the
analyte solution through this small window. The analyte diffuses to the
sensor surface unless (d) a system with injection and drain valves facilitate
the analyte transport. .............................................................................................19
Figure 1.8. Effect of the buffer electrolyte concentration on the sensitivity of
nanowire based FET sensors. This effect is illustrated for a p-type NW
device and a negatively charged analyte molecule. (a) At high electrolyte
concentration (1x PBS, short Debye length) most of the charge carried by
the captured analyte is screened by ions present in the buffer. This
screening causes the analyte charge to have little effect on the
accumulation phenomena that would provide an increase of the device
conductance. (b) Operating at a lower electrolyte concentration (0.01x
PBS) the charge carried by the analyte is poorly screened and thus, a
larger change in conductance can be observed. (c) In very dilute buffers,
charges located far away from the wire can still exert an influence on the
carrier density of the wire, resulting in extreme sensitivities. .............................23
xii

Figure 1.9. Configuration of an FET device operated with a liquid gate. In this
cartoon, a negative potential is applied between the source (ground) and
the liquid gate electrode. Cations in solution are attracted at the gate
electrode. Meanwhile, anions accumulate at the nanowire surface,
attracting positive carries from the p-type nanowire/nanotube. This
accumulation of carriers causes an increase in the channel conductivity.
An electrical double layer is formed at the channel/solution interface. .................25
Figure 2.1. Allotropic forms of carbon. Elemental carbon can be found in nature
under several allotropic forms, including amorphous, graphite, diamond,
carbon nanotubes, and fullerenes. .........................................................................35
Figure 2.2. Metallic and semiconducting carbon nanotubes. The chirality of the
nanotube determines whether the nanotube will be metallic (left) or
semiconducting (right). ........................................................................................36
Figure 2.3. Schematic representation of [Cp*Ru]
+
units coordinated to the surface
of a carbon nanotube. ...........................................................................................37
Figure 2.4. Some Cp*Ru-Arene complexes discussed in the literature. These
arenes are: (a) benzene, (b) naphthalene, (c) anthracene, (d) pyrene, and
(e) corannulene. .....................................................................................................38
Figure 2.5. Simulation of the displacement of pyrene by THF from the sandwich
compound Cp*Ru-pyrene. (a) The cation Cp*Ru-pyrene is represented
using two different graphics (space filling and ball-and-spoke). (b) The
first step toward the displacement of pyrene by THF molecules requires
tilting the Cp* ring, so a new coordination site is formed for the incoming
THF. Two more incoming THFs will eventually displace the pyrene from
the sandwich compound. ......................................................................................40
Figure 2.6. Monitoring the THF displacement of pyrene in Pyrene-RuCp* by
UV-Vis spectroscopy. (a) UV-Vis of a solution of pyrene-Ru-Cp* in
degassed CH
2
Cl
2
. After recording spectrum (a), THF was added into the
cuvette so the solution was 10% in THF v/v. (b) Another spectrum was
recorded 5 minutes later followed by nine more spectra within 475
minutes. These ten spectra basically overlap. (c) Another spectrum was
recorded 2 days later, and starts to show “free” pyrene. (d) 5 days later, the
spectrum of “free” pyrene was clearly distinguishable. ........................................40

xiii

Figure 2.7. Monitoring the MeCN displacement of pyrene in Pyrene-RuCp* by
UV-Vis spectroscopy. (a) UV-Vis of a solution of pyrene-Ru-Cp* in
degassed CH
2
Cl
2
. After recoring spectrum (a), MeCN was added into the
cuvett so the solution was 10% in MeCN v/v. (b) Another spectrum was
recording 3 minutes later followed by 13 more spectra within 465 minutes.
After 120 minutes, there was a sign of “free” pyrene. (c) Another
spectrum was recorded 2 days later, and shows “free” pyrene. The cuvett
used was capped with a Teflon cap and kept in the dark in between runs. ..........41
Figure 2.8. Simulation of the displacement of the nanotube from the Cp*Ru-
nanotube complex by THF. (a) The cation Cp*Ru-nanotube is represented
using two different graphics (space filling and ball-and-spoke). (b) The
first step toward the displacement of the nanotube by a THF molecule
would require tilting the Cp* ring. Unlike in the case of pyrene, the carbon
nanotube significantly prevents tilting, protecting the RuCp* from
incoming nucleophiles. ........................................................................................42
Figure 2.9. SEM pictures of the HiPCO nanotubes used in this experiment. .................43
Figure 2.10. Typical Raman spectra of “as received” carbon nanotubes. ......................44
Figure 2.11. Typical IR spectra of “as received” carbon nanotubes. ..............................45
Figure 2.12. Synthetic route 1. A suspension of carbon nanotubes in DCE is
refluxed in the presence of the precursor [Cp*Ru
(II)
-(MeCN)
3
][PF
6
]. .................46
Figure 2.13. Synthetic route 2. Carbon nanotubes are first reduced to the
corresponding Na salts using reduced naphthalene. The reduced nanotubes
are expected to reduce Ru(III) to Ru(II) and thus the Cp*Ru unit can
coordinate to the nanotubes. ................................................................................47
Figure 2.14. Possible product rearrangement following synthetic route 2. ....................47
Figure 2.15. Vibrational spectrum of [Cp*Ru-pyrene][PF
6
]. .........................................48
Figure 2.16. UV spectra of reagents and products involved in the synthesis
according to route 1. The spectra have been normalized or shifted for
clarity. (a)-Green: Suspension of nanotubes in THF. (b)-Red: Product(s) of
the reaction according to route 1--Cp*Ru-nanotube--suspended in THF.
(c)-Orange: Spectrum of Cp*Ru-Pyrene in DCE, the lambda max is
located at 312 and 355 nm. (d)-Blue: Spectrum of the precursor
Cp*Ru(MeCN)
3
in dry THF. ...............................................................................50
xiv

Figure 2.17. IR of [Cp*Ru(NCMe)
3
][PF
6
], route 1 starting material. The sample
was exposed to air for a few minutes during acquisition. The inset shows
the whole spectrum, in the range 4000-700 cm
-1
. ................................................51
Figure 2.18. IR spectrum of nanotube treated according to route 1. The expected
product is [Cp*Ru-nanotube][PF
6
]. .....................................................................52
Figure 2.19. IR spectrum of nanotube treated according to route 1 after the
mathematical subtraction of the IR spectrum of untreated nanotube. .................53
Figure 2.20. Raman spectra of the complex precursor [Cp*Ru(MeCN)
3
][PF
6
]. .............54
Figure 2.21. Raman spectra of route 1 product. The expected product is [Cp*Ru-
nanotube][PF
6
]. ....................................................................................................55
Figure 2.22. UV spectra of reagents and products involved in the synthesis
according to route 2. The spectra have been normalized or shifted for
clarity. (a) Nanotube suspension in THF. (b) The product (probably
Cp*Ru-nanotube) suspended in THF. (c) Absorption of a DCE solution of
Cp*Ru-Pyrene, the lambda max is located at 312 and 355 nm. (d)
Spectrum of the precursor Cp*Ru
(III)
-Cl
2
in dry THF. ........................................56
Figure 2.23. IR spectra of Cp*RuCl
2,
the Ru precursor according to route 2. ................57
Figure 2.24. IR spectrum of nanotube treated according to route 2 after the
mathematical subtraction of the IR spectrum of untreated nanotube. .................57
Figure 2.25. IR spectra of route 2 product compared to that of bare nanotube. .............58
Figure 2.26. Raman spectrum of the carbon nanotubes Na salt. .....................................59
Figure 2.27. Raman spectrum of the carbon nanotubes treated according to route
2 synthesis. ...........................................................................................................59
Figure 2.28. Effect of treating “as grown” nanotubes according to route 2
functionalization strategy. (Top) SEM picture of CVD grown nanotubes
on a Si wafer surface. (Bottom) SEM images of the same nanotubes after
chemical treatment according to route 2 synthesis. .............................................60
Figure 2.29. Proposed products of the carbon nanotubes treatments with Cp*-Ru
precursors according to (a) route 1 and (b) route 2. .............................................61
xv

Figure 2.30. Thermogravimetric analysis of the old batch of nanotubes used. The
TGA curve (left) shows a significant loss of mass below 400
o
C which is
associated with carbonaceous impurities. Also, the residual mass is
significantly high (~35%) which is associated to impurities related to
catalysts nanoparticles. The residual mass is shown in the inset as dark
gray mass. ..............................................................................................................64
Figure 2.31. TGA differential weight loss curves. The differential of the TGA
curve shown in figure 2.30 is shown on the left as the original set of data
points, and on the right after the application of a smoothing function to
increase clarity. ......................................................................................................65
Figure 2.32. TGA of as-purchased nanotubes. The loss of mass below 400 C
indicates the presence of amorphous carbon, impurities that may highly
interfere with my chemistry. The metal content is less than 20%. The
differential curve has been smoothed. ...................................................................66
Figure 2.33. TGA of purified nanotubes. There is no loss of mass below 400 C
indicating that traces of amorphous carbon have been removed. The metal
content is slightly above 20%, because amorphous carbon has been
eliminated thus increasing the % of metal content. The differential curve
has been smoothed for clarity. ...............................................................................67
Figure 2.34. The Raman spectra of as-received (red trace) and purified (blue
trace) Unidym nanotubes. Panel (a) shows the entire spectrum, panel (b)
focuses on the radial breathing mode region, panel (c) focuses on the D
and G lines, and panel (d) on the G’ lines. Both traces have been
normalized at the intensity of the G’ line. ..............................................................68
Figure 2.35. UV-Vis-nIR spectrum of suspended nanotubes in dry DCE. ......................69
Figure 2.36. Selected regions of the UV-Vis-nIR spectrum of nanotubes shown in
Figure 3.35. The left panel focuses on the 470-800 nm region where the
M11 and S22 absorptions take place. The right panel focuses on the
semiconducting fine structures (S11) in the 1050-1450 nm region. ......................70
Figure 2.37. UV-Vis spectroscopy product characterization. All spectra have
been recorded in the 260-1560 nm range using the same instrument, and
in the same solvent (dry DCE). (a, Red) Spectrum of the reaction product
Cp*-Ru-nanotube. (b, Green) Spectrum of suspended nanotubes. (c, Blue)
Spectrum of the reagent Cp*-Ru-(MeCN)
3
. (d, Wine) Spectrum of Cp*-
Ru-Pyrene. ............................................................................................................71
xvi

Figure 2.38. Detail of the UV-Vis spectra shown in Figure 2.37. (a, Red)
Spectrum of the reaction product Cp*-Ru-nanotube. This spectrum shows
a new broad absorption band centered at 466 nm. This band is absent is all
the other spectra. (b, Green) Spectrum of suspended nanotubes. (c, Blue)
Spectrum of the reagent Cp*-Ru-(MeCN)
3
. (d, Wine) Spectrum of Cp*-
Ru-Pyrene. ............................................................................................................72
Figure 2.39. The Raman spectra of purified Unidym nanotubes (blue trace) and
the Cp*-Ru modified nanotubes (black trace). Panel (a) shows the entire
spectrum, panel (b) focuses on the radial breathing mode region, panel (c)
focuses on the D and G line, and panel (d) on the G’ line. Both traces have
been normalized at the intensity of the G’ line in panels (a), (c), and (d),
and to the tallest RBM in panel (b). ......................................................................74
Figure 2.40. Selected region of Raman spectra of purified Unidym nanotubes
(blue trace) and the Cp*-Ru modified nanotubes (black trace) showing the
G’ line. ..................................................................................................................75
Figure 2.41. Oxidized carbons on the nanotube sidewall. Oxidized defect sites
can be obtained by oxygen plasma treatment. This method yields a number
of functional group, including carboxylic acids, aldehydes, ketones, and
alcohols. This oxidation breaks several C=C can create a hole in the
nanotube, affecting the electronic properties of the nanotube. ............................77
Figure 2.42. Electrical breakdown of metallic pathways nanotube-based FETs.
(a) Current (log scale) versus voltage for a nanotube FET (Vds = +500
mV). The On/Off ration is close to one, indicating the presence of metallic
pathways. (b) The same device exhibits an improvement in the On/Off
ratio after electrical breakdown. (c) Plot of drain current versus source-
drain voltage showing a typical curve for the electrical breakdown on
metallic metallic nanotubes (Vg = +30 V). ..........................................................78
Figure 2.43. Reaction mechanism for the addition of a nitrobenzene radical anion
to a carbon nanotube. This nitrobenzene radical is formed in situ from the
electrochemical reduction of a para-nitrobenzene diazonium salt. ......................79
Figure 2.44. Molecular modeling showing the addition of one (left) or two (right)
molecules of nitrobenzene radical anion to the C=C bond of a carbon
nanotube. The addition of nitrobenzene preserve the C-C sigma bond, and
only affect the C=C pi bond. ................................................................................80

xvii

Figure 2.45. Effect of the diazonium-modification on the conductivity of a
buckypaper. (a) An unmodified buckypaper is conductive and can be used
to observe the redox waves of [Fe(CN)
6
]
3-
. After the treatment with a
diazonium salt, the buckypaper becomes insulating, thus does not function
as working electrode, and the redox waves of [Fe(CN)
6
]
3-
cannot longer be
observed (flat CV trace). ......................................................................................82
Figure 2.46. The Bifunctional, electrochemically active molecule that can be
used to selectively functionalize carbon nanotube-based
electrodes/devices. ...............................................................................................84
Figure 2.47. Cyclic voltammetry traces a buckypaper (blue) and a buckypaper
fnctionalized with DMP-pyrene (red). The CV of the buckypaper is
essentially featureless as no redox active functional groups are present in
the nanotube paper. Upon binding DMP-pyrene, the CV trace shown the
typical redox waves of hydroquinone/paraquinone derivatives. ..........................85
Figure 2.48. Synthesis of 1-(4-(2,5-dimethoxyphenyl)butyl) pyrene. ...........................86
Figure 3.1. Schematic representation of the setup used for the synthesis of indium
oxide nanowire. The gray box and the red dots represent the furnace and
the heating coils, respectively. A piece of InAs is atomized by laser
ablation. Indium atoms are carried over the catalyst by the carrying gas,
where the VLS growth of indium oxide nanowires takes place. .........................94
Figure 3.2. Typical indium oxide nanowires grown in Professor Zhou’s
laboratory. (a) An individual In
2
O
3
nanowire. (b) A network of “as grown”
In
2
O
3
nanowires. ..................................................................................................95
Figure 3.3. Phosphonate-based bifunctional linker molecules as modifiers for the
surface of indium oxide nanowires. Freshly cleaned In
2
O
3
nanowires
possess numerous –OH groups on their surface (a), which can hydrolyze
phosphonate-based molecules (b) resulting in the immobilization of the
latter molecules on the nanowire surface (c). ......................................................96
Figure 3.4. Surface treatment to immobilize amine-rich biological molecules on
the surface of indium oxide nanowires. A phosphonate-based bifunctional
linker molecule is used to prepare the surface for bioconjugation. Terminal
COOH groups of the linker molecules are activated with EDC/NHS in
step (a) and then used to anchor biological molecules via amine groups
(step b). ................................................................................................................98
xviii

Figure 3.5. Fluorescence study using ITO substrates. Sample (a) was prepared
using a blue dye (AMCA-hydrazie, Pierce); control (b) was passivated
with ethanolamine prior to the reaction with the blue dye. Sample (c) was
prepared by immobilization of a single stranded DNA on the ITO and
hybriziation with the complementary single stranded DNA labeled with a
green dye; control (d) was obtained using a mismatched DNA (but still
labeled with the same green dye). Sample (e) was prepared by anchoring
biotin to the ITO and exposing it to streptavidin (labeled with a red dye);
control (f) was made by ethanolamine deactivation prior to biotin binding
and then exposed to the red-streptavidin. ............................................................99
Figure 3.6. Strategy for immobilization of an amino ferrocene to indium oxide
nanowires. ..........................................................................................................101
Figure 3.7. Cyclic voltammetry traces of a ferrocene derivative bound to ITO.
The sample (left panels) was prepared according to reaction steps outlined
in Figure 3.6. The control (right panels) however was not activated with
EDC/NHS, and retained its COOH surface. This carboxylic rich surface
might have formed an ammonium salt with the aminoferrocene. In fact,
the sample and control initially displayed a similar CV trace signal, but
washing with 10% acetic acid removed the non-covalently bound
ferrocene from the control. ................................................................................102
Figure 3.8. The gold nanoparticles used in this study are 25 nm in diameter, and
have streptavidin molecules bound to their surface. ..........................................103
Figure 3.9. Procedure to bind gold nanoparticle-labeled streptavidin to ITOs. (a)
EDC/NHS activation in MES buffer (pH ~4.5). (b) Exposure to a 0.1 mM
solution of amine terminated biotin, in PBS buffer, pH 7.4. (c) Exposure to
a solution of gold nanoparticle-labeled streptavidin in PBS buffer, pH 7.4. .......104
Figure 3.10. SEM images showing several examples of nanowires decorated with
gold nanoparticle-labeled streptavidin. ..............................................................104
Figure 3.11. Control samples of the experiments shown in Figure 3.10. Gold-
labeled streptavidin does not seem capable of binding to the passivated
nanowires . The large particles in some images are residual aggregates of
salts from the buffer. They are too big to be the 25 nm gold nanoparticles. .......105

xix

Figure 3.12. Technological goal and key technology for the selective
functionalization of dense arrays of FET devices. (a) The technological
goal is to functionalize a dense array of FET devices with different capture
probes (here represented by three different recognition groups on three
different devices). (b) The key technology for achieving the goal is to use
an electrochemical active molecule, such as the hydroquinone (Off state) /
benzoquinone (On state) redox pair. .................................................................107
Figure 3.13: Strategy for the selective functionalization of a dense array of FET
devices. This dense array is represented by four identical, parallel devices. ......108
Figure 3.14. Electrochemistry of an HQ-PA-coated ITO sheet. The chemical
structure of HQ-PA and Q-PA bound to ITO are shown on the left. The
graph on the right shows 15 consecutive CV traces demonstrating the
stability and reversibility of oxidation/ reduction of a SAM of HQ-PA on
an ITO glass sheet. The oxidation wave is centered at +330 mV; the
reduction wave at -200 mV. ...............................................................................110
Figure 3.15. Chronoamperometry traces. This electrochemical technique was
used to determine the amount of charge necessary to oxidize a predefined
area of HQ-PA: an average of 56.8 μC. When a thiol terminated DNA is
bond to Q-PA, according to the reaction scheme shown on the top right,
the charge necessary to oxidize all the unreacted HQ-PA in the same area
decreased to 50.95 μC. Interpretation of these results is that 10% of the Q-
PA have actually been reacted with the DNA. ..................................................111
Figure 3.16. (b) A fluorescence image of ITO surface that was oxidized to Q-PA,
reacted with SH-DNA, and the DNA paired to its complementary DNA
strand labeled with a fluorescence dye. (c) A fluorescence image of ITO
with the HQ-PA monolayer went through the same DNA attachment
procedure as the ITO sheet in (b), showing little or no DNA binding. ..............112
Figure 3.17. (a) A photograph of a nanowire mat sample contacted by two groups
of electrodes. Only the HQ-PA attached to the nanowires between the
upper electrodes were converted to Q-PA. (b) A SEM image of the In
2
O
3

nanowires before functionalization. The brighter stripes are gold
electrodes covering the nanowire mat. (c) The same sample imaged at
higher magnification, where the nanowire mat is clearly visible. (d) A
fluorescence image of the nanowires with Q-PA taken after DNA
attachment. The gold electrodes, passivated with an alkyl thiol, appear
dark under the fluorescence microscope. (e) A fluorescence image of the
nanowires with HQ-PA after DNA incubation. The nanowires appear
dark, indicating no DNA attached to HQ-PA. ...................................................114
xx

Figure 3.18. Overall synthetic approach to make HQ-PA. ..........................................116
Figure 3.19. (a) A planar indium oxide surface was functionalized with an Hq
derivative. Dissolved oxygen in the buffer oxidizes Hq to Pq. Hp then
binds to an amino terminated biotin. Biotin acts as a capture probe and
binds a dye-labeled streptavidin, resulting in a bright red surface under a
fluorescent light (box on the right). (b) The off state (DMP) is truly off.
An indium oxide planar surface coated with DMP does not get oxidized by
dissolved oxygen, does not bind biotin, and does not capture a dye-labeled
streptavidin. Thus it looks dark under fluorescence light (box on the right). ......121
Figure 3.20. Structure of DMP and electrochemical deprotection. DMP
derivatives can be converted to Pq on applying a working voltage ~1.0 V
(i) and Hp can then be used to immobilize biomolecules to the nanowire
surfaces (ii). ........................................................................................................122
Figure 4.1. Flow chart for the overall process leading to the real time detection of
biomolecules. The blue beveled boxes indicate the basic steps in the
process, and the green boxes indicate improvements/additions to a
particular operation. This process is illustrated for indium oxide
nanowires, but it would apply to carbon nanotube-based devices with little
modification. ......................................................................................................130
Figure 4.2. Initial set up for the synthesis of indium oxide nanowires. The
photographic image show a side view of the furnace. .......................................131
Figure 4.3. New setup for the synthesis of nanowires. The top photographic
images show the trace oxygen analyzer used to monitor the oxygen (top)
levels and the barometer (lower right corner). ...................................................132
Figure 4.4. The probe station and probe manipulators used to test FET devices. .........133
Figure 4.5. Sample holder. ............................................................................................135
Figure 4.6. Instruments and essential components for the automated survey of
devices. ...............................................................................................................135
Figure 4.7. Furnace. ......................................................................................................136

xxi

Figure 4.8. (a) Nanobiosensor chip has devices in center with electrial leads
running to the edges of the substrate. (b) The microfluidic chip is
constructed of PDMS elastomer, establishing a network of channels and
valves to direct fluid flow. (c) Our integrated platform combines the
microfluidic chip with the nanosensor array. .....................................................137
Figure 4.9. Details of the integrated platform. (a) Photo image of the 24-device
array of nanosensors. (b) Magnified photo image showing two sets of
three devices. (c) Further magnified photo image showing one device. (d)
Photo image of microfluidics integrated on top of the device shown in (a):
inset wholebody of the device with microfluidics. (e) Magnified photo
image of the devices with microfluidic system in place. (f) Further
magnified photo image showing a device, through the microfluidic chip. .........138
Figure 4.10. Attachment of the PDMS chip to an APTMS modified silicon oxide
surface. ...............................................................................................................139
Figure 4.11. Sequence of steps for the attachment of the PDMS chip to the sensor
array. ..................................................................................................................140
Figure 4.12. Delivery systems—microfluidic (a) and (b) and mixing cell (c)—
integrated with the sensor arrays, mounted on the PCB, and wired using
the pin connector card. .......................................................................................141
Figure 4.13. Experimental setup for monitoring the potential between two
electrodes immersed in solution while more buffer or protein solutions are
added to the mixing cell. ....................................................................................143
Figure 4.14. Monitoring the potential between noble metal wires and the
Ag/AgCl electrode over time after adding either buffer or streptavidin
(SA) solutions to the mixing cell. Notably, the baseline can fluctuate—10-
15%-- even if no other aliquots were added. ....................................................144
Figure 4.15. Monitoring the potential between two encapsulated Ag/AgCl
electrodes over time after adding either buffer or BSA solutions to the
mixing cell. The baseline fluctuations are short in duration--several
seconds—and often the change in potential is ~1% or less. .............................144

xxii

Figure 5.1. (a) Schematic diagram of the nanosensor. PSA-ABs are anchored to
the nanowire / nanotube surface and function as specific recognition
groups for PSA binding. (b) Reaction sequence for the modification of
In
2
O
3
nanowire: i, deposition of 3-phosphonopropionic acid; ii, DCC and
N-hydroxysuccinimide activation; iii, PSA-AB incubation (c) Reaction
sequence for the modification of nanotube: iv, deposition of 1-
pyrenebutanoic acid succinimidyl ester; v, PSA-AB incubation. ......................152
Figure 5.2. I-V and I-V
g
curves of an In
2
O
3
nanowire device (a, b) and a nanotube
device (c, d) before and after PSA incubation. Red and blue curves
represent measurements performed before and after PSA incubation,
respectively. .......................................................................................................154
Figure 5.3. Current recorded over time for an individual In
2
O
3
NW device (a) and
a nanotube mat device (b) when sequentially exposed buffer, BSA and
PSA. Insets: SEM images of respectively. .........................................................156
Figure 5.4. Solutions delivery system. We have used a Teflon cell to deliver
solutions of interest to the surface of the nanobiosensors. Devices placed
at the bottom of the cell are in contact with the solution via a micro-sized
aperture. Four screws guarantee sealing and prevent leaking. ...........................159
Figure 5.5. Current recorded over time for a blank nanotube mat device when
sequentially exposed to buffer and BSA. ...........................................................160
Figure 5.6. Ribbon structure of our engineered Fibronectin (Fn). The peptide
sequence after the Fn C terminus is “spelled out” to show the position of
our selective attachment point to the NW surface. ............................................163
Figure 5.7. Nanobiosensor devices based on indium oxide nanowires. (a)
Nanobiosensor chips as fabricated on a 3 inches silicon wafer. We
typically fabricate 9 chips with 2 different architectures per silicon wafer.
(b) One of the nanobiosensor chips. Distinguishable features include the
gold pads along the perimeter and an array of 24 sensors clustered at the
center of the chip. (c) One of the 24 nanobiosensor devices. The
interdigitated source and drain electrodes are clearly visible. (d) An SEM
image of the interdigitated source and drain interface showing several
indium oxide nanowires. (e) A single nanowire, the sensing element of our
sensors. ...............................................................................................................164

xxiii

Figure 5.8. (a) Schematic diagram showing Fn immobilized on the surface of an
In
2
O
3
nanowire FET device. The regions of Fn with the engineered
peptide sequence are highlighted in red. Fn was attached to the NWs via
the sulphydryl group of a cysteine near the C-terminus, remote from the
binding site. (b) A family of I
ds
-V
ds
curves and (c) a typical I
ds
-V
g
curve
(plotted both in linear (red) and logarithmic (blue)) obtained from one of
our devices operating with the liquid gate configuration. ..................................165
Figure 5.9. Normalized electrical output (I/Io) versus time of a single operating
device. (a)-(b) Response curves to passivation upon addition of successive
aliquots of BSA. Upon increasing the concentration of BSA (from pure
0.01x PBS), the baseline re-equilibrates at lower values of S-D current
until stability is ultimately reach at 40 µM BSA, in 0.01x PBS. (c)
Response for a nanowire device functionalized with Fn. The red arrows
indicate the times when the solution was raised to a given concentration of
N protein. Inset: configuration of our device during active sensing
measurements. BSA protein was used to block sites for non-specific
binding. The Fn probe molecule was then used to specifically capture the
target N protein. .................................................................................................168
Figure 5.10. Normalized response from three devices versus concentration of N
protein (dots). These plots can be fitted using a Langmuir isotherm model
(solid line). .........................................................................................................169
Figure 5.11. A control device without the Fn capture probe does not respond to
the presence of the SARS N protein. .................................................................170
Figure 5.12. Surface modification of our In
2
O
3
nanowire devices resulting in the
covalent immobilization of the Fibronectin probe on the nanowires. ...............174
Figure 5.13. Details and components are indicated in the figures. (A) Cross-
section schematic diagram of the setup used for real time N protein
sensing. This diagram is not on scale; the nanosensor device is shown
enlarged for clarity with respect to the rest of the set up. (B) A side view
of such setup. (C) A top-view photograph of the setup showing the Teflon
cell atop a circuit board. ....................................................................................175
Figure 5.14. Absolute response of the three nanowire device used to detect the N
protein. These are the same devices shown as relative response in Figure
3. .........................................................................................................................176
Figure 5.15. Establishing baseline in a protein-rich environment for the device
passivated with 2-mercaptoethanol prior to Fn. ...................................................177
xxiv

ABSTRACT

Semiconducting nanomaterials are strong candidates for sensing applications
because of their high surface to volume ratio. During my doctorate studies, I contributed
to the development of nanobiosensors, sensor devices based on field effect transistors that
utilize these nanomaterials as the sensing element in the device. I was particularly
involved in the surface modification of such nanomaterials.
Chapter One introduces nanobiosensors and discusses how certain factors related
to the device fabrication or experimental conditions influence the sensitivity, selectivity,
and settling time of sensor devices during the real time detection of biomolecules.
Chapter Two investigates the surface modification of carbon nanotubes. Several
methods were explored to address certain practical problems: a chemical method based
on the coordination of Cp*-Ru to the nanotube sidewalls resulted in the covalent
functionalization of nanotubes and introduction of dopants; and the electrochemical-
mediated addition of diazonium derived radicals. A related project involved investigating
an electrochemical method for the selective functionalization of nanotube-based FET
devices.
Chapter Three explores the surface modification of indium oxide nanowires.
Linker bifunctional molecules based on phosphonate derivatives were found to strongly
bind to the indium oxide surface. Moreover, a method for the electrochemical, selective
functionalization of In
2
O
3
based FETs was investigated.
xxv

Chapter Four details improvements to the overall process facilitating the real time
detection of biomolecules. These improvements include: growing nanowires under
controlled and reproducible conditions; testing devices in an automated fashion using a
testing station consisting of our own semiconductor parameter analyzer, switch matrix,
microfluidic accessories, among other components; and developing data analysis
software and a database.
Chapter Five explores the real time detection of biological molecules, specifically
the detection of biomarkers for two classes of malignancy: cancer and infectious diseases.
My research team was the first to demonstrate the detection of a cancer biomarker (PSA)
using FET nanobiosensors in a complementary fashion (using n-type and p-type
semiconductors). Our devices have also detected a biomarker related to the highly
contagious SARS’ coronavirus. Indium oxide nanowire devices were configured using
surface chemistry technique I developed.
Chapters Six provides concluding remarks.

1

CHAPTER 1:
INTRODUCTION TO NANOBIOSENSORS

Chapter Outline
1.1. Background
1.2. Fundamental Principles of FET Based Nanobiosensors
1.3. Carbon Nanotube- and Nanowire-based FET Sensors
1.4. Factors Affecting the Performance of Nanobiosensors
1.5. Conclusions
1.6. Chapter 1 References

1.1 Background
In the past 15 years, materials scientists and engineers progressively miniaturized
the materials that constitute the building blocks of various biomedical devices.
1-3
This
progressive downscaling has led to the creation of materials with at least one critical
dimension falling within the 1-100 nanometer range. There are two principal driving
forces that promote the exploration of matter in the nanometer range.
First, nanomaterials are comparable in size with most biological entities such as
proteins, nucleic acids, cells, viruses, etc., making them the ideal interface materials
between biological molecules and scientific instruments.
4-9

Second, this new class of materials possesses unique physical and chemical
properties directly arising from their size such as their high surface-to-volume ratio
2

(S/V). A direct consequence of this high S/V ratio is that a large fraction of the atoms in
the material are located at or near the surface. This proximity causes the surface atoms to
play an important role in determining the physical, chemical, and particularly electronic
properties of the nanomaterials. This dependence on the properties of the
nanomaterial/surrounding interface makes nanomaterials excellent substrates for
molecular sensing applications.
A diversity of sensor architectures have been designed and fabricated during the
last decade that utilize different nanomaterials as a sensing element (cantilevers, quantum
dots, nanotubes, nanowires, nanobelts, nanogaps, and nanoscale films).
10-24
Some of these
sensing devices, such as those based on cantilevers and quantum dots are highly specific,
ultra sensitive, and have short response times. However, these devices require integration
with optical components in order to translate surface binding phenomena into a readable
signal. The need for detection optics is expected to significantly increase the cost of
operation for such a device. In contrast, sensors designed to operate like field effect
transistors (FET) can directly translate the analyte-surface interaction into a readable
signal, without the need for elaborate optical components. These devices utilize the
electronic properties of the sensing element, such as its conductance, to produce the
signal output. Sensors based on FETs promise to revolutionize bioanalytical research by
offering the direct, real-time, highly specific, ultra sensitive, and label-free detection of
the desired biomolecule.
8, 25-27

My doctorate studies were devoted to the development of nanobiosensors based
on both indium oxide nanowires and carbon nanotubes.
3

1.2 Fundamental Principles of FET-Based Nanobiosensors
The typical structure of an FET sensor is illustrated in Figure 1.1(a). An FET
sensor has the arrangement of a common three electrode transistor: the semiconductor
material bridges the source and drain electrodes, forming the channel, and the gate
electrode is used to modulate the channel conductance. In the case of FET nanosensors,
the semiconductor channel is made of a nanomaterial and this nanomaterial is the
“sensing” component of the device. Semiconductor FET channels can be obtained using
several nanomaterials, including carbon nanotubes, silicon nanowires, and metal oxide
nanowires. S-D electrodes are often passivated with an inert, moisture blocking coating
layer (silicon nitride is widely used) that insulates the electrodes from the environment
(sample/solution under analysis). Figures 1.1 (b)-(d) show the top view of SEM images
of typical nano-FET devices prepared in our facilities. These examples include FET
devices manufactured using aligned carbon nanotubes (b), carbon nanotube networks (c),
and indium oxide nanowires (d). These nanomaterials are visible as “filament(s)”
between source (S) and drain (D) electrodes. In order to provide selectivity toward a
unique analyte, a specific recognition group (also called receptor, ligand, or probe) is
anchored to the surface of the semiconductor channel material. This receptor is typically
chosen to recognize its target molecule (also called analyte) with a high degree of both
specificity and affinity.

4

The sensing mechanism might be different depending whether sensor devices are
fabricated using nanowires or nanotubes.
11, 28
Signal generation for both silicon and
indium oxide nanowire-FET is produced by electrostatic interactions.
28, 29
The
mechanism of signal generation upon analyte binding is shown in Figure 1.2, using a p-
type nanowire as an example. For immunological-modified FET nanosensors, the
semiconductor channel has a uniform conductance determined by the main carrier density
in the nanowire (holes for a p-type semiconductor or electrons for an n-type). The
electrical current measured for the device at fixed source-drain and gate potentials is
constant over time, as illustrated schematically in Figure 1.2(a). Binding a charged
analyte molecule to a receptor anchored on the nanowire is equivalent to applying a

Semiconductor Channel / Nanowire
Receptor
Target
Buffer
Substrate
Dielectric
Source
Electrode
Drain
Electrode
Gate Electrode
Passivation
(a)
(S)
(d) (c) (b)
(D)
(S)
(S) (S)
(D)
(D) (D)
10 µm
3µm 3 µm

Figure 1.1. Structure of a field effect transistor (FET) nanobiosensor. (a) Cross sectional view: source
and drain electrodes bridge the semiconductor channel. The gate electrode can be used to modulate the
conductivity of the semiconductor channel. A receptor molecule attached to the surface of the
semiconductor material can specifically recognize and capture a target molecule from a buffer solution.
(b)-(d) Top view, SEM images of typical nano-FETs manufactured at our nanofabrication facility using
aligned carbon nanotubes (b), carbon nanotube networks (c), and indium oxide nanowires (d). These
nanomaterials are visible as “filament(s)” between S and D electrodes.
5

potential to the gate electrode, which alters the source-drain current. If the bound analyte
carries a charge opposite to the main carriers in the FET, charge carriers will accumulate
under the bound analyte, causing an increase in the device conductivity. This mechanism
is shown in Figure 1.2(b), where a negatively charged molecule such as DNA binds to the
p-type nanowire, causing a buildup of hole carriers, thus resulting in an increase in
conductivity. In contrast, analytes with molecular charges that are the same as the main
carriers in the FET lead to depletion of main carriers beneath the bound analyte, causing a
decrease in conductivity, as illustrated in 1.2(c).

Until recently, the mechanism leading to a signal transduction in a nanotube
biosensor has been poorly understood.
11, 30
For example, several studies have reported
that a nanotube device will show a decrease in resistance for every protein tested,

Time
Current
Carrier
depletion
Carrier
accumulation
(a)
(c)
Dielectric
p‐type nanowire
p‐type nanowire
p‐type nanowire
Time
Current
Current
Time
(b)
= positive charge
or hole carrier
= negative charge

Figure 1.2. Mechanism to modulate the conductance of a p-type nanowire FET. (a) A p-type nanowire
functionalized with a receptor exhibits uniform conductance prior to any interaction with the target
molecule. (b) When a negatively charged target molecule binds to the receptor, it attracts positive hole
carriers in the region of the wire below the receptor. This attraction causes carriers to accumulate,
resulting in an increase in conductance. (c) On the other hand, when a positively charged target
molecule is captured, it depletes the carrier density in the nanowire, causing a decrease in conductance.
6

independent of the overall charge on the protein.
30, 31
This observation conflicts with
those made on nanowire based transistors,
32, 33
and cannot be explained as an electrostatic
gating effect caused by the charged analyte perturbing the nanotube carriers. The key to
resolving this discrepancy was first proposed by Professor Dai’s group.
30
They suggested
that the dominant sensing mechanism is a modulation of the Schottky barrier at the
electrodes-nanotube interface caused by binding of the analyte and receptor. This
proposal has received support from other groups,
11, 34
whereas Professor Dekker’s group
has observed that another sensing mechanism, electrostatic gating, can also play an
important role depending on the nature of the analyte.
11

1.3 Carbon Nanotube- and Nanowire-Based FET Sensors
Carbon Nanotubes. Carbon nanotubes could be considered an ideal material for
sensing applications because every atom in a nanotube is located on the surface, leading
to extreme sensitivity to the surrounding environment.
35-38
Surface modified carbon
nanotubes are compatible (non-toxic) with living organisms such as cells,
39-43
thus
providing the appropriate interface between biological entities and electronic circuits.
Also, nanotubes can be readily synthesized from inexpensive precursors, such as
methane, or purchased from commercial sources in large quantities, with good purity.
Unfortunately, nanotubes suffer from important drawbacks in sensing
applications. These drawbacks include the need to separate semiconducting nanotubes
from metallic nanotubes, and a non uniform distribution of band gaps that leads to an
inability to fine tune the electronic properties. Long term, water-stable modification with
7

bio-receptor molecules has proved to be a challenge. Regardless, carbon nanotubes have
been shown to be able to detect proteins, oligonucleotides, and enzymatic activities with
high sensitivity and specificity.
36, 44

Nanowires. Semiconductor nanowire FET biosensors can be prepared with a
variety of semiconducting materials such as Si, Ge, or metal oxides (e.g. In
2
O
3
, SnO
2
,
ZnO, etc.), that are potentially useful in sensing applications and are currently the focus
of intense research.
45
Although larger in diameter than nanotubes, nanowires still possess
a large surface to volume ratio, resulting in good sensitivity to the surrounding
environment. The citotoxicity of nanowires is more difficult to evaluate compared to
nanotubes since important parameters such as material, size and shape, and surface
composition, can vary significantly, thus making it more difficult to study this class of
materials. However, a recent report shows neuron cell activity on silicon nanowires and
relative cellular signals recorded by silicon nanowire-based devices.
46
Nanowires can be
manufactured with high reproducibility, offer a well understood surface chemistry, and
provide the possibility of fine tuning the conductivity via introduction of dopants.
47

Drawbacks and advantages of nanowires depend on their method of production—
bottom up or top down--and are discussed below in Section 1.4.a-1.4.c. So far, nanowires
have been demonstrated to detect a variety of biological species such as oligonucleotides,
proteins, and viruses; moreover, they have been useful in monitoring cellular and
enzymatic activity, and determining the efficacy of potential drugs.
8, 25, 26, 46, 48-52

8

1.4. Factors Affecting the Performance of Nanobiosensors
Nanobiosensors with good performance should possess the following qualities,
especially if these devices are to be used for bioanalytical applications:
1) Outstanding selectivity or specificity
2) High sensitivity and reproducibility of results
3) Short settling time (time necessary to capture the analyte)
4) Fast recovering time (time to regenerate a device after a measurement)
These qualities are affected by several parameters related to fabrication
techniques, device geometries, and experimental conditions, discussed in the following
sections:
1.4.a. Nanowire and nanotube dimensions
1.4.b. Device geometries
1.4.c. Surface modification techniques
1.4.d. Delivery system
1.4.e. Active measurement parameters
1.4.f. Gating the device

1.4.a. Dimensions of Nanotubes and Nanowires
The size, geometry, and composition range for nanotubes are much more
restricted than that of nanowires. The former always have a circular cross-section, are
typically 1-2 nm in diameter and several microns in length, and their chemical
composition is difficult to alter (difficult to introduce doping atoms/elements). On the
9

other hand, nanowires (especially silicon nanowires) can be prepared with very different
cross-sections (circular, square, rectangular, trapezoidal), a variety of diameters (or
heights and widths) with lengths up to millimeters, and optimally tuned carrier densities
and mobilities. All these parameters can substantially affect the performance of the
sensor device.
For instance, cross section dimensions greatly affect the surface to volume ratio
and thus the sensitivity of the device, as illustrated in Figure 1.3. In this simulation, both
the nanowire (a) and the nanotube (b) have been configured with a similar linker
molecule, the same capture probe, and have been bound to the same analyte while
submerged in the same buffer solution (identical Debye length). The charged, bound
analyte exerts an electric field as deep as the buffer’s Debye length, attracting or repelling
charge carriers in the nanowire/nanotube.
53, 54
The small diameter of the nanotube allows
the bound analyte to influence the carrier concentration on the entire cross-section of the
nanotube, resulting in a strong signal generation. On the other hand, analytes’ region of
influence is curbed by the proximity of the surface, due to the large diameter of the
nanowire. This limitation on the analytes’ region of influence results in weak
accumulation/depletion of charge carriers and thus a weak signal generation for
nanowires. Binding of multiple analytes on the nanowire surface can certainly yield
stronger signals, whereas a single bound analyte might be sufficient to observing the
binding event for a nanotube. Figures 1.3(c) and (d) show how a small protein like PSA
(34 kDa) is sufficient to cover the entire nanotube cross-section when it binds to a peptide
aptamer capture probe with a lock-key mechanism. Covering the entire cross-section
10

might displace buffer ions from the nanotube surface. In this situation, the analyte is in
close contact with the entire section of the nanotube, resulting in an enhanced gating
effect and thus more strongly affecting the device conductivity.
Most of the studies elucidating the dependence of device characteristics on
nanomaterials’ dimensions and doping levels have been carried out on silicon nanowires.
For instance, Elfstrom et al. compared the electrical characteristics of Si nanowire based

Figure 1.3. Cross-sectional dimensions of nanowires and nanotubes and their importance in the
sensitivity of the sensor device. Both the nanowire (a) and the nanotube (b) have been configured with a
similar linker molecule, the same capture probe, and bound the same analyte while submerged in the
same buffer solution (identical Debye length). The charged bound analyte exerts an electric field as
deep as the Debye length, attracting or repelling charge carries in the nanowire or nanotube. The small
diameter of the nanotube allows for the bound analyte influencing the charge concentration on the
entire cross-section of the nanotube, resulting a strong signal generation. On the other hand, the large
diameter of the nanowire limits region of influence of the analyte to the proximity of the surface,
resulting weak accumulation/depletion of charge carries, and thus in a weak signal generation. Binding
of multiple analytes on the nanowire surface can certainly yield stronger signals, whereas a single
bound analyte might be sufficient to observing the binding event for a nanotube. Cartoons (c) and (d)
show how a small protein like PSA (34 kDa) is sufficient to cover the entire nanotube cross-section
when it bounds to a peptide amptamer capture probe with a lock-key mechanism. Covering the entire
cross-section might display buffer ions from the nanotube surface, helping generating a signal.
11

FETs upon exposure of the device to buffer solutions of different pH, as a function of the
Si nanowire diameter (50-170 nm).
55
They reported that environmental effects on the
nanowire FET properties decrease with increasing nanowire diameter, such that wires
larger than 150 nm behave similarly to those with micrometer-sized diameter. A similar
conclusion was reached by Stern et al. when devices fabricated using smaller nanowires
showed greater sensitivity to pH variation than larger nanowires.
32
The device sensitivity,
dependence on the channel dimensions, and doping concentration were also demonstrated
in other studies (see below).
56, 57
These studies underscore the importance of small
dimensions in order to achieve high levels of sensitivity to the environment and to the
effects brought about by analyte binding.
Several nanowire characteristics that can be altered during nanowire preparation,
such as nanowire dimension and doping levels, have been shown by Nair and Alam to
control device performances.
58-60
Their simulations indicate that devices fabricated using
small diameter nanowires (about 10 nm or less) would decrease the minimum amount of
analytes required for a detectable signal. Their work also indicates that for
nanobiosensors (nanowire sensors operating in high dielectric, aqueous buffers) the
doping density determines the device sensitivity, not the doping type (n- or p-type).
Lightly doped nanowires are expected to exhibit greater sensitivity than highly doped or
undoped devices, as experimentally demonstrated by Kim et al.
57
and noted by Cui et
al.
33
However, fabricating sensor devices with very small diameter nanowires and with
very low doping concentrations could introduce variability from device to device.
60

Dimensions and/or doping levels can be determined during the nanowire and
12

nanotube production. There are two general classes of techniques to produce either
nanotubes or nanowires: bottom up and top down approaches. While nanotubes are
always produced by bottom up methods, nanowires can be obtained by either method,
and each method has some advantages and disadvantages, as discussed below.
Nanotube and Nanowires by Bottom Up Technologies. The bottom up
approach involves preparing nanotubes or nanowires from molecular precursors, rather
than starting with the bulk semiconductor. Bottom up methods are used to produce both
group IV semiconductor (nanotubes and silicon) and metal oxide nanowires. “As grown”
nanotubes have essentially uniform diameter, lengths ranging from 3-5 microns to tenths
of microns, and a large distribution of chiralities. This variety of chiralities greatly affects
the electric properties of the nanotube FETs. Bottom-up techniques produce high-quality
nanowires, however, these nanowires grow with random orientation on the substrate and
are characterized by a distribution of lengths and diameters.
45
The variation in
nanotube/nanowire dimension, relative to top-down based devices, can impose limits on
bottom up sensors because of poor device uniformity and low fabrication yields.
61

Nanowires by Top-Down Technologies. Top down fabrication technologies start
with bulk materials and reduce the material dimensions using various techniques to cut,
pattern, etch, and shape these materials into the desired geometry and order.
45
Top down
production of nanowires employs a silicon on insulator (SOI) wafer as a substrate. These
top-down techniques can be either e-beam lithography, photolithography combined with
size reducing strategies such as the self-limiting oxidation, or a “nano-dimension”
transfer method such as the SNAP method.
51

13

Nanowires fabricated by top down techniques are uniform (nearly identical) and
well aligned. Top-down methods usually produce nanowires in high yields, in a
predetermined orientation and position on the substrate, making them easy to integrate
into functional devices. Disadvantages of top-down fabrication include high costs and a
slow rate of production. Also, nanowires produced by top-down technologies can rarely
approach the size of carbon nanotubes (1-2 nm). However, top down techniques offer an
excellent control over the nanowire length, which is crucial since the sensitivity is
inversely proportional to the length.
60

1.4.b. Device Fabrication: Geometry, Channel Width, and Channel Length
Once the nanotubes/nanowires have been prepared, the S-D electrodes and
passivation layer (optional) are deposited to complete the structure of the FET. The
silicon substrate can serve as the gate electrode. At this stage of the fabrication process
some important parameters that later will influence the device performance can be
defined, such as the channel length (distance between source and drain), channel width
(length of the source and drain), and the type of electrode passivation. These parameters
are shown in Figure 1.2 (a)-(b). The device dimensionality directly affects the response
time,
59
as illustrated in Figure 1.4. The kinetics of sensing, as a function of FET
dimensions, has been simulated theoretically by Sheehan and Whitman.
62
They found
that the channel length is the critical dimension for analyte accumulation on the nanowire
surface, since the sensitivity is a function of the total analyte flux over the sensing wire.
According to their simulations, fabricating FET devices with longer channel lengths will
14

significantly decrease the time required to produce a signal at a given concentration of
analyte.
62
A compromise has to be reached with regards to the best channel length. Long
channel lengths are desirable from a kinetic point of view but not from an electronic point
of view, where longer nanowires are known to be less sensitive.
60
A common channel
length is on the order of 2-10 µm.

1.4.c. Surface Functionalization Techniques, Linker Molecules and Receptors
While the nanotube/nanowire FETs described above will be sensitive to the
surrounding environment, they will not have the desired molecular recognition properties.
The surface of the sensing element (nanotube or nanowire) needs to be modified so that
the device can recognize a specific desired analyte. This selectivity is typically achieved
by anchoring a specific recognition group to the surface of the nanotube/nanowire. A
bifunctional linker molecule with two chemically different termini is used to help anchor
the receptor molecules to the nanotube/nanowire surface.
For carbon nanotubes, the unique structure of the nanotube (a network of sp
2

carbons) does not offer methods of covalent functionalization unless they drastically alter

Figure 1.4. Length of the channel device and device performance. Long channels are desired to
decrease the time required to capture an analyte since more capture probes can be immobilized on a
long channel, and thus it would be easier to capture the analyte from solution. However, long channels
are more resistant. For this reason, the short channels are expected to yield more sensitive devices since
the sensitivity is inversely proportional to the channel resistance. A compromise has to be reached with
regards to the best channel length. A common channel length is on the order of 2-10 µm.
15

the nanotube electronic properties. This concept is discussed in depth in Chapter Two.
Common surface modification methods for nanotubes involve the use of hydrophobically
bound linkers, such as pyrene derivatives or Tween 20. In the case of Si nanowires, the
linker molecule of choice depends on whether or not the wire has an oxide coating.
51
In
the case of metal oxide nanowires, a good choice for a linker molecule is one that is
terminated with a group capable of forming a non-hydrolizable conjugate, such as
siloxides or phosphonates. The optimum linker molecule was found to be a phosphonate
derivative, like 3-phosphonopropanoic acid.
63
This phosphonate spontaneously self
assembles on the nanowires from aqueous solutions or polar solvents.
63
The surface
modification of indium oxide nanowires is discussed in Chapter Three. After covalent
attachment of the receptor molecules, unreacted sites are usually deactivated with other
highly reactive molecules, such as ethanolamine, Tris buffer, or 2-mercaptoethanol.
The nature of the linker molecule can play a significant role in determining the
sensitivity of the resulting sensor device. For devices operating under the electrostatic
interaction mechanism, the size of the linker + probe molecule plays a crucial role in
determining the sensitivity of the biosensor. Short linkers/probes allow the bound analyte
to stay closer to the nanotube/nanowire where it can exert an electric field deeper into the
nanowire/nanotube, as shown in Figure 1.5.
After the nanowires/nanotubes are prepared with the linker molecules, capture
probes are bound to these linkers (often covalently) using standard bioconjugation
techniques. Examples of capture probes are provided in Chapter Five. The nature of the
capture probe has a critical role in two crucial parameters used to evaluate FET
16

nanobiosensors: the device selectivity and the sensitivity. The former is determined by
the binding affinity of the capture probe used toward the desired analyte, and is key to
eliminating false results. The latter arises from the size of the capture probe -- especially
when protein needs to be detected -- and is discussed in the next paragraph.
Importance of the Size of the Capture Probe. Monoclonal antibodies are
classical probe molecules for capturing target proteins. However, the relatively large size
of these antibodies (usually in the order of 10-15 nm, ~160 KDa) forces the detections of
proteins to be observed mainly under ideal conditions (in buffers with low ionic strength,
thus longer Debye length), as discussed later in Section 1.4.e. For this reason, alternative
capture probes have been investigated, such as aptamers and antibody mimetic proteins.
The relative size of these novel receptors to the size of conventional antibodies is
illustrated in a comparative cartoon in Figure 1.6.

Figure 1.5. Influence of the bound analyte electric field on the charge carries of the nanowire.
17

Aptamers are oligonucleotide-based affinity reagents that have recently been
employed in FET biosensors.
54
Aptamers solve this problem because of their tiny size (2-
3 nm, ~5 KDa). On the other hand, these aptamers are highly charged molecules due to
their phosphate backbone. Using highly charged probe molecules may be a disadvantage
since they could cause a decrease in sensitivity.
45

Antibody mimetic proteins (AMPs) are another class of capture probes, based on
relatively short peptide sequences. These AMPs can be engineered to improve
recognition properties such as selectivity and binding affinity,
64-68
with the potential to
surpass antibodies and nucleotide aptamers. These protein-based binding agents can be

Figure 1.6. Size of the three common capture probes on the surface on a nanowire based FET.
Classical antibodies are 10-15 nm “tall” and would place a captured analyte far away from the
nanowire (the “sensing” component of the device). On the other hand, aptamers are much smaller and a
capture analyte will be closer to the nanowire, thus resulting in the production of a stronger signal.
18

engineered and evolved until this high affinity for their target is achieved. The main
advantage of using these AMPs is their tiny size, an important factor to be considered
when designing nanobiosensors.
51
The use of AMPs leads to enhanced electrostatic
interaction between the captured molecules and the nanowire, resulting in stronger
signals. This enhanced signal due to AMPs is especially advantageous for performing
sensing measurements in media with high ionic strength, such as 1x PBS buffer or serum.
In these media, the Debye length (See Section 1.4.e) is significantly shorter than the size
of conventional antibodies.
53, 54
AMPs based on Fn are 3~5 nm in diameter whereas
antibodies are usually over 10-15 nm across, as shown in Figure 1.6.
There are many other advantages to using AMPs as receptors, including improved
binding affinity, higher selectivity, reduced cost, scalability, simplicity in the
manipulation of their structure, and their tiny size which allows for higher density of
probe molecules on the surface of the biosensor. In addition, AMPs can tolerate harsher
environmental changes such as high and low values of pH, salinity, and temperature.
Their robustness results in longer shelf life and higher tolerance of a wider variety of
chemical procedures. Moreover, it is expected that these peptide based affinity agents can
be produced in large quantity, at relatively low cost. The combination of low cost, high
binding affinity, chemical stability, and small size makes AMPs particularly attractive for
use with nanowire/nanotube biosensors.

19

1.4.d. Delivery System
Another important factor to be taken into consideration is the system used to
deliver the analyte solution, which mostly affect the settling time--the time required to
capture the analyte. The analyte must reach the active sensing surface in order to interact
with the capture agent. The time a receptor takes to capture its target molecule is affected
by the delivery strategy. Since fast responses are highly desirable, rapid analyte delivery
is crucial to the development of nanobiosensors. So far, two main systems have been
utilized for such a delivery: microfluidic channels and mixing cells,
51
each having its
advantages and disadvantages. These systems are shown in Figure 1.7.

Figure 1.7. Delivery systems. (a) Microfluidic channels can be used to precisely deliver a small
volume of analyte above the nanowire sensor. (b) View along the flow direction. The flux inside the
microfluidic channel is higher in the central part of the channel. The analyte has then to diffuse
perpendicularly to the flux in order to reach the sensor surface. (c) Alternatively, a mixing cell (or
solution chamber) can be used for analyte delivery. A mixing cell usually consists of a cone shaped
device with a narrow bottom aperture. A sensor is placed underneath in contact with the analyte
solution through this small window. The analyte diffuses to the sensor surface unless (d) a system with
injection and drain valves facilitate the analyte transport.
20

A microfluidic channel is usually made of molded elastomer such as PDMS with
injection and drain channels, as shown in Figure 1.7. Both microfluidic chips and mixing
cells used in my research tem are discussed in Chapter Four. The microfluidic devices are
placed on the top of the nanosensor so the solution can be directed over the nanowires. A
key benefit of a microfluidic device is that it allows the analysis to be conducted using
exceedingly small samples on the order of a nanoliter. The flow within the central part of
the channel is laminar and has a higher flux than at the periphery.
But microfluidic devices also come with several disadvantages. When a sample is
injected into the channel, in order for the analyte to reach the sensor surface, the analyte
has to diffuse normal to the flow, from the middle of the channel to the bottom where the
nanowires are located (Figure 1.7 (b)). Laminar microfluidic flow thus partially restricts
the ability of molecules to reach the nanosensor surface,
51, 62
especially for molecules
with high molecular weights ( above 100 kDa) that are known to diffuse an order of
magnitude slower than smaller biomolecules such as oligonucleotides.
62
Several
computational models of the sensing phenomena suggest that the analyte delivery to the
sensor surface might be the limiting step toward the detection of analytes at ultra-low
concentrations.
60, 62
Also, another disadvantage of PDMS channels is caused by the high
hydrophobic sidewalls present in those devices. Hydrophobic biomolecules with low
solubility in buffers are likely to adsorb and deposit along the PDMS walls. A passivation
strategy, using the proteins’ repelling properties of polyethylene glycol, was developed
by Wang et al. thus reducing undesirable, non-specific adsorption of biomolecules.
48

21

The other popular delivery method, shown schematically in Figure 1.7 (c), utilizes
a mixing cell (also called solution chamber). This cell, typically a cone-shaped, plastic
sample holder, is placed over the nanosensor chip and allows the solution to be delivered
from the top aperture. For simple cells where there is no continuous flow, different
solutions are delivered by replacement methods and the analyte diffuses isotropically
until it reaches the sensor surface. A more advanced mixing cell setup, shown in Figure
1.7(d), has been designed by Stern et al.
32
In this set up, injection of the solution
tangential to the nanowire-FET sensor significantly decreased the detection response
times compared to those observed in nanowire-FET that used microchannels for the
detection of similar target molecules.
61, 69

1.4.e. Active Measurement Conditions: pH and Ionic Strength
During sensing measurements, biological analytes need to be delivered to the
nanosensor surface. These analytes are usually dissolved in aqueous buffers, preferably at
a pH and electrolyte concentration similar to that of physiological solution. Phosphate
buffered saline (PBS) is an optimum model for human serum, which, like PBS, has a pH
value of 7.40 and 0.15 M electrolyte. The pH and especially the electrolyte concentration
are critical experimental variables to be considered.
Computational models for the response time of a nanobiosensor in a diffusion-
capture regime were executed by Nair and Alam in order to study the effects of
electrostatic screening caused by buffer solutions.
58
Their calculations predict that the
sensor response varies linearly with pH and logarithmically with electrolyte
22

concentration. Their simulation supports the data available in the literature, which indeed
show that nano-FET sensors respond linearly to pH variation (which is crucial for protein
detection) but non-linearly to electrolyte concentration. Thus, they suggest developing
analyte binding schemes at low ionic strength, in order to reduce the time taken to obtain
a detectable signal change. However, such a scheme might not meet certain minimum
electrolyte concentrations in order to retain a strong binding affinity between probes and
target molecules.
Debye Length. An important parameter that influences the device performance is
the Debye length ( λ
D
), defined as the maximum distance at which an external charge can
influence the nanowire carrier concentration.
60
The value of Debye length (in nm) in
water can be accurately calculated using the formula: λ
D
= 0.32 (I)
-1/2
where I is the ionic
strength of the buffer solution.
54
In aqueous media, the accumulation of carriers inside the
nanowire occurs over a depth equal to λ
D
, however, the Debye length decreases rapidly
with an increase in the ionic strength.
56
Probe molecules must therefore be attached as
close as possible to the nanowire surface, yet still retain their biological activity. The
Debye length plays an important role during immunological experiments where “large”,
relatively-low-charged biomolecules, such as antibodies, capture their target proteins.
When operating at the electrolyte concentration of serum, the Debye length (~ 0.7 nm) is
much smaller than the size of many antibodies (ca. 10-15 nm) and many proteins (ca. 5-
10 nm). Therefore, at such a short λ
D
, the electrolytes present in the buffer screen the
charges carried by the analyte, as shown in Figure 1.8 (a), resulting in a smaller nanowire
conductance change. Decreasing the salt concentration in the analyte solution allows for
23

detection of larger biomolecules. At longer Debye lengths, charged residues on the
analyte located several nanometers away from the nanowire will still exert an effect on
the charge carriers in the nanowire, as shown in Figure 1.8 (b-c).
One way to ensure longer Debye lengths is to use dilute buffer solutions with low
electrolyte concentrations. However, this practice could be problematic due to
complications caused by the necessary dilutions when preparing the sample. A second
problem with excessive dilutions is the fact that a minimum salt concentration is
necessary to retain biological activity of some proteins and is indispensable for DNA
hybridization.
56

1 x PBS
λ
D
~ 0.7 nm
p‐type nanowire
(a)
p‐type nanowire
(b)
p‐type nanowire
(c)
10
‐2
x PBS
λ
D
~ 7.3 nm
10
‐4
x PBS
λ
D
~ 75 nm
Time
Current
Time
Current
Time
Current
= negative charge
on the biomolecule
= electrolytes
in the buffer
= positive
hole carrier

Figure 1.8. Effect of the buffer electrolyte concentration on the sensitivity of nanowire based FET
sensors. This effect is illustrated for a p-type NW device and a negatively charged analyte molecule. (a)
At high electrolyte concentration (1x PBS, short Debye length) most of the charge carried by the
captured analyte is screened by ions present in the buffer. This screening causes the analyte charge to
have little effect on the accumulation phenomena that would provide an increase of the device
conductance. (b) Operating at a lower electrolyte concentration (0.01x PBS) the charge carried by the
analyte is poorly screened and thus, a larger change in conductance can be observed. (c) In very dilute
buffers, charges located far away from the wire can still exert an influence on the carrier density of the
wire, resulting in extreme sensitivities.
24

A recent experiment designed by Stern et al. clearly demonstrates that the
electrolyte concentration of buffers is a critical variable influencing the sensitivity of
these nanobiosensors.
53

The relationship between the spatial location of charge and chemical gating
effects was also investigated by Zhang et al.
70
Their results confirm that the detection
sensitivity of nanowire devices strongly depends on the location and strength of the
electric field produced by analyte molecules on the nanowire surface.
This sensitivity dependence on the buffer composition is an important limitation
for future applications of nanobiosensors when fast detection is required. Other
immunological assays based on optical detection, such as ELISA, can comfortably
operate at serum’s electrolyte concentrations.

1.4.f. Gating the Device
The device sensitivity of FET nanobiosensors can be tuned to its optimal value
using the gate electrode, for a given fixed S-D voltage. Applying a gate voltage close to
the inflection point of the I-V
g
curve, where the device transconductance is at its
maximum, allows working at high sensitivity. There are two types of gate electrodes that
can be used: the silicon substrate and liquid gate. The former uses heavily doped silicon
from the substrate as an electrode and its coating layer of silicon oxide as the dielectric
material. The latter uses noble metal wires (Pt or Au) or an encapsulated Ag, AgCl-
coated wire as the electrode (Ag/AgCl) and the buffer solution as the dielectric material.
25

This electrode arrangement is known in the field as “liquid gate” and is shown in Figure
1.9. The optimum type of electrode—metal wires or Ag/AgCl—is discussed in Section
4.1.
When an FET device is operated with the liquid gate configuration a voltage is
applied between the source electrode (ground) and liquid gate electrode. When a negative
voltage is applied cations accumulate at the liquid gate surface, while anions accumulate
on the channel surface. These anions will cause an accumulation of carriers (holes) in the
channel for a p-type semiconductor, or will cause a depletion of carries (electrons) for an
n-type semiconductor.
71
This situation is illustrated in Figure 1.9 for a p-type
semiconductor. Of course, if a positive voltage is applied, the reverse scenario occurs.
Rosenblatt et. al. showed that the gate electrode via liquid gate was about 20 times more
effective at modulating the current in the nanotube than the back gate.

= Hole carrier
= Anion
p‐type nanowire or nanotube
Silicon oxide
Source Drain
= Cation
Buffer
Electrical
double layer
Liquid
Gate
V
g

V
ds

Figure 1.9. Configuration of an FET device operated with a liquid gate. In this cartoon, a negative
potential is applied between the source (ground) and the liquid gate electrode. Cations in solution are
attracted at the gate electrode. Meanwhile, anions accumulate at the nanowire surface, attracting
positive carries from the p-type nanowire/nanotube. This accumulation of carriers causes an increase in
the channel conductivity. An electrical double layer is formed at the channel/solution interface.
26

1.5 Conclusions
This field of nanobiosensors is expanding rapidly and has already demonstrated
several potential applications including health monitoring and drug discovery. In this
chapter, I have introduced nanobiosensors and discussed several parameters influencing
the sensing curves in real time detection experiments, such as sensitivity, selectivity, and
settling time. The sensitivity is mainly affected by nanowire and nanotube dimensions,
doping levels, device geometry, gating method (back gate or liquid gate) ionic strength of
the buffer, size of the capture probe, and applied gate voltage affect. The selectivity of
these devices is directly related from the binding affinity of the capture probe for the
analyte. The settling time, the time it takes to capture the analyte and produce a binding
signal, is mainly affected by the type of delivery system used (microfluidic or mixing
cell).
I devoted my doctorate studies to the development of nanobiosensors along with
my colleagues Chao Li, Fumiaki Ishikawa, Rui Zhang, and HsiaoKang Chang. This team
was quite unique. In fact, we are the only research team that carries out parallel work on
two types of sensing platforms, and can clearly identify the advantages and disadvantages
of each platform. Although the nanobiosensing project is a team project, where the
success depends on each team member, I have provided some unique ideas and
contributions to this field of science. The next four chapters summarize our efforts in
developing nanobiosensors.
27

1.6 Chapter 1 References
1. Holleman, A.; Cheok, M. H.; den Boer, M. L.; Yang, W. J.; Veerman, A. J. P.;
Kazemier, K. M.; Pei, D. Q.; Cheng, C.; Pui, C. H.; Relling, M. V.; Janka-Schaub,
G. E.; Pieters, R.; Evans, W. E., Gene-expression patterns in drug-resistant acute
lymphoblastic leukemia cells and response to treatment. New England Journal of
Medicine 2004, 351, (6), 533-542.
2. Ferrari, M., Cancer nanotechnology: Opportunities and challenges. Nature
Reviews Cancer 2005, 5, (3), 161-171.
3. Heath, J. R.; Davis, M. E., Nanotechnology and cancer. Annual Review of
Medicine 2008, 59, 251-265.
4. Englebienne, P.; Van Hoonacker, A., Bionanotechnology: The science of
revealing life with nanostructures. Current Nanoscience 2005, 1, (2), 97-106.
5. Bogunia-Kubik, K.; Sugisaka, M., From molecular biology to nanotechnology
and nanomedicine. Biosystems 2002, 65, (2-3), 123-138.
6. Tamerler, C.; Sarikaya, M., Molecular biomimetics: nanotechnology and
bionanotechnology using genetically engineered peptides. Philosophical
Transactions of the Royal Society a-Mathematical Physical and Engineering
Sciences 2009, 367, (1894), 1705-1726.
7. Gao, J. H.; Xu, B., Applications of nanomaterials inside cells. Nano Today 2009,
4, (1), 37-51.
8. Patolsky, F.; Zheng, G.; Lieber, C. M., Nanowire sensors for medicine and the life
sciences. Nanomedicine 2006, 1, (1), 51-65.
9. Yang, W. R.; Thordarson, P.; Gooding, J. J.; Ringer, S. P.; Braet, F., Carbon
nanotubes for biological and biomedical applications. Nanotechnology 2007, 18,
(41), 412001-412013.
10. Raymo, F. M.; Yildiz, I., Luminescent chemosensors based on semiconductor
quantum dots. Physical Chemistry Chemical Physics 2007, 9, (17), 2036-2043.
11. Heller, I.; Janssens, A. M.; Mannik, J.; Minot, E. D.; Lemay, S. G.; Dekker, C.,
Identifying the mechanism of biosensing with carbon nanotube transistors. Nano
Letters 2008, 8, (2), 591-595.
28

12. Tansil, N. C.; Gao, Z. Q., Nanoparticles in biomolecular detection. Nano Today
2006, 1, (1), 28-37.
13. Rosi, N. L.; Mirkin, C. A., Nanostructures in biodiagnostics. Chemical Reviews
2005, 105, (4), 1547-1562.
14. Liang, X. G.; Chou, S. Y., Nanogap detector inside nanofluidic channel for fast
real-time label-free DNA analysis. Nano Letters 2008, 8, (5), 1472-1476.
15. Chen, Y.; Wang, X.; Hong, M. K.; Erramilli, S.; Mohanty, P.; Rosenberg, C.,
Nanoscale field effect transistor for biomolecular signal amplification. Applied
Physics Letters 2007, 91, (24), 243511-243514.
16. Neff, P. A.; Serr, A.; Wunderlich, B. K.; Bausch, A. R., Label-free electrical
determination of trypsin activity by a silicon-on-insulator based thin film resistor.
Chemphyschem 2007, 8, (14), 2133-2137.
17. Kang, B. S.; Wang, H. T.; Lele, T. P.; Tseng, Y.; Ren, F.; Pearton, S. J.; Johnson,
J. W.; Rajagopal, P.; Roberts, J. C.; Piner, E. L.; Linthicum, K. J., Prostate
specific antigen detection using AlGaN/GaN high electron mobility transistors.
Applied Physics Letters 2007, 91, (11).
18. Huang, X. J.; Choi, Y. K., Chemical sensors based on nanostructured materials.
Sensors and Actuators B-Chemical 2007, 122, (2), 659-671.
19. Park, I. Y.; Li, Z. Y.; Li, X. M.; Pisano, A. P.; Williams, R. S., Towards the
silicon nanowire-based sensor for intracellular biochemical detection. Biosensors
& Bioelectronics 2007, 22, (9-10), 2065-2070.
20. Poghossian, A.; Ingebrandt, S.; Abouzar, M. H.; Schoning, M. J., Label-free
detection of charged macromolecules by using a field-effect-based sensor
platform: Experiments and possible mechanisms of signal generation. Applied
Physics a-Materials Science & Processing 2007, 87, (3), 517-524.
21. Ingebrandt, S.; Offenhausser, A., Label-free detection of DNA using field-effect
transistors. Physica Status Solidi a-Applications and Materials Science 2006, 203,
(14), 3399-3411.
22. Neff, P. A.; Wunderlich, B. K.; Lud, S. Q.; Bausch, A. R., Silicon-on-insulator
based thin film resistors for quantitative biosensing applications. Physica Status
Solidi a-Applications and Materials Science 2006, 203, (14), 3417-3423.
29

23. Han, Y.; Offenhausser, A.; Ingebrandt, S., Detection of DNA hybridization by a
field-effect transistor with covalently attached catcher molecules. Surface and
Interface Analysis 2006, 38, (4), 176-181.
24. Lud, S. Q.; Nikolaides, M. G.; Haase, I.; Fischer, M.; Bausch, A. R., Field effect
of screened charges: Electrical detection of peptides and proteins by a thin-film
resistor. Chemphyschem 2006, 7, (2), 379-384.
25. Patolsky, F.; Timko, B. P.; Zheng, G. F.; Lieber, C. M., Nanowire-based
nanoelectronic devices in the life sciences. Mrs Bulletin 2007, 32, (2), 142-149.
26. Patolsky, F.; Zheng, G. F.; Lieber, C. M., Nanowire-based biosensors. Analytical
Chemistry 2006, 78, (13), 4260-4269.
27. Zheng, G. F.; Patolsky, F.; Cui, Y.; Wang, W. U.; Lieber, C. M., Multiplexed
electrical detection of cancer markers with nanowire sensor arrays. Nature
Biotechnology 2005, 23, (10), 1294-1301.
28. Stern, E.; Vacic, A.; Reed, M. A., Semiconducting Nanowire Field-Effect
Transistor Biomolecular Sensors. Ieee Transactions on Electron Devices 2008,
55, (11), 3119-3130.
29. Ishikawa, F. N.; Curreli, M.; Chen, P. C.; Zhang, R.; Cote, R. J.; Thompson, M.
E.; Zhou, C., A Calibration Method for Nanowire Biosensors to Suppress Device-
to-device Variation. ACS Nano 2009, 3, (12), 3969-3976.
30. Chen, R. J.; Choi, H. C.; Bangsaruntip, S.; Yenilmez, E.; Tang, X. W.; Wang, Q.;
Chang, Y. L.; Dai, H. J., An investigation of the mechanisms of electronic sensing
of protein adsorption on carbon nanotube devices. Journal of the American
Chemical Society 2004, 126, (5), 1563-1568.
31. Chen, R. J.; Bangsaruntip, S.; Drouvalakis, K. A.; Kam, N. W. S.; Shim, M.; Li,
Y. M.; Kim, W.; Utz, P. J.; Dai, H. J., Noncovalent functionalization of carbon
nanotubes for highly specific electronic biosensors. Proceedings of the National
Academy of Sciences of the United States of America 2003, 100, (9), 4984-4989.
32. Stern, E.; Klemic, J. F.; Routenberg, D. A.; Wyrembak, P. N.; Turner-Evans, D.
B.; Hamilton, A. D.; LaVan, D. A.; Fahmy, T. M.; Reed, M. A., Label-free
immunodetection with CMOS-compatible semiconducting nanowires. Nature
2007, 445, (7127), 519-522.
30

33. Cui, Y.; Wei, Q. Q.; Park, H. K.; Lieber, C. M., Nanowire nanosensors for highly
sensitive and selective detection of biological and chemical species. Science 2001,
293, (5533), 1289-1292.
34. Tang, X. W.; Bansaruntip, S.; Nakayama, N.; Yenilmez, E.; Chang, Y. L.; Wang,
Q., Carbon nanotube DNA sensor and sensing mechanism. Nano Letters 2006, 6,
(8), 1632-1636.
35. Kim, S. N.; Rusling, J. F.; Papadimitrakopoulos, F., Carbon nanotubes for
electronic and electrochemical detection of biomolecules. Advanced Materials
2007, 19, (20), 3214-3228.
36. Allen, B. L.; Kichambare, P. D.; Star, A., Carbon nanotube field-effect-transistor-
based biosensors. Advanced Materials 2007, 19, (11), 1439-1451.
37. Balasubramanian, K.; Burghard, M., Biosensors based on carbon nanotubes.
Analytical and Bioanalytical Chemistry 2006, 385, (3), 452-468.
38. Gruner, G., Carbon nanotube transistors for biosensing applications. Analytical
and Bioanalytical Chemistry 2006, 384, (2), 322-335.
39. Dubin, R. A.; Callegari, G. C.; Kohn, J.; Neimark, A. V., Carbon nanotube fibers
are compatible with mammalian cells and neurons. Ieee Transactions on
Nanobioscience 2008, 7, (1), 11-14.
40. Lacerda, L.; Bianco, A.; Prato, M.; Kostarelos, K., Carbon nanotubes as
nanomedicines: From toxicology to pharmacology. Advanced Drug Delivery
Reviews 2006, 58, (14), 1460-1470.
41. Lacerda, L.; Raffa, S.; Prato, M.; Bianco, A.; Kostarelos, K., Cell-penetrating
CNTs for delivery of therapeutics. Nano Today 2007, 2, (6), 38-43.
42. Popov, A. M.; Lozovik, Y. E.; Fiorito, S.; Yahia, L., Biocompatibility and
applications of carbon nanotubes in medical nano robots. International Journal of
Nanomedicine 2007, 2, (3), 361-372.
43. Mattson, M. P.; Haddon, R. C.; Rao, A. M., Molecular functionalization of carbon
nanotubes and use as substrates for neuronal growth. Journal of Molecular
Neuroscience 2000, 14, (3), 175-182.
44. Star, A.; Gabriel, J. C. P.; Bradley, K.; Gruner, G., Electronic detection of specific
protein binding using nanotube FET devices. Nano Letters 2003, 3, (4), 459-463.
31

45. Gao, Z.; Agarwal, A.; Trigg, A. D.; Singh, N.; Fang, C.; Tung, C. H.; Fan, Y.;
Buddharaju, K. D.; Kong, J., Silicon Nanowire Arrays for Label-Free Detection
of DNA. Analitical Chemisty 2007, 79, (9), 3291-3297.
46. Patolsky, F.; Timko, B. P.; Yu, G. H.; Fang, Y.; Greytak, A. B.; Zheng, G. F.;
Lieber, C. M., Detection, stimulation, and inhibition of neuronal signals with
high-density nanowire transistor arrays. Science 2006, 313, (5790), 1100-1104.
47. Kumar, M. J.; Reed, M. A.; Amaratunga, G. A. J.; Cohen, G. M.; Janes, D. B.;
Lieber, C. M.; Meyyappan, M.; Wernersson, L. E.; Wang, K. L.; Chau, R. S.;
Kamins, T. I.; Lundstrom, M.; Yu, B.; Zhou, C. W., Special Issue on Nanowire
Transistors: Modeling, Device Design, and Technology. Ieee Transactions on
Electron Devices 2008, 55, (11), 2813-2819.
48. Wang, W. U.; Chen, C.; Lin, K. H.; Fang, Y.; Lieber, C. M., Label-free detection
of small-molecule-protein interactions by using nanowire nanosensors.
Proceedings of the National Academy of Sciences of the United States of America
2005, 102, (9), 3208-3212.
49. Lieber, C. M.; Wang, Z. L., Functional nanowires. Mrs Bulletin 2007, 32, (2), 99-
108.
50. Timko, B. P.; Cohen-Karni, T.; Yu, G. H.; Qing, Q.; Tian, B. Z.; Lieber, C. M.,
Electrical Recording from Hearts with Flexible Nanowire Device Arrays. Nano
Letters 2009, 9, (2), 914-918.
51. Curreli, M.; Zhang, R.; Ishikawa, F. N.; Chang, H.-K.; Cote, R. J.; Zhou, C.;
Thompson, M. E., Real-Time, Label-Free Detection of Biological Entities Using
Nanowire-Based FETs. IEEE Transactions on Nanotechnology 2008, 7, (6), 651 -
667.
52. Patolsky, F.; Zheng, G. F.; Hayden, O.; Lakadamyali, M.; Zhuang, X. W.; Lieber,
C. M., Electrical detection of single viruses. Proceedings of the National
Academy of Sciences of the United States of America 2004, 101, (39), 14017-
14022.
53. Stern, E.; Wagner, R.; Sigworth, F. J.; Breaker, R.; Fahmy, T. M.; Reed, M. A.,
Importance of the Debye Screening Length on Nanowire Field Effect Transistor
Sensors. Nano Letters 2007, 7, (11), 3405-3409.
54. Maehashi, K.; Katsura, T.; Kerman, K.; Takamura, Y.; Matsumoto, K.; Tamiya,
E., Label-free protein biosensor based on aptamer-modified carbon nanotube
field-effect transistors. Analytical Chemistry 2007, 79, (2), 782-787.
32

55. Elfstrom, N.; Juhasz, R.; Sychugov, I.; Engfeldt, T.; Karlstrom, A. E.; Linnros, J.,
Surface charge sensitivity of silicon nanowires: Size dependence. Nano Letters
2007, 7, (9), 2608-2612.
56. Heitzinger, C.; Klimeck, G., Computational aspects of the three-dimensional
feature-scale simulation of silicon-nanowire field-effect sensor for DNA
detaction. Journal of Computational Electronics 2007, 6, 387-390.
57. Kim, A.; Ah, C. S.; Yu, H. Y.; Yang, J. H.; Baek, I. B.; Ahn, C. G.; Park, C. W.;
Jun, M. S.; Lee, S., Ultrasensitive, label-free, and real-time immunodetection
using silicon field-effect transistors. Applied Physics Letters 2007, 91, (10).
58. Nair, P. R.; Alam, M. A., Screening-limited response of nanobiosensors. . Los
Alamos National Laboratory, Preprint Archive, Condensed Matter 2007, 1-7.
59. Nair, P. R.; Alam, M. A., Performance limits of nanobiosensors. Applied Physics
Letters 2006, 88, (23), 233120-233123.
60. Nair, P. R.; Alam, M. A., Design Considerations of Silicon nanowire biosensors.
Ieee Transactions on Electron Devices 2007, 54, 3400-3407.
61. Bunimovich, Y. L.; Shin, Y. S.; Yeo, W. S.; Amori, M.; Kwong, G.; Heath, J. R.,
Quantitative real-time measurements of DNA hybridization with alkylated
nonoxidized silicon nanowires in electrolyte solution. Journal of the American
Chemical Society 2006, 128, (50), 16323-16331.
62. Sheehan, P. E.; Whitman, L. J., Detection limits for nanoscale biosensors. Nano
Letters 2005, 5, (4), 803-807.
63. Curreli, M.; Li, C.; Sun, Y. H.; Lei, B.; Gundersen, M. A.; Thompson, M. E.;
Zhou, C. W., Selective functionalization of In2O3 nanowire mat devices for
biosensing applications. Journal of the American Chemical Society 2005, 127,
(19), 6922-6923.
64. Liu, R.; Barrick, J.; Szostak, J. W.; Roberts, R. W., Optimized Synthesis of RNA-
Protein Fusions for In Vitro Protein Selection. Methods in Enzymology 2000, 317,
268-293.
65. Barrick, J. E.; Takahashi, T. T.; Ren, J.; Xia, T.; Roberts, R. W., Large Libraries
Reveal Diverse Solutions to an RNA Recognition Problem. Proceedings of the
National Academy of Sciences of the United States of America 2001, 98, 12374-
12378.
33

66. Roberts, R. W.; Szostak, J. W., RNA-peptide fusions for the in vitro selection of
peptides and proteins. Proceedings of the National Academy of Sciences of the
United States of America 1997, 94, 12297-12302.
67. Takahashi, T. T.; Austin, R. J.; Roberts, R. W., mRNA display: ligand discovery,
interaction analysis and beyond. Trends in Biochemical Sciences 2003, 28, (3),
159-165.
68. Olson, C. A.; Liao, H. I.; Sun, R.; Roberts, R. W., nRna display selection of a
high-affinity, modification-specific phospho-IkBalpha-binding fibronectin. ACS
Chemical Biology 2008, 3, (3), 480-485.
69. Hahm, J.; Lieber, C. M., Direct ultrasensitive electrical detection of DNA and
DNA sequence variations using nanowire nanosensors. Nano Letters 2004, 4, (1),
51-54.
70. Zhang, G.-J.; Zhang, G.; Chua, J. H.; Chee, R.-E.; Wong, E. H.; Agarwal, A.;
Buddharaju, K. D.; Singh, N.; Gao, Z.; Balasubramanian, N., DNA Sensing by
Silicon Nanowire: Charge Layer Distance Dependence. Nano Letters 2008, 8, (4),
1066-1070.
71. Rosenblatt, S.; Yaish, Y.; Park, J.; Gore, J.; Sazonova, V.; McEuen, P. L., High
performance electrolyte gated carbon nanotube transistors. Nano Letters 2002, 2,
(8), 869-872.

34

CHAPTER 2:
SURFACE MODIFICATION OF CARBON NANOTUBES
Chapter Outline
2.1. Introduction to Carbon Nanotubes
2.2. Coordination of RuCp* to the Carbon Nanotube Sidewalls
2.3. Recent Achievements on the Coordination of RuCp* to Nanotubes
2.4. Controlled Covalent Surface Modification
2.5. Selective Functionalization of Carbon Nanotubes
2.6. Conclusions
2.7. Chapter 2 References

2.1 Introduction to Carbon Nanotubes
Carbon nanotubes were discovered in 1991,
1
and rapidly became the most
investigated material of the past decade. Carbon nanotube is an allotropic form of carbon
along with diamond, graphene, amorphous, and fullerenes, as shown in Figure 2.1.
Carbon nanotubes are the ideal material for a wide range of applications because
they possess unique physical, chemical (relatively inert), thermal, electronic, and
mechanical properties.
2-10
Some of the nanotubes’ outstanding properties relative to my
project include:
1) A very high surface to volume ratio which is optimal for sensing applications.
11-14
In
fact, every atom in a carbon nanotube is located on the surface.

35

2) Relatively low cytotoxicity.
8, 15, 16
While the safety of nanotubes is still a controversial
subject, nanotubes appear to be non-toxic to cells when immobilized on a solid matrix
such as an electrical circuit,
17
making them the ideal material for electrical
circuits/biology interface.

Despite these outstanding properties, the large scale usage of nanotubes is still
facing several practical drawbacks, the most important of which can be summarized as
follows:
1) Difficulty of covalently functionalizing the nanotube surface.
18-20
The covalent
functionalization of the nanotube sidewalls has not been achieved without the
disruption of the sp
2
hybridized carbon network, which severely affects the
nanotubes’ electronic properties.

Figure 2.1. Allotropic forms of carbon. Elemental carbon can be found in nature under several
allotropic forms, including amorphous, graphite, diamond, carbon nanotubes, and fullerenes.

36

2) Low solubility, which translates into difficult solution processing.
21, 22
Soluble
nanotubes are highly desirable for bulk chemical processes. However, suspensions of
individual, unbundled nanotubes can only be achieved in low concentrations and
under drastic conditions.
3) Difficulty in fine-tuning the electrical properties via the introduction of dopants.
23

The electronic properties of semiconducting nanotubes cannot be easily fine-tuned by
introducing a doping element (nitrogen, boron, phosphorus) as in the case of silicon.
The fabrication of field effect transistors using nanotubes was first demonstrated
in 1998 by Professor Cees Dekker’s research group.
24
Immediately after this
demonstration, researchers envisioned a large number of applications for nanotube
electronics. However, they soon realized that carbon nanotubes come in two types:
semiconducting (chiral) and metallic (achiral), as illustrated in Figure 2.2. Transistors
made using mainly semiconductive nanotubes are highly desirable since very large
On/Off current ratios can be obtained. The synthesis of nanotubes always yields both
Figure 2.2. Metallic and semiconducting carbon nanotubes. The chirality of the nanotube determines
whether the nanotube will be metallic (left) or semiconducting (right).

37

metallic (~30%) and semiconducting nanotubes (~70%). A significant amount of research
is devoted to the efficient separation of nanotube mixtures,
25, 26
conversion of metallic
into semiconducting nanotubes,
27
and synthesis of single chirality nanotubes,
28
but so far
it has not been economically feasible for industry to apply any of these methods.

2.2 Coordination of RuCp* to the Carbon Nanotube Sidewalls
Background. I authored an original proposal (as a requirement for my qualifying
exams) for an experiment that would investigate a chemical method to address the issues
of covalent functionalization, low solubility, and nanotube doping. This method might
also improve other properties of nanotubes such as thermal conductivity, interfacing with
biology, possible sorting of metallic and semiconducting nanotubes, new optical
properties (absorption/emission), etc. This method is based on the synthesis of a
ruthenium-based metal sandwich compound on the nanotube sidewalls. Specifically, this
compound would be penthamethylcyclopentadienyl-ruthenium or, in short, Cp*-Ru. A
molecular modeling of such compound is shown in Figure 2.3.

Figure 2.3. Schematic representation of [Cp*Ru]
+
units coordinated to the surface of a carbon
nanotube.

38

Metal sandwich compounds. Metal sandwich compounds are a class of
organometallic compounds in which the central metal atom is bound between two cyclic
organic molecules containing a delocalized π system. This metal is covalently bound to
all the atoms of the organic ring, thus preserving its aromaticity. In the literature,
29
there
are numerous examples of RuCp*- units η
6
bound to aromatic molecules such as benzene,
naphthalene, anthracene, pyrene, as shown in Figure 2.4. More recently, the crystal
structure of Cp*-Ru-corannulene has also been published.
30-33
This trend shows that Cp*-
Ru could be attached to increasingly large conjugated systems. All these Cp*Ru-arene
complexes seem to be air stable. Moreover, the surface of corannulene has a curvature
similar to ~1.5 nm carbon nanotubes.

The synthesis of the Cp*-Ru complex on the nanotube sidewalls has never been
reported in the literature since it faces several new synthetic challenges. One challenge is
the curvature of the nanotube surface, but it can be overcome by using large diameter
(>1.5nm) nanotubes. Another challenge may come from the much extended aromatic
conjugation of the nanotube: the larger the conjugation, the longer the Ru-arene bond,
Ru (II)
Ru (II) Ru (II) Ru (II)
(C H
3
)
5 (C H
3
)
5
(C H
3
)
5
(C H
3
)
5
Ru (II)
(C H
3
)
5
(a)
(b)
(c)
(d)
(e)

Figure 2.4. Some Cp*Ru-Arene complexes discussed in the literature. These arenes are: (a) benzene,
(b) naphthalene, (c) anthracene, (d) pyrene, and (e) corannulene.

39

resulting in a weaker Ru-arene bond.
32
Thus, it is expected that the Ru-nanotube bond
may also be weak, due to the extended conjugation of the nanotube.

Stability of Cp*-Ru Sandwich Compounds. In the case of Ru-arene complexes,
the aromatic ring becomes more labile and easily displaced by a competing, incoming
ligand (either thermal or photochemical displacement) as a function of extended
conjugation. This process is illustrated in Figure 2.5 for a molecule of THF coordinating
to Cp*-Ru-pyrene. The first step of the THF coordination is the ability of Cp*-Ru-pyrene
complex to create another ligand site so that THF will coordinate to Ru. This creation of
a new ligand site might involve tilting the Cp* ring tilts, and consequent Ru becomes η
4
coordinated to pyrene. Other two incoming THFs will eventually displace pyrene. This
displacement process can easily be monitored by UV-Vis spectroscopy, as shown in
Figure 2.6. In this experiment, a solution of pyrene-Ru-Cp* in degassed CH
2
Cl
2
was left
standing in 10% THF v/v in a sealed cuvette, and the absorption spectrum was recorded
at the times indicated in the figure legend. No significant displacement occurred within
the first 475 minutes (curves (b)). However, after two days standing in 10% THF, the
typical shapeline of “free” pyrene started to appear, and five days later, (d), the typical
spectrum of “free,” non-coordinated pyrene was clearly distinguishable. The cuvette was
sealed with a Teflon cap and kept in the dark in between runs.

40

(b)
(a)
THF THF

Figure 2.5. Simulation of the displacement of pyrene by THF from the sandwich compound Cp*Ru-
pyrene. (a) The cation Cp*Ru-pyrene is represented using two different graphics (space filling and ball-
and-spoke). (b) The first step toward the displacement of pyrene by THF molecules requires tilting the
Cp* ring, so a new coordination site is formed for the incoming THF. Two more incoming THFs will
eventually displace the pyrene from the sandwich compound.
300 360 420
0.0
0.8
1.6
absorbance
wav elenght (nm)
(a)
(b)
(c)
(d)

Figure 2.6. Monitoring the THF displacement of pyrene in Pyrene-RuCp* by UV-Vis spectroscopy.
(a) UV-Vis of a solution of pyrene-Ru-Cp* in degassed CH2Cl2. After recording spectrum (a), THF
was added into the cuvette so the solution was 10% in THF v/v. (b) Another spectrum was recorded 5
minutes later followed by nine more spectra within 475 minutes. These ten spectra basically overlap. (c)
Another spectrum was recorded 2 days later, and starts to show “free” pyrene. (d) 5 days later, the
spectrum of “free” pyrene was clearly distinguishable.

41

A similar experiment, this time carried out in acetonitrile (MeCN), demonstrated
that the size and nucleophilicity of the solvent affects the rate of pyrene displacement, as
shown in Figure 2.7. Monitoring the MeCN displacement of pyrene from Pyrene-RuCp*
was accomplished by UV-Vis spectroscopy. After 120 minutes, there was already a sign
of “free” pyrene in 10% MeCN v/v. After two days of standing in the presence of
MeCN, the presence of “free” pyrene became very evident (c).

However, in the case of the complex Cp*-Ru-nanotube, ring tilting of Cp*, and
therefore the displacement of Cp*-Ru from the nanotube, might be more difficult because
the nanotube is bulky, as illustrated in Figure 2.8. Obstructed Cp* ring tilting may
300 350 400
0
1
2
absorbance
wav elenght (nm)
(a)
(b)
(c)

Figure 2.7. Monitoring the MeCN displacement of pyrene in Pyrene-RuCp* by UV-Vis spectroscopy.
(a) UV-Vis of a solution of pyrene-Ru-Cp* in degassed CH2Cl2. After recoring spectrum (a), MeCN
was added into the cuvett so the solution was 10% in MeCN v/v. (b) Another spectrum was recording 3
minutes later followed by 13 more spectra within 465 minutes. After 120 minutes, there was a sign of
“free” pyrene. (c) Another spectrum was recorded 2 days later, and shows “free” pyrene. The cuvett
used was capped with a Teflon cap and kept in the dark in between runs.

42

prevent a competing ligand from accessing the central atom of Ru, giving the Cp*-Ru-
nanotube complex a higher stability than analogous arene complexes. This conclusion has
been deduced by computational modeling and has not been demonstrated experimentally.
A ligand much smaller than THF, such as OH
-
, or MeCN might be able to easily display
Cp*-Ru from the nanotubes.

Strategies developed to synthesize these complexes must take into account the
lower thermal and photo stability of very large conjugated systems. The synthesis of any
metal sandwich complex on the sidewalls of carbon nanotubes has never been reported in
the literature.

(a)
(b)
THF THF

Figure 2.8. Simulation of the displacement of the nanotube from the Cp*Ru-nanotube complex by
THF. (a) The cation Cp*Ru-nanotube is represented using two different graphics (space filling and ball-
and-spoke). (b) The first step toward the displacement of the nanotube by a THF molecule would
require tilting the Cp* ring. Unlike in the case of pyrene, the carbon nanotube significantly prevents
tilting, protecting the RuCp* from incoming nucleophiles.

43

The Carbon Nanotubes Used in This Study. Raw carbon nanotubes used in
this experiment were provided by one of our collaborators. These “as received” HiPCO
nanotubes were characterized by SEM, UV-Vis, Raman, and IR spectroscopy. Samples
for SEM were prepared by suspending the nanotubes in dichloroethane (DCE) with the
aid of a bath sonicator and spin coating the solution on a silicon wafer, followed by
solvent evaporation. SEM images of these nanotubes (Figure 2.9) show that they still
contain a significant amount of amorphous carbon and/or catalyst nanoparticles.
Therefore, these nanotubes should have been purified using a probe sonicator and an
ultracentrifuge, but these instruments were not available at that time.

Carbon nanotubes were also characterized by Raman spectroscopy (532 nm
laser). A typical spectrum of a nanotube is shown in Figure 2.10. Distinctive features are:
 The radial breathing mode (RBM) at 175 cm
-1
. A single line indicates that the
nanotubes have a uniform diameter, and by using the formula RBM=A/d + B, where
A=234 and B=10, the resulting diameter is 1.4 nm.
Figure 2.9. SEM pictures of the HiPCO nanotubes used in this experiment.

44

 The D-line given by the presence of defective/disordered sites, mainly a combination
of 5-7-7-5 rings.
 The G-Line, tangential mode of the phonon traveling longitudinally on the nanotube.
 The G’-line as overtone of the D-Line.
The IR spectrum of the unmodified carbon nanotubes is shown in Figure
2.11. It is basically featureless. Carbon nanotubes are expected to have the same IR
active modes of graphite, which are at 1590 cm
-1
(E
1u
) and 868 cm
-1
(A
2u
). However,
for commercial nanotubes, intense bands of absorption should be located at 1110,
1535, and 1700 cm
-1
with a weak band at 830 cm
-1
.
0 500 1000 1500 2000 2500 3000
0
2000
4000
wav enumber (1/cm)
175
1340
1599
1747
2673
24 43
RBM
D-Line
G-Line
G'-Line
Second order modes

Figure 2.10. Typical Raman spectra of “as received” carbon nanotubes.

45

The UV-Vis spectrum of row nanotubes, recorded in DCE, is shown in Figures
2.16(a) and 2.22(a). The spectrum is essentially featureless. This means that the
experiments did not start with suspensions of individual nanotubes.

2.2.a Synthetic Routes for Cp*-Ru-Nanotube
In this study, the Cp*-Ru- complex of nanotubes was prepared using two different
synthetic procedures:
Route 1. The first route requires refluxing the precursor [Cp*Ru
(II)
-
(MeCN)
3
][PF
6
] in a suspension of nanotubes in DCE,
29
as schematically shown in Figure
2.12. The expected product is shown in this scheme.
3000 2500 2000 1500 1000
35
40
45
% Transmittance
wav enumber (1/cm)
2978
292 6
235 3
173 4
1 695
822
1076
1335
1536

Figure 2.11. Typical IR spectra of “as received” carbon nanotubes.

46

Route 2. The first step of the second route requires reducing the carbon
nanotubes to their corresponding Na
+
salt, as shown in Figure 2.13. This can be
accomplished by reacting a suspension of nanotubes in dry THF with a powerful
reducing agent such as the sodium salts of naphthalene or benzophenone. The reduced
carbon nanotubes are then stirred in a THF solution of Cp*Ru
(III)
-Cl
2
. Electron transfer
between the nanotubes and the Ru
(III)
chloride salt reduces Ru
(III)
to Ru
(II)
, which then
coordinates to the sidewalls of the nanotubes. Later it was found out that THF, even if
considered a weak ligand, may still “strongly” coordinate Cp*-Ru, thus impeding the
coordination of Cp*Ru to the nanotubes. To prevent this effect we used diglyme as
solvent. The product indicated in Figure 2.13 is the expected product; the actual product
of this reaction may be different.
In the case of route 2, there is a controversy on the charge of the resulting
complex. The IR spectrum (Figure 2.25) shows very little PF
6
-
present. Absence of the
PF
6
counter ion might be explained by a “negative” η
6
carbon ring coordinating Cp*-Ru.
This “negative” carbon ring on the nanotube might quickly form by electron migration

[Cp*Ru(CH
3
-CN)
3
]
+

-
PF
6
Ru(II)
Reflux in DCE
PF
6
Ru(II)
N N
N
C
C
C
P
F
F
FF
F
F

Figure 2.12. Synthetic route 1. A suspension of carbon nanotubes in DCE is refluxed in the presence of
the precursor [Cp*Ru
(II)
-(MeCN)
3
][PF
6
].

47

from an adjacent reduced region of the nanotube, as illustrated in Figure 2.14. If this is
the case, the reaction product Cp*-Ru-nanotube is neutral, and no counter ions such as
Cl
-
or PF
6
-

are necessary to balance to the positive Cp*-Ru
(II)
moiety. The actual product
might also be a mixture of the two structures (positive and neutral Cp*-Ru-nanotube),
with a predominance of the second structure (neutral).

Ru (II)
Ru
(III)
Cl
Cl
Cl
Cl
Ru
(III)
Cl
Cp* Ru
(III)
Cl
2
polym er
Cl
Cl
TH F
Dry TH F
Stirr at RT
R u(III)
2N aC l
Na P F
6
PF
6
Na
SW N T
Na
Na
Na
Na
Na
O

Figure 2.13. Synthetic route 2. Carbon nanotubes are first reduced to the corresponding Na salts using
reduced naphthalene. The reduced nanotubes are expected to reduce Ru(III) to Ru(II) and thus the
Cp*Ru unit can coordinate to the nanotubes.
Cl
Cl
TH F
R u(III)
Na
Na
Ru(II)
Na
Na C l
Cl
Ru(II)
2N aC l

Figure 2.14. Possible product rearrangement following synthetic route 2.

48

Cp*-Ru-Pyrene as a Model Compound for Cp*-Ru-Nanotube. The synthesis
of the complex [Cp*Ru-pyrene][PF
6
] was attempted following both routes 1 and 2, with
the intention to use this compound as a model reference for the characterization of the
Cp*Ru-Nanotube. This complex, when stored dry in a closed vial, appeared to be stable
within the laboratory atmosphere for over 3 years (checked by UV spectroscopy and
1
H
NMR, no free pyrene was observed).
The vibrational spectrum of [Cp*Ru-pyrene][PF
6
] is shown in Figure 2.15, and
the strongest band of absorption is given by P-F stretch at 831 cm
-1
produced by the
counter ion PF
6
(see inset). Other important features that will help identify our products
are the C-H stretches of the Cp* methyl groups at 2916 cm
-1
and several vibrational
modes associated with the Cp* ring (1685, 1473, 1386, and 1028 cm
-1
). The Cp* ring

3000 2500 2000 1500 1000
115
120
125
130
135
140
1027.9
1186
1386.6
1473.3
1576.5
1685.5
1793.5
2368.2
2422.2
2916.8
3080.7
% Transmittance
Wavenumber (1/cm)
Py rene Ru Cp*
3000 2500 2000 1500 1000
30
40
50
60
70
80
90
100
110
120
130
140
831.2
1386.6
Ru(II)
PF
6

Figure 2.15. Vibrational spectrum of [Cp*Ru-pyrene][PF
6
].

49

peaks have been assigned with the aid of computer modeling and quantum mechanical
calculations to distinguish the Cp* from vibrational modes of pyrene. C-H stretches at
3080.7 cm
-1
belong to pyrene.
The Uv-Vis spectrum of Cp*-Ru-Pyrene was also recorded and is shown in
Figures 2.6 (a), 2.7 (a), 2.16, and 2.22.

2.2.b. Cp*-Ru-Nanotube Product Characterization
The synthetic products of route 1 and route 2 were characterized by UV/Vis, IR,
and Raman spectroscopy.
Route 1 Product Characterization: UV-Vis Spectroscopy. UV-Vis
spectroscopy can be used to monitor reactions when both the starting material and
products have different absorption spectra. The spectra of starting reagents carbon
nanotubes and Cp*Ru(MeCN)
3
is shown in Figure 2.16 (a) and (d), respectively. The
carbon nanotube spectrum is essentially featureless, whereas that of Cp*Ru(MeCN)
3
shows a very weak, broad shoulder centered at 410 nm. The reaction product, Cp*Ru-
Nanotube, exhibits a “strong” absorption centered at 312 nm, a weak shoulder centered at
345 nm, and a broad absorption around 410 nm. When this spectrum is compared to that
of Cp*Ru-Pyrene, both have a transition positioned at 312 nm but the lower energy
transition is significantly red-shifted for Cp*Ru-nanotube.
This difference might be due to the higher conjugation of the nanotube with
respect to pyrene. Table 2.1 lists the position of this absorption peak for different Cp*Ru-

50

Arene complexes (small conjugation) discussed in the literature.
29
Lamda max is mostly
in the range 360-370 nm, but anthracene has λ
max
at 480 nm.

300 350 400 450 500 550 600
-0.2
-0.1
0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
1.0
1.1
Absorbance (Arb. units)
Wav elenght (nm)
312
355
(a)
(b)
(c)
(d)

Figure 2.16. UV spectra of reagents and products involved in the synthesis according to route 1. The
spectra have been normalized or shifted for clarity. (a)-Green: Suspension of nanotubes in THF. (b)-
Red: Product(s) of the reaction according to route 1--Cp*Ru-nanotube--suspended in THF. (c)-Orange:
Spectrum of Cp*Ru-Pyrene in DCE, the lambda max is located at 312 and 355 nm. (d)-Blue: Spectrum
of the precursor Cp*Ru(MeCN)
3
in dry THF.
Cp*Ru
Cp*Ru Cp*Ru
Cp*Ru

Table 2.1. Electronic absorption spectra for several Ru-based sandwich compounds. Those compounds
highlighted in the red box involve complexes of RuCp* η
6
-coordinated to conjugated arenes such as
naphthalene, anthracene, pyrene, and chrysene. The counter ion of all these complexes is PF
6
-
.

51

Route 1 Product Characterization: Infrared Spectroscopy. The IR spectrum of
the starting material (Figure 2.17) and [Cp*Ru-Pyrene][PF
6
] (Figure 2.15) was compared
to that of the reaction product (Figures 2.18 and 2.19). In the starting material, [Cp*-Ru-
(MeCN)
3
][PF
6
], the main absorption peak in the spectrum results from the P-F stretch at
828 cm
-1
produced by the counter ion PF
6
-
. Other important features in the starting
material are the C-H stretches of the Cp*’s methyl groups at 2988-2924 cm
-1
and several
vibrational modes associated with the Cp* ring (1674, between 1455 and 1385, and 1031
cm
-1
).

The IR spectrum of the Route 1 product, [Cp*Ru-Nanotube][PF
6
] is shown in
Figure 2.18. This spectrum shows a progressively dropping baseline, similar to the
spectrum of unmodified nanotubes (Figure 2.11). The raw nanotube spectrum was

3500 3000 2500 2000 1500 1000
96
100
104
% Transmittance
wavenum ber (1/cm )
3649
2988
2924
2369
2320
2265 1736
1674
1455
1419
1385
1072
1031
828
4000 3000 2000 1000
20
40
60
80
100
830
1033
1421
2267 2923
Ru(II)
N N
N
C
C
C
P
F
F
FF
F
F

Figure 2.17. IR of [Cp*Ru(NCMe)
3
][PF
6
], route 1 starting material. The sample was exposed to air for
a few minutes during acquisition. The inset shows the whole spectrum, in the range 4000-700 cm
-1
.

52

subtracted from that of the product from route 1, and is shown in Figure 2.19. This
spectrum has the characteristic vibrational modes of Cp*-Ru unit. The C-H stretch of the
Cp* methyl groups are visible around 2972 and 2926 cm
-1
, similar to the spectra of the
pyrene analog and the tris-MeCN precursor reagent. The vibrations associated with the
Cp* ring modes, located at 1670, 1460, 1382, 1282-45, 1132, 1022 cm
-1
are also similar
to those from the pyrene analog (1685, 1473, 1386, 1186, and 1028 cm
-1
). However, the
intensity of the P-F band at 834 cm
-1
is much weaker than expected, suggesting less PF
6
-

than predicted.

3000 2500 2000 1500 1000
64
66
68
70
72
74
76
78
80
82
84
86
% Transmittance
wav enumber (1/cm)
296 2
235 5
19 85
16 66
1 560
1 424
1 020
83 4

Figure 2.18. IR spectrum of nanotube treated according to route 1. The expected product is [Cp*Ru-
nanotube][PF
6
].

53

Route 1 Product Characterization: Raman Spectroscopy. The Raman
spectrum of the precursor [Cp*Ru(MeCN)
3
][PF
6
] was recorded using a 532 nm excitation
wavelength, and is shown in Figure 2.20. Notably, the vibrations from the methyl groups
of the Cp* at 2941 cm
-1
and the CN stretch at 2268 cm
-1
produce relatively intense
bands. The Cp* ring vibrations show up in the broad peak between 1430 cm
-1
and 1560
cm
-1
. The counter ion PF
6
seems invisible under the conditions governing these
measurements.
3000 2500 2000 1500 1000
170
180
190
wav enumber (1/cm)
2 972 2926
2372
2 345
198 5
16 70
1450
12 82
1245
1132
10 22
83 4

Figure 2.19. IR spectrum of nanotube treated according to route 1 after the mathematical subtraction of
the IR spectrum of untreated nanotube.

54

The Raman spectrum of the route 1 product is significantly different from that of bare
nanotubes, and is shown in Figure 2.21. Clearly, there is a broadening of the G line,
which overlaps with the D line and some lines from the Cp* ring. This broadening
indicates a decrease in symmetry on the sidewalls on carbon nanotubes, suggesting that
Cp*Ru is coordinated to the nanotube sidewalls. Also, the vibrations from the Cp*
methyl groups are visible at 2919 cm
-1
, a blue shifted with respect to the starting material
by 22 wavenumbers but by 137 with respect to pyrene-RuCp*.

500 1000 1500 2000 2500 3000
0
200
400
600
800
1000
2941.84
2268.5
1376.62
941.763
264.136
wavenumber (1/cm)
444.6
1560.7
Ru(II)
N N
N
C
C
C
P
F
F
FF
F
F

Figure 2.20. Raman spectra of the complex precursor [Cp*Ru(MeCN)
3
][PF
6
].

55

Route 2 Product Characterization: UV-Vis Spectroscopy. As discussed earlier,
bare nanotubes have a featureless spectrum whereas the precursor Cp*Ru
(III)
-Cl
2
shows a
strong absorption centered around 438 nm. The spectrum of the route 2 product shows
two features: a “strong” absorption at 312 nm and a very weak shoulder around 410 nm,
as shown in Figure 2.22. When this spectrum is compared to that of Cp*Ru-Pyrene, both
have a transition positioned at 312 nm but the lower energy transition is significantly red-
shifted for Cp*Ru-nanotube. This difference might be due to the higher conjugation of
the nanotube with respect to pyrene.
500 1000 1500 2000 2500 3000
0
100
200
300
400
2919.2
2414.2
2030.2
1573.4
437.1

wavenumber (1/cm)

Figure 2.21. Raman spectra of route 1 product. The expected product is [Cp*Ru-nanotube][PF
6
].

56

Route 2 Product Characterization: Infrared Spectroscopy. The starting
material Cp*RuCl
2
shows the characteristic vibrational modes of a coordinated Cp* ring
(Figure 2.23). The C-H stretches are centered at 2983 and 2914 cm
-1
, while vibrational
bands associated with the Cp* ring, can be seen at 1672, 1444, 1371, and 1022 cm
-1
.
Figures 2.24 and 2.25 show the IR spectra of modified nanotubes according to
route 2. The very weak band corresponding to the P-F stretch indicates that PF
6
might not
be present on the sidewalls of nanotubes. The absence of PF
6
can be interpreted as an
unsuccessful anion exchange (Cl
-
for PF
6
-
). Thus, chloride might still be the counter ion,
or the reduced carbon nanotube itself can be the counter ion for Cp*Ru according to the
reduction mechanism illustrated in Figure 2.13. In any case, this IR spectrum shows the
300 350 400 450 500 550 600
0.0
0.2
0.4
0.6
0.8
1.0
Abs units
wav elenght (nm)
438
312
355
(a)
(b)
(c)
(d)

Figure 2.22. UV spectra of reagents and products involved in the synthesis according to route 2. The
spectra have been normalized or shifted for clarity. (a) Nanotube suspension in THF. (b) The product
(probably Cp*Ru-nanotube) suspended in THF. (c) Absorption of a DCE solution of Cp*Ru-Pyrene,
the lambda max is located at 312 and 355 nm. (d) Spectrum of the precursor Cp*Ru
(III)
-Cl
2
in dry THF.

57

presence of Cp*Ru unit in the product, evidenced by bands at 2967 and 2914 cm
-1
for the
C-H stretches of the CP* methyl groups, and the vibrational modes associated with the
Cp* ring at1656, 1438, 1371, 1214, and 1020 cm
-1
.

3000 2500 2000 1500 1000
90
95
100
% Transmittance
wav enumber (1/cm)
29 83
2 914
2 362
16 72
156 0
1 444
13 71
102 2

Figure 2.23. IR spectra of Cp*RuCl
2,
the Ru precursor according to route 2.
3000 2500 2000 1500 1000
188
190
192
194
196
wav enumber (1/cm)
2967
2914
1955
1656 1438
1371
1214
1020
837

Figure 2.24. IR spectrum of nanotube treated according to route 2 after the mathematical subtraction of
the IR spectrum of untreated nanotube.

58

Route 2 Product Characterization: Raman Spectroscopy. The first step of
route 2 synthesis requires the reduction of the nanotube to their corresponding Na salt.
The success of the reduction reaction can be monitored by Raman spectroscopy. In fact,
the spectrum of reduced nanotubes (Figure 2.26) does not have the RBM line, due to the
strong disturbance in the hexagonal lattice of the sidewalls by the sodium salt. Also, the
relative intensity of the D line increased, meaning an increase in disorder within the sp
2
-
bonded carbon framework.
After the reduced nanotubes were stirred in a solution of Cp*Ru
(III)
, the resulting
product has the Raman spectra shown in Figure 2.27. This spectrum is very similar to that
of unmodified nanotubes, suggesting that very little functionalization had occurred.

3000 2500 2000 1500 1000
72
80
88
As received SWNT
Cp*Ru-SWNT via Ru(III)
% Transmittance
wavenumber (1/cm)
2910
2983
1372
1661
1018
SWNT
SWNT-RuCp*

Figure 2.25. IR spectra of route 2 product compared to that of bare nanotube.

59

500 1000 1500 2000 2500 3000
-500
0
500
1000
1500
2000
2500
3000
2658.8
1595.5
1333.2
wav enumber(1/cm)

Figure 2.26. Raman spectrum of the carbon nanotubes Na salt.
500 1000 1500 2000 2500 3000
0
10000
20000
30000
40000
50000
2665.8
1587.2
1340.2
1079.4
67.1
w a v e nu m b er (1 /c m )

Figure 2.27. Raman spectrum of the carbon nanotubes treated according to route 2 synthesis.

60

Route 2 Product Characterization: SEM Imaging Using Nanotube Mats. A
mat of carbon nanotubes was grown on a Si wafer by CVD methods, as shown in the top
SEM image in Figure 2.28. Typically, nanotubes grown by CVD have a larger diameter
(between 1 and 3 nm) than HipCO produced nanotubes (1.4 nm). These nanotubes were
treated according to route 2 synthesis. After chemical treatment, the nanotubes show
bumps on their surface (lower SEM images in Figure 2.28), which may indicate a
successful modification of their surface.

Figure 2.28. Effect of treating “as grown” nanotubes according to route 2 functionalization strategy.
(Top) SEM picture of CVD grown nanotubes on a Si wafer surface. (Bottom) SEM images of the same
nanotubes after chemical treatment according to route 2 synthesis.

61

Comparing Route 1 to Route 2. The investigation of the synthesis of
pentamethylcyclopentadienyl ruthenium sandwich compounds on the sidewalls of carbon
nanotubes led to controversial findings. Two synthetic methods were developed, possibly
leading to two structurally similar but electronically different products, as shown in
Figure 2.29.

Both routes yielded a product whose UV-Vis spectrum has a typical shapeline of
Cp*Ru coordinated to fused-rings arenes. IR spectroscopy confirmed the presence of
Cp*Ru in both products. However, IR spectroscopy is also raising questions about the
product electronic structure and nature of the counter ion to [Cp*Ru]
+
. In both IR spectra,
especially for route 2, the band of absorption for the P-F bond (from PF
6
-
) is relatively
weak, indicating low abundance of this anion in the products. This low abundance of the
PF
6
might be explained by a “negative” six-member

carbon ring

η
6
coordinated to Cp*-

Ru(II)
(b)
Ru(II)
PF
6
(a)

Figure 2.29. Proposed products of the carbon nanotubes treatments with Cp*-Ru precursors according
to (a) route 1 and (b) route 2.

62

Ru, as shown in Figure 2.29 (b). If this is the case, the reaction product Cp*-Ru-nanotube
is neutral, and no counter ions such as Cl
-
or PF
6
-

are necessary to balance to the positive
Cp*-Ru
(II)
moiety. This structure is highly probable for route 2 since this synthetic
procedure uses the sodium salt of reduced nanotubes. However, the actual product might
also be a mixture of the two structures (positive and neutral Cp*-Ru-nanotube), with a
predominance of the second structure (neutral). The proposed product of route 1
synthesis is [Cp*Ru-nanotube][PF
6
] with a neutral carbon nanotube, a cationic Cp*Ru,
and PF
6
as a counter ion, (Figure 2.29 (a)).
Route 1 yielded results that were more consistent with the proposed product than
route 2. In fact, the characterization methods utilized in route 1 (Uv-Vis, IR, and Raman)
supported the structure shown in Figure 2.29(a).

63

2.3 Recent Achievements on the Coordination of RuCp* to Nanotubes
The work described in Section 2.2 was carried out during my 3
rd
year of graduate
studies, and the equipment available at that time to process the nanotubes was not
adequate to properly obtain good dispersions. Moreover, the instruments used to
characterize my products were old, unreliable, or poorly maintained. Recent upgrades in
my lab’s equipment and in the department’s instrumentation facilities allowed for
adequate nanotube processing and product characterization.
Quality Evaluation of the Old Nanotubes by ThermoGravimetric Analysis
(TGA). As a first step in this section, I evaluated the quality of nanotubes used in the
work described in Section 2.2 (the old nanotubes). This evaluation was carried out by
thermogravimetric analysis (TGA), an analytical technique that has been recently
suggested to be a very practical method for the bulk characterization of nanotubes both at
the macroscale (carbon-to-metal ratio) and at the nanoscale (single-walled to multi-
walled ratio).
34
TGA data were collected by burning the nanotube sample under the
following conditions: temperature ramp = 10º C/min; air flow = 50 mL/min. These
conditions were also used for every other TGA analysis described in this Section. The
TGA curve of the old nanotubes is shown in Figure 2.30 as percentage weight loss versus
temperature. The TGA curve shows a significant loss of mass below 400º

C which is
associated with carbonaceous impurities, such as graphite residues, fullerenes, polycyclic
aromatics, and amorphous carbon. Also, the residual mass is significantly high (~35%)
which is associated with impurities related to catalyst nanoparticles. The residual mass is
shown in the photo (the inset in Figure 2.30) as dark gray mass.

64

The differential weight loss versus temperature is shown in Figure 2.31.The
decomposition of amorphous carbon occurs from 200º to 400º C, whereas nanotubes
oxidize above 400º C.
34
This allows the quantitative determination of the amorphous
carbon within the sample. The old nanotubes clearly have a lot of carbon-based
impurities that burn at 350 ºC. The two peaks associated with nanotube decomposition
are centered at 410º C and 435º C, indicating nanotubes of different lengths, rather than
different diameters since Raman showed that the diameter is fairly uniform (Figure 2.10).
Or the 435º C peak maybe belong to the oxidation of double walled- or multiwalled-
nanotubes within the sample.
34

0 100 200 300 400 500 600 700 800
0
10
20
30
40
50
60
70
80
90
100
% Weight Loss
Temperature (C)

Figure 2.30. Thermogravimetric analysis of the old batch of nanotubes used. The TGA curve (left) shows
a significant loss of mass below 400
o
C which is associated with carbonaceous impurities. Also, the
residual mass is significantly high (~35%) which is associated to impurities related to catalysts
nanoparticles. The residual mass is shown in the inset as dark gray mass.

65

Although the work described in Section 2.2 was carried out using low quality and
poorly dispersed nanotubes, that work was very valuable because now I know some
weakness and strengths of the chemistry involved. This time, I have used nanotubes
purchased from Unidym (batch p0261), and had their purity promptly evaluated by TGA.

Unidym Nanotubes: ThermoGravimetric Analysis (TGA). The quality of the
“as-received” nanotubes was evaluated again using TGA. The weight percentage versus
temperature is shown in Figure 2.32(a) and the weight derivative in Figure 2.32(b).
Although the residual metal catalyst is much less than the old nanotubes (~20%),
the Unidym nanotubes contains a significant amount of carbonaceous impurities, which
can interfere with the coordinating Cp*Ru, giving false results. Thus these nanotubes had
to be further purified to eliminate any traces of contaminates such as carbonaceous
species and catalyst metal nanoparticles.
340 360 380 400 420 440 460
0
1
2
3
4
5
6
Differential Weight Loss (%/C)
Temperature (C)
250 300 350 400 450 500
0
1
2
3
4
5
6
Differential Weight Loss (%/C)
Temperature (C)
349
410
435

Figure 2.31. TGA differential weight loss curves. The differential of the TGA curve shown in figure 2.30
is shown on the left as the original set of data points, and on the right after the application of a smoothing
function to increase clarity.

66

Unidym Nanotubes: Bulk Purification. The purification process starts with
annealing the as-received nanotubes in vacuum at 900º C for 1 hour with a slow
temperature ramping (1º C/min).
35
This process removes any carbonaceous contaminants
from the nanotubes,
35
avoiding centrifuging suspensions of nanotubes which would be
incompatible with air-sensitive working conditions.

Purified Unidym Nanotubes: Quality Control by TGA. The TGA curves are
shown in Figure 2.33. Clearly, there is very minimal loss of mass below 400º C,
indicating that any traces of amorphous carbon have been removed. The metal content is
increased to slightly above 20% because amorphous carbon has been eliminated. This
elimination increases the percentage of metal content in the overall sample. The
differential weight curve has been smoothed for clarity.
100 200 300 400 500 600 700 800
0
10
20
30
40
50
60
70
80
90
% Weight Loss
Temperature (C)
100 200 300 400 500 600 700
-0.1
0.0
0.1
0.2
0.3
0.4
0.5
Derivative Weigh Loss (%/C)
temperature (C)

Figure 2.32. TGA of as-purchased nanotubes. The loss of mass below 400 C indicates the presence of
amorphous carbon, impurities that may highly interfere with my chemistry. The metal content is less than
20%. The differential curve has been smoothed.

67

Raw and Purified Unidym Nanotubes: Raman Spectroscopy. Raman
spectroscopy was used to monitor the efficacy of the purification process. The Raman
spectrum of the as-received (red trace) and purified (blue trace) nanotubes is shown in
Figure 2.34. Panel (a) shows the entire spectrum, panel (b) focuses on the radial breathing
mode region, panel (c) focuses on the D and G lines, and panel (d) focuses on the G’ line.
Both spectra were collected using a 532 nm laser and the resulting traces have been
normalized at the intensity of the G’ line.
35
Interestingly, the radial breathing mode shows
at least four distinct peaks (229, 237, 247, and 274 nm), indicating the presence of
nanotubes with different diameters within the sample. The diameter of these nanotubes
has been calculated to be 1.1, 1.05, 1, and 0.9 nm, respectively. These diameters are
slightly smaller than the old nanotubes (1.4 nm). This could be a disadvantage for the
coordination of Cp*-Ru since the curvature is smaller.
0 100 200 300 400 500 600 700 800
0
10
20
30
40
50
60
70
80
90
100
% Weight Loss
Temperature (C)
350 400 450 500
0
5
10
15
20
25
Differential Weight Loss (%/C)
Temperature (C)
413
401
435

Figure 2.33. TGA of purified nanotubes. There is no loss of mass below 400 C indicating that traces of
amorphous carbon have been removed. The metal content is slightly above 20%, because amorphous
carbon has been eliminated thus increasing the % of metal content. The differential curve has been
smoothed for clarity.

68

A small G/G’ line ratio is an indication of successful purification.
35
My
purification process produced a G/G’ line ratio smaller than that of raw nanotubes (panel
(c)), a clear sign of the elimination of impurities. Also, this broader G line shows a strong
shoulder at 1540 cm
-1
, characteristic of electron-phonon coupling in bundles of
nanotubes. This shoulder also indicates the presence of a large fraction of metallic
nanotubes in my sample.
-500 0 500 1000 1500 2000 2500 3000
0.0
0.2
0.4
0.6
0.8
1.0
1.2
1.4
1.6
Normalized Counts
Wav enumber (1/cm)
(a)
100 150 200 250 300 350 400
0.00
0.05
0.10
0.15
0.20
0.25
0.30
Normalized Counts
Wav enumber (1/cm)
274
229
237
250
(b)

1200 1300 1400 1500 1600 1700 1800
0.0
0.2
0.4
0.6
0.8
1.0
1.2
1.4
1.6
Normalized Counts
Wav enumber (1/cm)
1330
1520
1541
1590
(c)
2200 2300 2400 2500 2600 2700 2800 2900 3000
0.0
0.2
0.4
0.6
0.8
1.0
Normalized Counts
Wav enumber (1/cm)
2634
(d)

Figure 2.34. The Raman spectra of as-received (red trace) and purified (blue trace) Unidym nanotubes.
Panel (a) shows the entire spectrum, panel (b) focuses on the radial breathing mode region, panel (c)
focuses on the D and G lines, and panel (d) on the G’ lines. Both traces have been normalized at the
intensity of the G’ line.

69

Purified Unidym Nanotubes: UV-Vis Spectroscopy. Annealed nanotubes were
placed in a round bottom flask and heated under vacuum for 5 minutes to remove
moisture and other absorbed molecules. Dichloroethane was added in the ratio of 1 ml of
DCE/0.05 mg of nanotubes. DCE was previously distilled over CaH
2
to remove traces of
water and stored in a Schlenk flask. The nanotubes were then sonicated in a cup horn
ultrasonicator for 30 minutes at the power of 80 W. The sonicator cup was filled with a
circulating mixture of water/ethylene glycol (50/50) chilled at 0 C with a chiller. The
efficacy of this dispersion procedure is evaluated by UV-Vis spectroscopy. The
suspended nanotube spectrum is shown in Figure 2.35 in the 260-1560 nm range.

Selected regions of this spectrum are shown in Figure 2.36, where the nanotube
electronic transitions are easier to visualize. The intensity of these transitions is lower
than those produced by nanotubes suspended in water with the aid of surfactants, which
can suspend nanotubes at a concentration at least 20 times higher than dry DCE.
400 600 800 1000 1200 1400 1600
0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
Abs (arb units)
w avelength (nm )

Figure 2.35. UV-Vis-nIR spectrum of suspended nanotubes in dry DCE.

70

Coordination of Cp*Ru to Nanotubes. The coordination of Cp*-Ru to carbon
nanotubes was carried out according to Route 1, as described in Section 2.2, but with a
few modifications. Highly purified nanotubes can be easily suspended in dry DCE, but at
lower concentrations (less than 0.05 mg/mL) than the old nanotubes, where carbonaceous
impurities may have favored the suspension. A typical suspension is obtained by
sonicating 1 mg of nanotube soot (~80% pure by TGA) in 20 mL of dry DCE. I know
define 1 equivalent of nanotube to be 30 carbon atoms in the nanotube. Computer
modeling shows that Cp*Ru moieties can coordinated to a nanotube with a “footprint” of
24-30 carbon atoms, before bumping into each other. Therefore, my nanotube suspension
was about 100 µM in nanotubes (~2 µmol in 20 mL). Addition of 1 equivalent Cp*-Ru-
(MeCN)
3
increases the ionic strength of the DCE suspension, and the nanotubes slowly
re-aggregate. Addition of fresh DCE (10 mL more) followed by 30 seconds bath
500 550 600 650 700 750
0.14
0.16
0.18
0.20
0.22
0.24
0.26
Abs (arb units)
w av elength (nm)
M1 1
S22
1100 1200 1300 1400
0.06
0.07
0.08
0.09
0.10
Abs (arb units)
wav elength (nm)
S11

Figure 2.36. Selected regions of the UV-Vis-nIR spectrum of nanotubes shown in Figure 3.35. The left
panel focuses on the 470-800 nm region where the M11 and S22 absorptions take place. The right panel
focuses on the semiconducting fine structures (S11) in the 1050-1450 nm region.

71

sonication separates the nanotubes once again, and keeps them suspended. The rest of the
synthesis procedure is as described in Section 2.2.

Product characterization: UV-Vis Spectroscopy. All the UV-Vis spectra were
collected in the 260-1560 nm range in dry DCE, using the same cuvettes and the same
instrument. The entire collected spectra are shown in Figure 2.37, and a zoom of the 260-
650 nm region is presented in Figure 2.38. Spectrum (a), red trace, belongs to the reaction
product, Cp*-Ru-nanotube. Spectrum (b), green trace, is the absorption curve of
suspended nanotubes. Spectrum (c), blue trace, belongs to the reagent Cp*-Ru-(MeCN)
3
.
Spectrum (d), wine trace, was recorded from a solution of Cp*-Ru-Pyrene.
400 600 800 1000 1200 1400
0.0
0.5
1.0
1.5
Abs (arb units)
wav elength (nm)
(a)
(b)
(c)
(d)

Figure 2.37. UV-Vis spectroscopy product characterization. All spectra have been recorded in the 260-
1560 nm range using the same instrument, and in the same solvent (dry DCE). (a, Red) Spectrum of the
reaction product Cp*-Ru-nanotube. (b, Green) Spectrum of suspended nanotubes. (c, Blue) Spectrum of
the reagent Cp*-Ru-(MeCN)
3
. (d, Wine) Spectrum of Cp*-Ru-Pyrene.

72

Distinctive bands of absorption in spectrum (a) are given by the Cp* moiety
located at 315 nm, a nanotube transition at 370 nm, and the Cp*Ru coordinated to the
nanotube at 466 nm. The band of Ru-Cp* is lower in intensity with respect to Figure
2.16(b), and also red-shifted by 40 nm. The position of λ
max
is significantly different from
the old nanotubes ( λ
max
= 415 nm) and the pyrene analog ( λ
max
= 360). However, the
naphthalene analog has a λ
max
at 480 nm (Table 2.1). Thus, it is not unusual for Cp*Ru-
arenes to have such a red-shifted λ
max
.

These time differences in the UV-Vis spectrum
with respect to the old nanotubes have several origins. Beside the fact that the two spectra
250 300 350 400 450 500 550 600 650
0.0
0.2
0.4
0.6
0.8
1.0
1.2
Abs (arb units)
wav elength (nm)
315
358
415
466
(a)
(b)
(c)
(d)
370

Figure 2.38. Detail of the UV-Vis spectra shown in Figure 2.37. (a, Red) Spectrum of the reaction
product Cp*-Ru-nanotube. This spectrum shows a new broad absorption band centered at 466 nm. This
band is absent is all the other spectra. (b, Green) Spectrum of suspended nanotubes. (c, Blue) Spectrum
of the reagent Cp*-Ru-(MeCN)
3
. (d, Wine) Spectrum of Cp*-Ru-Pyrene.

73

were collected using different instruments and different solvents, the nanotubes used for
spectrum 2.16(b) contained a significant amount of impurities, such as polycyclic
aromatics that may have also coordinated Cp*-Ru. Moreover, the Unidym nanotubes
have a much smaller diameter than the old nanotubes, which affects the nanotube
electronic structure.

Product characterization: Raman Spectroscopy. The spectrum of the reaction
product is shown in Figure 2.39 as a black trace, and is juxtaposed to the purified
nanotube spectrum (blue trace). Panel (a) shows the entire spectrum, panel (b) focuses on
the radial breathing mode (RBM) region, panel (c) focuses on the D and G lines, and
panel (d) on the G’ line. Both spectra were collected using a 532 nm laser and the
resulting traces have been normalized at the intensity of the G’ line
35
in panels (a), (c),
and (d), and to the tallest RBM in panel (b).
The radial breathing mode of the modified nanotubes--black trace in panel (b)--is
significantly lower in intensity, indicating a decrease in symmetry on the nanotube cross
section. This is an indication of successful surface modification. Panel (c) shows a
narrowing of the G line, a phenomenon correlated with electronic doping of the
nanotubes. Panel (d) is shown in detail in Figure 2.40. The position rather than the
intensity of the G’ is an indication of electronic doping.
35
My modified nanotubes have a
G’ line that has shifted by 5 wavenumbers to higher energies, an indication of electronic
doping.
35
The asymmetrical shape of the G’ line indicates that these nanotubes have been
functionalized to different degrees. This non-uniform functionalization is expected since

74

this sample contains nanotubes with at least four different diameters and a number of
different chiralities, which in turn possess different reactivity.

500 1000 1500 2000 2500 3000
0.0
0.2
0.4
0.6
0.8
1.0
1.2
Normalized Counts
Wav enumber (1/cm)
(a)
150 200 250 300 350
0.0
0.2
0.4
0.6
0.8
1.0
Normalized Counts
Wav enumber (1/cm)
(b)

1200 1300 1400 1500 1600 1700 1800 1900 2000 2100
0.0
0.2
0.4
0.6
0.8
1.0
Normalized Counts
Wav enumber (1/cm)
(c)
2450 2500 2550 2600 2650 2700 2750 2800 2850 2900 2950 3000 3050
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
1.0
Normalized Counts
Wav enumber (1/cm)
(d)

Figure 2.39. The Raman spectra of purified Unidym nanotubes (blue trace) and the Cp*-Ru modified
nanotubes (black trace). Panel (a) shows the entire spectrum, panel (b) focuses on the radial breathing
mode region, panel (c) focuses on the D and G line, and panel (d) on the G’ line. Both traces have been
normalized at the intensity of the G’ line in panels (a), (c), and (d), and to the tallest RBM in panel (b).

75

2550 2600 2650 2700 2750
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
1.0
Normalized Counts
Wav enumber (1/cm)
2634
2640

Figure 2.40. Selected region of Raman spectra of purified Unidym nanotubes (blue trace) and the Cp*-
Ru modified nanotubes (black trace) showing the G’ line.

76

2.4 Covalent Controlled Surface Modification
Background. Networks of carbon nanotubes in the nanobiosensing project are
used as semiconducting channels in FET devices (See Chapters One and Five). In order
to have optimally performing nanobiosensors, it is important to eliminate every metallic
pathway in the FET channel. In addition, to produce more durable/stable sensors
(especially for nucleic acid sensors) it is important to covalently attach the capture probe
to the surface of the nanotubes. This experiment, involving diazonium salt, was designed
to address both challenges affecting these nanotube-based devices. At the time I was
working on this experiment, metallic pathways were eliminated by electrical breakdown
(see below). The surface modification was either achieved using non-covalent linkers,
such as pyrene derivatives or Tween 20.
As discussed in Section 2.1, the surface functionalization of carbon nanotubes
using chemical reagents is still a challenge. When modifying FET devices, the ideal
surface modification method would use a reagent that:
1) Distinguish between metallic and semiconducting nanotubes.
2) Covalently bind to the nanotube, but preserve most of the sp
2
network so the
electronic properties are only slightly affected.
One method we initially explored involved using the nanotube’s defect sites to
covalently attach linker molecules to the nanotube sidewalls. These defect sites are
created during CVD growth of nanotubes and are often pending methyl groups that can
be easily oxidized to COOH by nitric acid treatment. The carboxylic moiety can then be
used as an anchoring site to attach other molecules via standard coupling chemistry.

77

However, these defect sites are not as common on the nanotube as we would like.
Therefore this technique can yield only a low degree of functionalization. We then
explored using oxidized defect sites obtained by oxygen plasma treatment. This treatment
utilizes highly reactive oxygen plasma species, mainly O
2
+
, to oxidize the nanotube
sidewalls, creating a number of functional groups, including carboxylic acids, aldehydes,
ketones, and alcohols. In order to create a COOH group, several C=C need to be broken
and often a hole is created in the nanotube, as shown in Figure 2.41. This hole greatly
affects the electronic properties of the nanotube. This method is fast and can be applied to
a large number of devices simultaneously. However, when we explored this method, it
was difficult to obtain the optimal degree of oxidation and often devices were destroyed.
A method to remove metallic pathways in a device requires the electrical
breakdown of metallic nanotubes. This electrical breakdown is accomplished by
increasing the current between source and drain electrodes while the gate electrode is

Figure 2.41. Oxidized carbons on the nanotube sidewall. Oxidized defect sites can be obtained by
oxygen plasma treatment. This method yields a number of functional group, including carboxylic
acids, aldehydes, ketones, and alcohols. This oxidation breaks several C=C can create a hole in the
nanotube, affecting the electronic properties of the nanotube.

78

used to maintain the device in an off state by holding the gate potential at a fixed positive
high voltage. In this FET configuration, the semiconductive pathways are non-conductive
and thus the current only passes through the metallic pathways, heating the nanotubes
until they burn off. This method is slow but effective at improving the On/Off ration of
nanotube devices, as shown in Figure 2.42. However, this method is also known to
damage adjacent semiconducting nanotubes. I have used this method to improve the
On/Off ratio of nanotube devices used as biosensors, but I started to explore the
possibility of using diazonium salts following the work of Professor Klaus Kern
36, 37
(see
below). Recently, our team, in particular Fumiaki Ishikawa demonstrated that by
carefully tuning the surface density of nanotubes in the FET channel it is possible to
obtain semiconducting nanotube networks where metallic pathways are very improbable.

(a)
(b)
(c)

Figure 2.42. Electrical breakdown of metallic pathways nanotube-based FETs. (a) Current (log scale)
versus voltage for a nanotube FET (Vds = +500mV). The On/Off ration is close to one, indicating the
presence of metallic pathways. (b) The same device exhibits an improvement in the On/Off ratio after
electrical breakdown. (c) Plot of drain current versus source-drain voltage showing a typical curve for
the electrical breakdown on metallic metallic nanotubes (Vg = +30 V).

79

2.4.a. Diazonium Salts Chemistry on Carbon Nanotubes
The electrochemically controlled addition of diazonium salts is a method that
allows the surface modification of nanotubes directly on the FET device.
36, 37
In this
method, a nanotube–based FET is used as the working electrode in an electrochemical
cell. A reducing voltage applied to the FET (~-250 mV) reduces the diazonium salt,
creating arene radical anions that add to the nanotube C=C, as illustrated in Figure 2.43.

This method was investigated because it can yield three results that are useful to
biosensing with a single pot:
1) Remove metallic pathways in nanotube-based FETs.
2) Provide an anchoring point for the covalent immobilization of capture probes in a
controlled way so the electrical characteristics of the device will be unaltered. In fact,

Figure 2.43. Reaction mechanism for the addition of a nitrobenzene radical anion to a carbon nanotube.
This nitrobenzene radical anion is formed in situ from the electrochemical reduction of a para-
nitrobenzene diazonium salt.

80

C-C sigma bonds are not broken during the addition of diazonium derivatives, as
shown in Figure 2.44.
3) The number of linker molecules can be controlled by optimizing the applied voltage
(in an electrochemical cell), concentration of diazonium, and time.

2.4.b. Preliminary Investigation of Diazonium Chemistry on Nanotubes
Initial testing was carried out on buckypapers, which are tangible models for the
nanotube networks in FET devices. These nanotube based papers are conductive and thus
can be used as working electrodes in an electrochemical cell. The buckypapers were
fabricated by an undergraduate student working under my supervision, Kunal Patel.
Buckypapers were supported by a substrate such as a plastic sheet or glass due to their

Figure 2.44. Molecular modeling showing the addition of one (left) or two (right) molecules of
nitrobenzene radical anion to the C=C bond of a carbon nanotube. The addition of nitrobenzene
preserve the C-C sigma bond, and only affect the C=C pi bond.

81

fragility. Notably, the conductivity of these nanotube films varies significantly from day
to day and is not precisely reproducible.
Addition of diazonium salts to buckypapers was simple and straightforward. The
buckypaper is assembled in a custom-designed holder, and a Teflon cone-shaped cell is
placed on top of the buckypaper and sealed at the bottom. The Teflon cell is filled with a
1 mM solution of para-nitrobenzene diazonium salt in PBS buffer. The buckypaper is
clamped with a clip alligator and connected to the potentiostat (working electrode port);
platinum gauze was used as counter electrode; and a standard Ag/AgCl electrode was the
references. A reductive voltage of -250 mV applied for 10 seconds resulted in the
addition nitrobenzene radicals to the nanotubes.
The conductivity of nitrobenzene-modified nanotube films was characterized by
an electrochemical method. Initially, the buckypaper is a conductive material and can be
used to observe the redox signal of [Fe(CN)
6
]
3-
. The Teflon cell was filled with a 0.01
mM solution of [Fe(CN)
6
]
3-
in PBS buffer and the redox waves were recorded, as shown
in Figure 2.45(a). The Fe(III) solution was extensively washed away and replaced by the
1 mM solution of para-nitrobenzene diazonium salt in PBS buffer. Ten seconds at -250
mV ensured the completed addition of nitrobenze to the nanotubes. Extensive
functionalization of nanotubes disrupts the sp
2
network, causing the nanotubes to become
an insulating material. The buckypaper thus becomes non-conductive and therefore
cannot serve as a working electrode any longer. The Teflon cell is refilled with the Fe(III)
solution, but this time it is not possible to observe its redox signal, since the modified
nanotubes are non-conductive, producing a flat CV trace, Figure 2.45(b).

82

While this procedure to treat nanotube networks in FET devices generated
encouraging preliminary results, we halted the investigation before attempting anything
on nanotube devices. At that time, our team developed a method of obtaining
semiconducting networks of nanotubes by carefully tuning the surface density of
nanotubes in the FET channel. Moreover, the non-covalent functionalization of nanotubes
gives reliable results in terms of stability for immobilized protein capture probes.

-800 -600 -400 -200 0 200 400 600 800 1000 1200
-250
-200
-150
-100
-50
0
50
100
150
Current (micro Amps)
Potential Vs Ag/AgCl (mV)
(a)
(b)

Figure 2.45. Effect of the diazonium-modification on the conductivity of a buckypaper. (a) An
unmodified buckypaper is conductive and can be used to observe the redox waves of [Fe(CN)
6
]
3-
. After
the treatment with a diazonium salt, the buckypaper becomes insulating, thus does not function as
working electrode, and the redox waves of [Fe(CN)
6
]
3-
cannot longer be observed (flat CV trace).

83

2.5 Selective Functionalization of Carbon Nanotubes
Background. The selective functionalization of carbon nanotube-based devices
has not yet been reported in the literature. The rationale and background for the selective
functionalization is discussed in Chapter Three for indium oxide nanowires, and the same
concepts apply to the selective functionalization of nanotube devices.
Just as the preliminary investigation for nanowires was carried out on model
surfaces (ITO coated glass), so the preliminary investigation for nanotubes was carried
out on buckypapers. Buckypapers are thin films of carbon nanotubes, often supported by
a substrate such as a plastic sheet or glass. These buckypapers can be used as working
electrodes in electrochemical cells. Notably, the conductivity of these films varies
significantly from day to day and is not easily reproducible. The electrochemically
switchable molecule designed for this experiment is 1-(4-(2,5-dimethoxyphenyl)-butyl)-
pyrene (in short, DMP-pyrene) and is shown in Figure 2.46. This bifunctional molecule
binds to the nanotube surface by Pi-Pi interaction via the pyrene moiety and the other
terminal entails of the electrochemically active hydroquinone (protected by methoxy
groups in this case). Initially, the deprotection of hydroquinone was slated to be
chemically carried out using BBr
3
or cerium(IV) ammonium nitrite (CAN). Ultimately
however, this deprotection was achieved electrochemically by applying an oxidizing
voltage of +1.5V at the working electrode, following a procedure proposed by my
colleague Rui Zhang.

84

A solution of DMP-pyrene in methanol (0.1 mM) is prepared with the aid of
sonication and gentle heating. The buckypaper is assembled in a custom-designed holder,
and a Teflon cone-shaped cell is placed on top of the buckypaper and sealed at the
bottom. The solution containing DMP-pyrene is placed in the cell and allowed to sit for
over 1 hour so the pyrene derivative would self assemble on the nanotubes. After
extensive washing, the Teflon cell is filled with PBS buffer and the buckypaper is
clamped with a clip alligator and connected to the potentiostat (working electrode port).
The cyclicvoltammetry traces of the DMP-pyrene modified nanotubes (red) and
bare buckypaper (blue) are shown in Figure 2.47. The red CV trace shows the typical
redox waves of hydroquinone derivatives. A control scan performed using bare nanotubes
shows a featureless CV trace.
After having demonstrated the functionality of this technology using buckypapers,
this investigation was never carried over to actual nanotube devices.

Figure 2.46. The Bifunctional, electrochemically active molecule that can be used to selectively
functionalize carbon nanotube-based electrodes/devices.

85

2.5.a Synthesis of DMP-pyrene
The overall synthetic procedure used to synthesize DMP-pyrene is shown in
Figure 2.48. A solution of 1-bromo-pyrene (590 mg, 2.1 mmol) is dry ether was cooled to
-78 C in a dry ice/acetone bath. Two equivalents of tert-butyl lithium (3 mL of 1.44 M
solution in hexane) were added drop wise, under nitrogen. Within seconds, the reaction
mixture turned orange. The mixture was stirred at -78 C for 20 minutes, the mixture
turned dark orange/red. A solution of 2-(4-Bromobuthyl)-1,4-dimethoxybenzene (0.751
mg, 0.75 mmol) in dry ether was added to 1-lithium-pyrene solution. The synthesis of
-400 0 400 800 1200 1600
-100
-50
0
50
100
150
Current (micro Amps)
Potential (mV)

Figure 2.47. Cyclic voltammetry traces a buckypaper (blue) and a buckypaper fnctionalized with
DMP-pyrene (red). The CV of the buckypaper is essentially featureless as no redox active functional
groups are present in the nanotube paper. Upon binding DMP-pyrene, the CV trace shown the typical
redox waves of hydroquinone/paraquinone derivatives.

86

this molecule is detailed in Section 3.3. This mixture was allowed to warm up to room
temperature while stirring and stirred for 1 more hour. The reaction mixture was first
quenched with methanol and then with 6 M ammonium chloride. Extraction with ether
and purification on a column chromatography (Hexane/ethyl acetate, 6:4) yielded DMP-
pyrene as sticky mass.

2.6. Conclusions
In this chapter I have discussed the investigation of novel methods for the surface
modification of carbon nanotubes for potential applications in FET devices, such as
nanobiosensors. Carbon nanotubes are a very promising material for sensing applications
since every atom in the material is at the surface, but their use is limited due to drawbacks
such as the presence of metallic nanotubes and challenging surface chemistry and doping.
I have proposed and explored two surface modification methods that address the above
mentioned problems.
The first method I proposed and investigated is a chemical method that results in
the coordination of Cp*-Ru to the fused, six-member carbon rings of the nanotube. This
Br
2eq. t-BuLi
-78
o
C, ether
Li
MeO OMe
Br
MeO OMe
ether

Figure 2.48. Synthesis of 1-(4-(2,5-dimethoxyphenyl)butyl) pyrene.

87

method provides a technique to covalently modify the nanotube sidewalls without
breaking any of the nanotube C=C (thus preserving the conjugation system), and at the
same time n-dope the semiconducting nanotubes via electron donation from the
coordinated Cp*-Ru. This project was highly innovative since metal sandwich
compounds using nanotubes have never been reported before. Two synthetic routes were
investigated to make the Cp*Ru-nanotube complex. Both routes yielded a product whose
UV-Vis spectrum has a typical shapeline of Cp*Ru coordinated to fused-rings arenes. IR
spectroscopy confirmed the presence of Cp*Ru in both products. However, IR
spectroscopy is also raising questions about the product electronic structure and nature of
the counter ion to [Cp*Ru]
+
. In both IR spectra, especially for route 2, the band of
absorption for the P-F bond (from PF
6
-
) is relatively weak, indicating low abundance of
this anion in the products. This low abundance of the PF
6
might be explained by a
“negative” six-member

carbon ring

η
6
coordinated to Cp*-Ru. If this is the case, the
reaction product Cp*-Ru-nanotube is neutral, and no counter ions such as Cl
-
or PF
6
-

are
necessary to balance to the positive Cp*-Ru
(II)
moiety. This structure is highly probable
for route 2 since this synthetic procedure uses reduced nanotubes. However, the actual
product might also be a mixture of the two structures (positive and neutral Cp*-Ru-
nanotube), with a predominance of the second structure (neutral). The proposed product
of route 1 synthesis is [Cp*Ru-nanotube][PF
6
] with a neutral carbon nanotube, a cationic
Cp*Ru, and PF
6
as a counter ion. The expected n-doping effect of the Cp*Ru on the
electrical characteristics of nanotube FET was not further investigated.

88

The Cp*Ru coordination to nanotubes was recently resumed using nanotubes of
higher purity. Individual nanotube suspensions were achieved with the aid of an
ultrasonicator. Cp*Ru coordination was accomplished according to route 1 and the
products characterized by UV-Vis-nIR and Raman spectroscopy. Both characterization
techniques confirmed the coordination of Cp*Ru to the nanotubes.
A second method to modify the nanotube surface utilizes the electrochemical-
mediated addition of arene radicals (generated in situ from reduction of diazonium
precursors) to the nanotube C=C. This method was investigated for two reasons. First,
provides us to destroy metallic pathways in the nanotube network in the FET channel
after drastic derivatization using radicals generated from precursors such as para-
diazonium salt of nitrobenzene. Second, this method offers an attachment points for
bioconjugation after controlled derivatization using a precursor such as the para-
diazonium salt of benzoic acid. Nanotube-based biosensors modified as described above
have never been reported before. The addition of diazonium salts was demonstrated on
buckypapers, but was not further investigated on nanotube FET devices.
A related project involved investigating an electrochemical method for the
selective functionalization of dense arrays of nanotube-based FET devices. A
hydroquinone derivative of pyrene was designed and synthesized, inspired by
achievements in nanowire surface modification (see Chapter Three). The functionality of
this switchable molecule was demonstrated on buckypapers, but was not further
investigated on nanotube FET devices.

89

2.7 Chapter 2 References
1. Iijima, S., Helical Microtubules of Graphitic Carbon. Nature 1991, 354, (6348),
56-58.
2. Avouris, P.; Chen, J., Nanotube electronics and optoelectronics. Materials Today
2006, 9, (10), 46-54.
3. Salvetat, J. P.; Bhattacharyya, S.; Pipes, R. B., Progress on mechanics of carbon
nanotubes and derived materials. Journal of Nanoscience and Nanotechnology
2006, 6, (7), 1857-1882.
4. Endo, M.; Hayashi, T.; Kim, Y. A.; Muramatsu, H., Development and application
of carbon nanotubes. Japanese Journal of Applied Physics Part 1-Regular Papers
Brief Communications & Review Papers 2006, 45, (6A), 4883-4892.
5. Hu, Y. H.; Shenderova, O. A.; Hu, Z.; Padgett, C. W.; Brenner, D. W., Carbon
nanostructures for advanced composites. Reports on Progress in Physics 2006,
69, (6), 1847-1895.
6. Trojanowicz, M., Analytical applications of carbon nanotubes: a review. Trac-
Trends in Analytical Chemistry 2006, 25, (5), 480-489.
7. Anantram, M. P.; Leonard, F., Physics of carbon nanotube electronic devices.
Reports on Progress in Physics 2006, 69, (3), 507-561.
8. Lam, C. W.; James, J. T.; McCluskey, R.; Arepalli, S.; Hunter, R. L., A review of
carbon nanotube toxicity and assessment of potential occupational and
environmental health risks. Critical Reviews in Toxicology 2006, 36, (3), 189-217.
9. Moulton, S. E.; Minett, A. I.; Wallace, G. G., Carbon nanotube based electronic
and electrochemical sensors. Sensor Letters 2005, 3, (3), 183-193.
10. Avouris, P., Molecular electronics with carbon nanotubes. Accounts of Chemical
Research 2002, 35, (12), 1026-1034.
11. Lee, M.; Baik, K. Y.; Noah, M.; Kwon, Y. K.; Lee, J. O.; Hong, S., Nanowire and
nanotube transistors for lab-on-a-chip applications. Lab on a Chip 2009, 9, (16),
2267-2280.
12. Byon, H. R.; Kim, S.; Choi, H. C., Label-Free Biomolecular Detection Using
Carbon Nanotube Field Effect Transistors. Nano 2008, 3, (6), 415-431.

90

13. Patra, M. K.; Manzoor, K.; Manoth, M.; Negi, S. C.; Vadera, S. R.; Kumar, N.,
Nanotechnology applications for chemical and biological sensors. Defence
Science Journal 2008, 58, (5), 636-649.
14. Sinha, N.; Ma, J. Z.; Yeow, J. T. W., Carbon nanotube-based sensors. Journal of
Nanoscience and Nanotechnology 2006, 6, (3), 573-590.
15. Popov, A. M.; Lozovik, Y. E.; Fiorito, S.; Yahia, L., Biocompatibility and
applications of carbon nanotubes in medical nano robots. International Journal of
Nanomedicine 2007, 2, (3), 361-372.
16. Lacerda, L.; Bianco, A.; Prato, M.; Kostarelos, K., Carbon nanotubes as
nanomedicines: From toxicology to pharmacology. Advanced Drug Delivery
Reviews 2006, 58, (14), 1460-1470.
17. Hussain, M. A.; Kabir, M. A.; Sood, A. K., On the cytotoxicity of carbon
nanotubes. Current Science 2009, 96, (5), 664-673.
18. Peng, X. H.; Wong, S. S., Functional Covalent Chemistry of Carbon Nanotube
Surfaces. Advanced Materials 2009, 21, (6), 625-642.
19. Singh, S.; Kruse, P., Carbon nanotube surface science. International Journal of
Nanotechnology 2008, 5, (9-12), 900-929.
20. Banerjee, S.; Hemraj-Benny, T.; Wong, S. S., Covalent surface chemistry of
single-walled carbon nanotubes. Advanced Materials 2005, 17, (1), 17-29.
21. Jin, W. J.; Sun, X. F.; Wang, Y., Solubilization and functionalization of carbon
nanotubes. New Carbon Materials 2004, 19, (4), 312-318.
22. Tasis, D.; Tagmatarchis, N.; Georgakilas, V.; Prato, M., Soluble carbon
nanotubes. Chemistry-a European Journal 2003, 9, (17), 4001-4008.
23. Terrones, M.; Souza, A. G.; Rao, A. M., Doped carbon nanotubes: Synthesis,
characterization and applications. Carbon Nanotubes 2008, 111, 531-566.
24. Tans, S. J.; Verschueren, A. R. M.; Dekker, C., Room-temperature transistor
based on a single carbon nanotube. Nature 1998, 393, (6680), 49-52.
25. Chaturvedi, P.; Verma, P.; Singh, A.; Chaudhary, P. K.; Harsh; Basu, P. K.,
Carbon nanotube - Purification and sorting protocols. Defence Science Journal
2008, 58, (5), 591-599.

91

26. Pillai, S. K.; Ray, S. S.; Moodley, M., Purification of single-walled carbon
nanotubes. Journal of Nanoscience and Nanotechnology 2007, 7, (9), 3011-3047.
27. Gomez, L. M.; Kumar, A.; Zhang, Y.; Ryu, K.; Badmaev, A.; Zhou, C. W.,
Scalable Light-Induced Metal to Semiconductor Conversion of Carbon
Nanotubes. Nano Letters 2009, 9, (10), 3592-3598.
28. Harutyunyan, A. R.; Chen, G. G.; Paronyan, T. M.; Pigos, E. M.; Kuznetsov, O.
A.; Hewaparakrama, K.; Kim, S. M.; Zakharov, D.; Stach, E. A.; Sumanasekera,
G. U., Preferential Growth of Single-Walled Carbon Nanotubes with Metallic
Conductivity. Science 2009, 326, (5949), 116-120.
29. Schrenk, J. L.; Mann, K. R., Photochemical Studies of [(Eta-5-C5r5)M(Eta-6-
Cot)]Pf6 and [(Eta-5-C5r5)M(Eta-4-1,5-Cot)L]Pf6 Complexes of Iron and
Ruthenium. Inorganic Chemistry 1986, 25, (11), 1906-1908.
30. Angelici, R. J.; Zhu, B.; Fedi, S.; Laschi, F.; Zanello, P., Electrochemical and
EPR studies of the corannulene ruthenium(II) sandwich complex [(eta(6)-
C6Me6)Ru(eta(6)-C20H10)](SbF6)(2). Inorganic Chemistry 2007, 46, (25),
10901-10906.
31. Vecchi, P. A.; Alvarez, C. M.; Ellern, A.; Angelici, R. J.; Sygula, A.; Sygula, R.;
Rabideau, P. W., Synthesis and structure of a dimetallated buckybowl:
Coordination of one {Cp*Ru}(+) unit to each side of corannulene. Angewandte
Chemie-International Edition 2004, 43, (34), 4497-4500.
32. Zhu, B. L.; Ellern, A.; Sygula, A.; Sygula, R.; Angelici, R. J., eta(6)-coordination
of the curved carbon surface of corannulene (C20H10) to (eta(6)-arene)M2+ (M =
Ru, Os). Organometallics 2007, 26, (7), 1721-1728.
33. Vecchi, P. A.; Alvarez, C. M.; Ellern, A.; Angelici, R. J.; Sygula, A.; Sygula, R.;
Rabideau, P. W., Flattening of a curved-surface buckybowl (corannulene) by
eta(6) coordination to {Cp*Ru}(+). Organometallics 2005, 24, (19), 4543-4552.
34. Mansfield, E.; Kar, A.; Hooker, S. A., Applications of TGA in quality control of
SWCNTs. Analytical and Bioanalytical Chemistry 2010, Article in Press, DOI
10.1007/s00216-009-3319-2.
35. Moonoosawmy, K. R.; Kruse, P., To Dope or Not To Dope: The Effect of
Sonicating Single-Wall Carbon Nanotubes in Common Laboratory Solvents on
Their Electronic Structure. Journal of the American Chemical Society 2008, 130,
(40), 13417-13424.

92

36. Balasubramanian, K.; Burghard, M.; Kern, K., Effect of the electronic structure of
carbon nanotubes on the selectivity of electrochemical functionalization. Physical
Chemistry Chemical Physics 2008, 10, (16), 2256-2262.
37. Kooi, S. E.; Schlecht, U.; Burghard, M.; Kern, K., Electrochemical modification
of single carbon nanotubes. Angewandte Chemie-International Edition 2002, 41,
(8), 1353-1355.

93

CHAPTER 3:
SURFACE MODIFICATION OF INDIUM OXIDE NANOWIRES

Chapter Outline
3.1. Indium Oxide Nanowires: Synthesis And Electronic Properties
3.2. Surface Modification
3.3. Selective Functionalization of In
2
O
3
Nanowire-Based FET Devices
3.4. Recent Achievements in the Selective Functionalization of In
2
O
3

3.5. Conclusions
3.6. Chapter 3 References

3.1. Indium Oxide Nanowires: Synthesis and Electronic Properties
Indium oxide nanowires are synthesized using the laser ablation-assisted
technique, which promotes growth according to the Vapor-Liquid-Solid (VLS)
mechanism.
1-4
In this procedure, gold nanoparticles with a mean diameter of 9.4 nm are
used as catalysts for the VLS growth. These Au nanoparticles are uniformly dispersed,
from a colloidal aqueous solution, onto the substrate (usually a piece of Si wafer) to be
used as support for the nanowire growth. Theoretically, these nanowires could be grown
on any substrate that can sustain the high temperatures used during growth (770
o
C).
After the catalyst deposition, the substrate is washed, dried with a stream of air, and
inserted into a quartz tube placed inside a furnace. A schematic diagram of this set up is
94

shown in Figure 3.1. A source of indium, typically a small piece of InAs, is also placed
inside the quartz tube. The internal temperature of the furnace is rapidly raised to 770
o
C.
Then, the InAs piece is atomized by laser ablation using a laser (458 nm; 130 mW).
The atomized indium is carried over the catalyst by a stream of inert gas, such as
argon, containing about 40 ppm of oxygen. When the atomized indium reaches the
catalyst nanoparticles, the gold and indium fuse forming an In/Au alloy. Continuous
feeding of such alloy with indium causes the molten solution to reach saturation; and
each crystalline indium oxide grows in only one direction from the catalyst nanoparticle,
as clearly shown by the SEM image in Figure 3.2(a). However, the entire batch of
nanowires is randomly oriented, rather than parallel to the carrying gas, as evident in
Figure 3.2(b).

Figure 3.1. Schematic representation of the setup used for the synthesis of indium oxide nanowire. The
gray box and the red dots represent the furnace and the heating coils, respectively. A piece of InAs is
atomized by laser ablation. Indium atoms are carried over the catalyst by the carrying gas, where the
VLS growth of indium oxide nanowires takes place.
95

Typically, “as grown” nanowires exhibit dimensions of 10-20 nm for the diameter
and 3-8 m for the length. TEM analysis of these nanowires demonstrated that they grow
as single crystals.
1
This analysis was carried out years ago and it is not repeated every
time a new batch of nanowires is grown. It is known that there is a significant variation in
the quality of these nanowires, from batch to batch. However, for many years we have
assumed that these nanowires grow as single crystals, hoping that the best results would
be reproducible, but many times the electronic properties of certain batches of nanowires
were below expectations.

Indium oxide nanowires are semiconducting materials and thus can be used to
fabricate FETs. Our In
2
O
3
nanowires display n-type semiconductor behavior. Oxygen
vacancies are believed to provide the necessary level of doping to exhibit the n-type
characteristics.
2-4
Although devices were always fabricated by my engineering
colleagues, Chao Li and Fumiaki Ishikawa, I have been involved in testing devices,
(a)
(b)

Figure 3.2. Typical indium oxide nanowires grown in Professor Zhou’s laboratory. (a) An individual
In
2
O
3
nanowire. (b) A network of “as grown” In
2
O
3
nanowires.
96

analyzing certain electrical characteristics, and selecting the most suitable devices from a
large array to be used as nanobiosensors. Device characteristic measurements and
analysis for biosensing applications are discussed in Chapters Five.

3.2 Surface Modification
The surface of In
2
O
3
nanowires—the In
2
O
3
/air interface-- is rich in –OH terminal
groups, as illustrated in Figure 3.3(a). Also, the In
2
O
3
surface is very similar to the
surface of indium-tin-oxide (ITO) coated glass. Therefore, we hypothesized that chemical
procedures used to modify the surface of ITO would also apply to our indium oxide
nanowires. Several functional groups, such as amines, carbonates, and siloxanes have
been used as anchoring points to self assemble monolayers of desired coating molecules.
However, the strongest binders to indium oxide surfaces appeared to be phosphonate-
based molecules (Figure 3.3(b)). Hydroxyl functional groups on the surface of In
2
O
3

nanowires hydrolyze the terminal phosphonate group, resulting in the covalent
immobilization of the phosphonate-based molecules to the nanowire surface (Figure
3.3(c)). Usually, a freshly cleaned nanowire/ITO surface is submerged overnight in a 0.1

Figure 3.3. Phosphonate-based bifunctional linker molecules as modifiers for the surface of indium
oxide nanowires. Freshly cleaned In
2
O
3
nanowires possess numerous –OH groups on their surface (a),
which can hydrolyze phosphonate-based molecules (b) resulting in the immobilization of the latter
molecules on the nanowire surface (c).
97

mM aqueous solution of the phosphonate-based molecule, resulting in the self assembly
of the coating layer. A baking step under rigorously controlled conditions (inert
atmosphere, 120
o
C, and 12 hours) has recently been added to the procedure to ensure the
strong immobilization of the molecule to the surface.
Such phosphonate monolayers are commonly characterized by XPS or surface
FTIR. However, both the XPS and the FTIR at USC were either out of service or not
sufficiently sensitive for the characterization of such monolayers. Therefore, the
phosphonate monolayer on ITO/nanowires had to be characterized using different
methods:
3.2.a. Contact angle measurement of hydrophilic/hydrophobic modified ITOs
3.2.b. Observation of fluorescence signal from immobilized dye
3.2.c. Observation of the redox waves from immobilized ferrocene derivatives
3.2.d. SEM imaging of bound streptavidin labeled with Au nanoparticles

3.2.a. Contact angle measurement of hydrophilic/hydrophobic modified ITOs
A simple method to determine whether a given molecules can self-assemble on a
surface entails contact angle measurements. Of course, this method applies well only if
there is a significant difference between the modified and unmodified surface. A freshly
cleaned ITO piece has contact angle ranging between 10-15 degrees. If a freshly cleaned
ITO is submerged overnight in an ethanolic solution of 1-phosphonohexanoic acid, the
resulting surface displays contact angles over 120 degrees. Clearly, the phosphonate-
based molecule was assembled as a densely packed monolayer. Other phosphonate
98

derivatives resulted in more hydrophilic coating layers, such as 3-phosphonopropamine
(contact angle 80-90 degrees) or 3-phosphonopropanoic acid (contact angle 65-75
degrees). The ability to create hydrophilic and hydrophobic surfaces indicates that our
surface modification strategy is effective.
After having gained confidence in the use of different bifunctional linker
molecules, I moved forward and anchored biomolecules to the modified ITO/nanowires.
The overall goal of this portion of the project was to immobilize biological molecules to
be used as capture probes for biosensing applications according to the strategy outlined in
Figure 3.4.

Figure 3.4. Surface treatment to immobilize amine-rich biological molecules on the surface of indium
oxide nanowires. A phosphonate-based bifunctional linker molecule is used to prepare the surface for
bioconjugation. Terminal COOH groups of the linker molecules are activated with EDC/NHS in step
(a) and then used to anchor biological molecules via amine groups (step b).
99

3.2.b. Observation of fluorescence signals from immobilized dyes
This study entails using a phosphonate-based bifunctional linker molecule to
covalently bind either a dye molecule or a ligand that will eventually coordinate a
secondary, dye-labeled biomolecule. The surface chemistry used here is very similar to
the procedure used to immobilize biomolecules in Figure 3.4. This procedure was carried
out mainly on ITO coated pieces of glass, since it is easier to obtain and handle such
substrates. Three different emitting dyes were used in this study, and in each case the
sample substrate emitted much brighter light than the control pieces, as shown in Figure
3.5. For each sample/control pair, the same exposure time was used during image

Figure 3.5. Fluorescence study using ITO substrates. Sample (a) was prepared using a blue dye
(AMCA-hydrazine, Pierce); control (b) was passivated with ethanolamine prior to the reaction with the
blue dye. Sample (c) was prepared by immobilization of a single stranded DNA on the ITO and
hybriziation with the complementary single stranded DNA labeled with a green dye; control (d) was
obtained using a mismatched DNA (but still labeled with the same green dye). Sample (e) was prepared
by anchoring biotin to the ITO and exposing it to streptavidin (labeled with a red dye); control (f) was
made by ethanolamine deactivation prior to biotin binding and then exposed to the red-streptavidin.
100

capturing. In Figure 3.5, sample (a) was prepared using a blue dye (AMCA-hydrazine,
Pierce) that bound directly to the EDC activated surface. The corresponding control piece
of ITO (b) was passivated with ethanolamine prior to the reaction with the blue dye.
Sample (c) was prepared by immobilization of a single stranded DNA on the ITO. Upon
hybriziation with the complementary single stranded DNA labeled with a green dye, the
sample appeared brighter than the control (d), which was obtained using a mismatched
DNA (but labeled with the same green dye). Sample (e) was prepared by anchoring biotin
to the ITO. Upon exposure to streptavidin (labeled with a red dye), the sample appears
much brighter than the control (f). The control was made by ethanol amine deactivation
prior to biotin binding. This fluorescence study demonstrates that the overall surface
modification technique is valid.

3.2.c. Observation of the redox waves from immobilized ferrocene derivatives
Another useful method to determine the efficacy of the immobilization techniques
entails covalently binding an electrochemically active molecule and then observing the
redox signal. ITO coated glass can be used as a working electrode in an electrochemical
cell. Thus, if I immobilize a ferrocene derivative to the ITO, I should be able to observe
its redox signal. For this reason, 2-ethylaminoferrocene was synthesized by reduction of
the commercially available 1-cyanomethylene ferrocene. This amino-ferrocene derivative
was immobilized according to the strategy outlined in Figure 3.6.
101

The ferrocene modified ITO is placed in an electrochemical cell and the
cyclovoltammetry (CV) trace was recorded. The resulting CV trace is shown in Figure
3.7. The control ITO piece was not activated with EDC/NHS, but was exposed to
aminoferrocene. Therefore the aminoferrocene was not supposed to bind to the control
ITO. However, as soon as I took the CV trace, I observed the redox signal from both
pieces, which had a comparable current signal: the sample (top left graph in Figure 3.7)
and the control (top right). After being washed with a few mL of 10% acetic acid in
water, the control piece did not show the redox signal anymore (lower right graph in
Figure 3.7), while the sample still produced the signal (lower left). I believed that amino
ferrocene might have formed an ammonium salt with the COOH terminated ITO surface
and that only washing with acidic solution removed it. In any case, our strategy to
immobilize a molecule to the ITO was clearly working. The lesson learned here was to
apply activation reagents to the control piece anyway and then passivate it with either

Figure 3.6. Strategy for immobilization of an amino ferrocene to indium oxide nanowires.
102

ethanol amine or other alkyl amines so the resulting surface is not ionic anymore and
does not trap other molecules by forming ammonium salts.

Figure 3.7. Cyclic voltammetry traces of a ferrocene derivative bound to ITO. The sample (left panels)
was prepared according to reaction steps outlined in Figure 3.6. The control (right panels) however was
not activated with EDC/NHS, and retained its COOH surface. This carboxylic rich surface might have
formed an ammonium salt with the aminoferrocene. In fact, the sample and control initially displayed a
similar CV trace signal, but washing with 10% acetic acid removed the non-covalently bound ferrocene
from the control.
103

3.2.d SEM imaging of bound streptavidin labeled with Au nanoparticles
A fourth method to study surface immobilization strategies entails visualizing the
immobilized, labeled molecule by electron microscopy. This time the label was not a
fluorescence dye nor an electrochemically active molecule, but a gold nanoparticle
readily visible with an SEM microscope. An SEM image of these streptavidin-modified
gold nanoparticles is shown in Figure 3.8. The immobilization strategy is outlined in
Figure 3.9. After the usual 3-phosphonopropanoic acid coating and activation with
EDC/NHS, an amine-terminated biotin was bound to the nanowire surface from an
aqueous solution. Subsequent exposure of the nanowires to a solution of streptavidin-
modified gold nanoparticles (25 nm) promotes the biotin-streptavidn complex formation
on the nanowire surface, thus bringing the Au nanoparticles close to the nanowires.
Upon biotin-streptavidin complex formation, the nanowires appear to be decorated with
nanoparticles, as show in the SEM images in Figure 3.10. Even though there are many
nanoparticles on the substrate, there is a high concentration of them on the nanowires.
Clearly, the functionalization strategy is working.

Figure 3.8. The gold nanoparticles used in this study are 25 nm in diameter, and have streptavidin
molecules bound to their surface.
104

The control experiments supported the functionalization method. As a control
experiment, I deactivated the NHS-coating layer with ethanolamine, so that biotin would
not bind to the nanowire surface, and consequently no streptavidin-gold would attach to

Figure 3.9. Procedure to bind gold nanoparticle-labeled streptavidin to ITOs. (a) EDC/NHS activation
in MES buffer (pH ~4.5). (b) Exposure to a 0.1 mM solution of amine terminated biotin, in PBS buffer,
pH 7.4. (c) Exposure to a solution of gold nanoparticle-labeled streptavidin in PBS buffer, pH 7.4.

Figure 3.10. SEM images showing several examples of nanowires decorated with gold nanoparticle-
labeled streptavidin.
105

the nanowire. The results of this control experiment are shown in the SEM images in
Figure 3.11.
This nanowire modification strategy—based on phosphonate-containing
bifunctional linker molecules—will be used for the experiments related to biosensing
described in Chapter 5.

3.3 Selective Functionalization of In
2
O
3
Nanowire-Based FET Devices
3.3.a Overview of Selective Functionalization Techniques
For certain biosensing applications it would be valuable to have dense arrays of
electrodes/devices with different probes attached. This site-selective surface

Figure 3.11. Control samples of the experiments shown in Figure 3.10. Gold-labeled streptavidin does
not seem capable of binding to the passivated nanowires . The large particles in some images are
residual aggregates of salts from the buffer. They are too big to be the 25 nm gold nanoparticles.
106

functionalization has been investigated extensively and is based on inducing surface
reactions on demand. This technique has been shown to be useful in protein micro-
patterning
5-14
and electrically functionalization of multi-electrode devices.
15-17
.
Electrochemical activation is a particularly popular approach to selective surface
functionalization due to its ability to independently address individual electrodes.
18
To be
functionalized in a controlled manner, the surface of the electrodes needs to be activated
or deactivated on demand so that another desired molecule can be site-selectively
immobilized. The activation and deactivation processes are achieved through a redox-
active monolayer on the surface. By controlling the voltage on a designated electrode, the
monolayer can be oxidized or reduced. In general, one of the two redox states will
constitute the “OFF” state for the monolayer and this state will be chemically inert. The
other redox state will on the other hand be reactive toward a certain chemical (“ON”
state).
Electrochemically controlled selective functionalization of metal
5-14
and
semiconductor
16, 17
surfaces have been studied by several groups. However these studies
were never aiming at creating nanobiosensors for multiplex analysis. We envisioned
creating dense arrays of electrodes/devices, where each electrode/device has a different
capture probe anchored to its surface, as conceptually shown in figure 3.12(a). In this
cartoon, three different capture probes (or recognition groups) are immobilized on three
independent electrodes that might be spaced by just a few microns. While we
individuated the “right” molecule for the job, Professor James Heath’s group managed to
publish a paper using the same technology just a few months earlier than our paper.
16
The
107

“desired” molecule is a redox active molecule based on benzoquinone (ON) /
hydroquinone (OFF) (shown in Figure 3.12 (b)) which is well suited for this job, as
largely demonstrated on gold electrodes by Professor Milan Mrksich’s group.
10-14

The overall strategy for the selective functionalization of a dense array of FET
devices is outlined in Figure 3.13. The dense array of nanowire FETs is represented by
four identical, adjacent devices. Each device has been functionalized with a
hydroquinone derivative (blue dots) and the coating layer is resting in its OFF state, as
shown in cartoon (a). If the device that is second to the left needs to be functionalized
with a particular capture probe, this device will be used as working electrode in an
electrochemical cell. The hydroquinone coating layer will be oxidized to benzoquinone
(ON state, orange dots) (cartoon (b)). The electron poor C=C is a site for nucleophilic

Figure 3.12. Technological goal and key technology for the selective functionalization of dense arrays
of FET devices. (a) The technological goal is to functionalize a dense array of FET devices with
different capture probes (here represented by three different recognition groups on three different
devices). (b) The key technology for achieving the goal is to use an electrochemical active molecule,
such as the hydroquinone (Off state) / benzoquinone (On state) redox pair.
108

addition from a thiol terminated biomolecule (c), which covalently binds to the C=C
according to the Michal addition mechanism.
16
Other reactive functional groups toward
benzoquinone include azides, cyclopentadiene, and amines. Any unreacted benzoquinone
will be either switched off (reduced back to hydroquinone) or passivated with 2-
mercaptoethanol. This procedure only functionalizes the electrochemically activated
device. Repeating the same procedure will result in the selective functionalization of all
the other devices in the array, as illustrated for the third device from the left in Figure
3.13 (cartoon (c)-(e)).

Figure 3.13: Strategy for the selective functionalization of a dense array of FET devices. This dense
array is represented by four identical, parallel devices.
109

3.3.b Selective Functionalization of In
2
O
3
Nanowire Mat Devices
This work started in 2004 and was my first project in Professor Thompson’s
group. At that time, nanomaterials, such as carbon nanotubes
19-22
and silicon
nanowires,
23, 24
had been employed in the first generation of nanoscale biosensing
devices. In these devices, selectivity to a given analyte had been accomplished by
anchoring an analyte-specific recognition group to the surface of the nanotubes or
nanowires.
Prior to my doctorate studies, bare indium oxide nanowire devices had been
exclusively employed as gas/chemical sensors.
25, 26
Although these devices exhibit great
sensing properties, such as ultrahigh sensitivity down to ppb level, fast response, and fast
recovery time,
26
these devices had never been used as biosensors
25, 26
The two major
differences between gas and biological sensors is that in the latter, the In
2
O
3
nanowire
surface must be coated with a capture probe and the entire device operates while
submerged in solutions. To this end, my project focused on extending the surface
chemistry reported by Mrksich and coworkers
10, 12, 14, 27, 28
toward the selective
functionalization of arrays of In
2
O
3
nanowire FETs with capture probes. At the same
time, Professor James Heath’s group reported a related process for the selective
functionalization of silicon nanowires,
29
in which the linking molecules are anchored to
silicon nanowire surfaces using UV radical photoactivation, followed by deprotection of
the hydroquinone residue using BBr
3
. My approach achieves the In
2
O
3
nanowire surface
functionalization via a simple self-assembly process, utilizing 4-(1,4-dihydroxybenzene)-
butyl-phosphonic acid (HQ-PA), shown in Figure 3.14. This eliminates the use of UV
110

photoactivation and BBr
3
, a very corrosive Lewis acid we have found to attack many
metal oxide nanowires. Oxidized HQ-PA (Q-PA, Figure 3.14) reacts with a range of
functional groups, which can be easily incorporated into biomolecules and other
materials, such as thiols, azides, cyclopentadienes, and primary amines.
Preliminary studies were carried out on ITO coated glass, since derivatives of
phosphonic acid are well known to bind to indium tin oxide (ITO),
30
which has a surface
composition very similar to that of In
2
O
3
nanowires. After the chemical synthesis of HQ-
PA, a self-assembled monolayer (SAM) of HQ-PA was generated on freshly cleaned ITO
surfaces by submerging the substrate into a 0.1 mM aqueous solution of HQ-PA for 16
hr. HQ-PA is an electrochemically active molecule containing a hydroquinone group that
undergoes reversible oxidation/reduction at low potentials,
27
as shown in Figure 3.14.

Figure 3.14. Electrochemistry of an HQ-PA-coated ITO sheet. The chemical structure of HQ-PA and
Q-PA bound to ITO are shown on the left. The graph on the right shows 15 consecutive CV traces
demonstrating the stability and reversibility of oxidation/ reduction of a SAM of HQ-PA on an ITO
glass sheet. The oxidation wave is centered at +330 mV; the reduction wave at -200 mV.
111

The surface coverage and stability were characterized by electrochemical
analysis. A cyclic voltammetry trace (CV) of a derivatized film is shown in Figure 3.14.
The film was scanned multiple times (shown for 15 consecutive scans) in the range from
-450mV to +600mV. The oxidation wave is centered at +330mV and the reduction wave
is centered at -200mV; these results corroborate previously published data.
10, 12, 14, 27-29
Also, chronoamperometric measurements on a predefined area of the ITO sheet (0.57
cm
2
) reveal that a charge of 57 μC is consumed for the complete oxidation of the HQ-PA
SAM, at +400 mV (Figure 3.15). This gives a surface coverage of 4.9 10
-10
mol/cm
2
or
33.7 Å
2
/molecule, indicating that HQ-PA forms a densely-packed monolayer.
30, 31

Figure 3.15. Chronoamperometry traces. This electrochemical technique was used to determine the
amount of charge necessary to oxidize a predefined area of HQ-PA: an average of 56.8 μC. When a
thiol terminated DNA is bond to Q-PA, according to the reaction scheme shown on the top right, the
charge necessary to oxidize all the unreacted HQ-PA in the same area decreased to 50.95 μC.
Interpretation of these results is that 10% of the Q-PA have actually been reacted with the DNA.
112

The potential of the cell was then brought to +450mV and held for 5 seconds to
create a monolayer of Q-PA (active state). A separate ITO sheet was held at -350mV to
ensure the complete reduction of the monolayer of HQ-PA (inactive state). Both ITO
sheets were then submerged in a solution of a 10 M thiol-terminated DNA
32
for 2 hours
(solvent = PBS buffer containing a catalytic amount of triethanolamine, pH 7.40), Figure
3.16, step ii. The ITO sheets were next incubated for 30 minutes in an aqueous solution
containing the complementary DNA
32
tagged with a red fluorescence dye (Figure 3.16,
step iii). The ITO surface was rinsed with a PBS buffer (pH 7.00) containing 1M NaCl to
wash away any excess of complementary DNA. Fluorescence microscopy images of the
activated (Figure 3.16, bright) and inactivated (Figure 3.16, dark) ITO sheets demonstrate

Figure 3.16. (b) A fluorescence image of ITO surface that was oxidized to Q-PA, reacted with SH-
DNA, and the DNA paired to its complementary DNA strand labeled with a fluorescence dye. (c) A
fluorescence image of ITO with the HQ-PA monolayer went through the same DNA attachment
procedure as the ITO sheet in (b), showing little or no DNA binding.
113

the success of this selective functionalization strategy. This demonstrates that DNA binds
only to the activated ITO sheet in a uniform manner. In control experiments using a
mismatched DNA-Dye,
32
the ITO sheet appears dark indicating no pairing (not shown).
Chronoamperometry 10% of the ITO surface was covered with DNA as demonstrated by
in Figure 3.15.
Having established the successful functionalization strategy on the ITO surfaces,
the same reaction sequence was applied to In
2
O
3
nanowires. A mat sample of In
2
O
3
nanowires (average diameter of 10 nm and length of 3 m) was grown on a SiO
2
/Si wafer,
followed by photolithography and metal deposition (Ti/Au, 3nm/50nm) to pattern an
electrode array. The resulting device is shown in Figure 3.17. We chose to work with
nanowire mat devices since they have numerous advantages such as great reliability, high
sensitivity and ease of fabrication when compared to individual nanowire devices. Figure
3.17 (b) and 3.17(c) show typical SEM images of the nanowire mat sample used in this
study. Multiple nanowires were found bridging the Au electrodes. A SAM of HQ-PA was
created on a freshly cleaned In
2
O
3
nanowire mat sample using the same procedure as for
the ITO sheets. The In
2
O
3
nanowire device was placed into the electrochemical cell and
completely reduced to HQ-PA. In order to prevent the thiol terminated DNA from
attaching to the gold electrodes, the sample was treated with dodecane-1-thiol after HQ-
PA SAM formation. This resulted in the formation of a SAM of a C
12
alkyl chain on the
Au electrodes surface (Figure 3.17(a)). The device was then placed into an
electrochemical cell with both electrodes submerged in the electrolyte but with potential
applied across only select Au electrodes (upper electrodes in Figure 3.17). The potential
114

of the cell was held at +450mV for 5 seconds. In this way, the monolayer of HQ-PA
coating on specific nanowires was oxidized to Q-PA. Then, the entire device was
submerged in the SH-DNA solution for 2 hours, resulting in the selective, covalent
linkage of DNA to the nanowires with Q-PA.

Fluorescence studies were used to confirm the selective functionalization of the
In
2
O
3
nanowires array. Typical fluorescence images from similar nanowires devices with

Figure 3.17. (a) A photograph of a nanowire mat sample contacted by two groups of electrodes. Only
the HQ-PA attached to the nanowires between the upper electrodes were converted to Q-PA. (b) A
SEM image of the In
2
O
3
nanowires before functionalization. The brighter stripes are gold electrodes
covering the nanowire mat. (c) The same sample imaged at higher magnification, where the nanowire
mat is clearly visible. (d) A fluorescence image of the nanowires with Q-PA taken after DNA
attachment. The gold electrodes, passivated with an alkyl thiol, appear dark under the fluorescence
microscope. (e) A fluorescence image of the nanowires with HQ-PA after DNA incubation. The
nanowires appear dark, indicating no DNA attached to HQ-PA.
115

Q-PA and HQ-PA, which have been treated with the complementary DNA strand
containing a fluorescent dye label, are shown in Figure 3.17(d) and 3.17(e), respectively.
In Figure 3.17 (d) the gold electrodes appear as dark lines, whereas the nanowire mat
with Q-PA, derivatized with DNA, appears as bright network. In contrast, the nanowires
with HQ-PA, which underwent the same DNA treatment, do not show any fluorescence,
as seen in Figure 3.17(e). This demonstrates there is no DNA binding to the nanowires
with HQ-PA.

3.3.c. Materials
Dry Dichloromethane (DCM) was purchased from DriSolv (max water content:
50ppm); tetrahydrofuran (THF) was freshly distilled from sodium/ketone; triethyl
phosphine (Aldrich) was freshly distilled over sodium; Phosphate buffer saline (PBS)
was purchased from VWR and the pH adjusted to 7.40. Bromotrimethylsilane (SiMe
3
Br)
was purchased from Alfa Aesar. All other chemicals were purchased from Aldrich and
used without further purification. ITO coated glass was provided by Universal Display
Corporation (UDC).

3.3.d. Synthesis of HQ-PA
The electrochemically active molecule HQ-PA was not commercially available
and was synthesized according the procedure outline in Figure 3.18.
116

2-(4-Bromobuthyl)-1,4-dimethoxybenzene (1) was synthesized according to
literature procedures.
16
A solution of 1,4-dimethoxybenzene (12.061g, 87.29mmol) in
120 mL of THF was stirred for 1 minute at RT and then cooled to -78
o
C in a nitrogen
purged Schlenk flask. A solution of 1.0 M n-butyllithium in hexane (100 mmol) was
added dropwise. The mixture was stirred at -78
o
C for 15 minutes, warmed to RT and
allowed to stir for 1 hour, and then transferred to a second flask containing a solution of
1,4-dibromebutane (31.0 mL, 262mmol) in 100 mL of THF at 0
o
C. The mixture was
stirred for 4 hours at RT and the reaction was then quenched with 30 mL of saturated
NH
4
Cl. The solution was extracted with 3x40mL of CH
2
Cl
2
. The combined organic
OMe
OM e
OM e
Me O
Br
OM e
Me O
P
OE t
OE t
OH
HO
P
OE t
OE t O
O
OH
HO
P
OH
OH O
1) B u-Li
2) B r-C 4H 8-B r
P(O E t)3
BB r3
SiM e3B r
(1)
(2)
(3)
(H Q -PA )

Figure 3.18. Overall synthetic approach to make HQ-PA.
117

layers were washed with brine, dried over MgSO
4
and the solvent removed under reduced
pressure. Unreacted 1,4-dimethoxybenzene was removed by vacuum distillation. The
crude product was purified by column chromatography (hexane/DCM 90:10) yielding a
colorless oil (16.33g, 69.8%).
1
H-NMR (CDCl
3
):  1.72 (quintet, 2H), 1.87 (quintet, 2H),
2.61 (t, 2H), 3.43 (s, 3H), 3.76 (s, 3H), 3.77 (s, 3H), 6.68-6.78 (m, 3H).
2-(1,4-dimethoxybenzene)-butyl phosphonic acid diethyl ester (2). A solution
of compound (1) (6.35g, 23.26mmol) in 15mL of triethylphosphine (127.7 mmol) was
refluxed overnight under nitrogen. Excess triethylphosphine was removed in vacuo and
the crude product purified by column chromatography using hexane/DCM (50:50) as
eluent, yielding a colorless oil (7.18g, 93.5%).
1
H-NMR (CDCl
3
):  1.31 (t, 6H), 1.61-
1.83 (m, 6H), 2.59 (t, 2H), 3.76 (s, 3H), 3.77 (s, 3H), 4.03–4.16 (m, 4H), 6.68-6.78 (m,
3H).

2-(1,4-dihydroxybenzene)-butyl phosphonic acid diethyl ester (3). A solution
of (2) (3.5g, 10.6mM) in dry CH
2
Cl
2
(8mL) was cooled to -78
o
C and a 1M solution of
BBr
3
in CH
2
Cl
2
(31mL, 31mM) was added dropwise. The mixture was stirred at RT for
16 hours, quenched with 3mL of saturated NH
4
Cl, and extracted with diethylether. The
combined organic layers were washed with brine, dried over MgSO
4
, and the solvent
removed under reduced pressure. Purification by column chromatography
(hexane/acetone, 85:15) afforded a thick, yellowish oil (2.85g, 89.0%).
1
H-NMR
(CDCl
3
):  1.31 (t, 6H), 1.61-1.83 (m, 6H), 2.58 (t, 2H), 2.98 (broad s, 2H), 4.03–4.16
(m, 4H), 6.59-6.68 (m, 3H).
118

2-(1,4-dihydroxybenzene)-butyl phosphonic acid (HQ-PA). SiMe
3
Br (4.0mL,
30.32mmol) was added drop wise under nitrogen to a solution of (3) (2.85g, 9.44mmol)
in 10mL of dry CH
2
Cl
2
and stirred for 6 hours. Unreacted SiMe
3
Br and solvent were
removed under vacuum, yielding a yellowish oil. This oil was stirred for 2 hours in 10mL
of methanol after which the methanol removed under vacuum. The crude product was
dissolved in a minimum amount of methanol and recrystalized from diethylether, yielding
2.18g (93.9%).
1
H-NMR (D
2
O):  1.53-1.91 (m, 6H), 2.58 (t, 2H), 6.79-6.92 (m, 3H).

3.3.e. Device fabrication
In
2
O
3
nanowires synthesis and device fabrication. In
2
O
3
nanowires were grown
via a laser ablation method on Si/SiO
2
substrate.
30
Au clusters with a diameter of 10 nm
worked as catalyst, and the growth followed vapor-solid-liquid mechanism. Devices were
made directly on as-grown nanowires samples by photolithography, followed by Ti
(5nm)/Au (50nm) deposition. After lift-off, the device was carefully cleaned before
further modification.

3.3.f. Surface functionalization
ITO and In
2
O
3
nanowires device cleaning procedure. ITO sheets were boiled
for 5 minutes each in trichloroethylene, acetone, and finally, ethanol. The sheets were
then placed in an ozone/UV chamber for 10 minutes. The nanowires devices were
cleaned by the same procedure of the ITO but placed in the ozone/UV chamber for 3
minutes.
119

Deposition of HQ-PA and monolayer formation. Freshly cleaned ITO sheets or
nanowires were placed in an aqueous solution of HQ-PA (0.1 mM) at RT, for at least 16
hours. The samples were then washed extensively with water and dried under nitrogen.
For the In
2
O
3
nanowires a further step was taken to passivate the gold electrodes. The
sample was soaked into a 1mM solution of dodecanethiol in hexane at RT overnight.
Electrochemical Activation of the Monolayer. Oxidation and reduction of the
HQ-PA monolayer were performed in an electrochemical cell using a solution of PBS
buffer (pH 7.40) and methanol (95:5) with Pt as counter electrode and Ag/AgCl as a
reference electrode. Determination of the monolayer density was done by counting the
charge used to oxidize an area of 0.573 cm
2
and dividing by Faraday’s constant and then
by 2 (#of electrons consumed per reaction). This showed that the monolayer is closely
packed with a surface coverage of 4.9 x10
-10
mol/cm
2
or 33.7 Å
2
/molecule.
Functionalization of ITO and In
2
O
3
nanowires with DNA oligonucleotides.
DNA oligonucleotides used herein were 20 bases in length and were modified with the 5’
thiol modifier C6 (Integrated DNA Technologies, Inc). After deprotecting the thiol
modifier, the oligonucleotide was purified by chromatography and used immediately. The
probe DNA (5’–HS-GCT TTG AGG TGC GTG TTT GT, 10 M in PBS buffer pH7.40)
was applied to the sample surface and stored in a humidity chamber in the dark for
overnight. The sample was then carefully rinsed with buffer solution to remove excess
DNA. For the DNA hybridization, the target DNA (5’-Alex-ACA AAC ACG CAC CTC
AAA GC, 5 M in PBS buffer, PH 7.40, Integrated DNA Technologies, Inc) was applied
to the sample surface for 20 minutes, followed by washing with buffer solution.
120

3.3.g. Fluorescence microscopy
A Zeiss Axiovert 200M epi-fluorescence microscope (Carl Zeiss, Germany) with 63x
water-immersion objective was used for the fluorescence imaging of Alexa Fluor 546
NHS ester bound to DNA. Alex Fluor 546 from Integrated DNA Technologies had a 555
nm absorbance maximum and 571 nm emission maximum. The observation filter set
consisted of 540/25 nm excitation filter, 565 nm dichroic mirror, and 580 nm long-pass
emission filter. Images were captured and analyzed with an AxioCam MRm camera and
AxioVision3.1 software (Carl Zeiss, Germany).

3.4. Recent Achievements in the Selective Functionalization of In
2
O
3

After our preliminary investigation (Section 3.3),
32
we envisioned that our
hydroquinone/benzoquinone technology could be employed to configure multiplexed
sensor with specific capture probes. While this approach initially looked very promising,
we later (with my colleague Rui Zhang) discovered that the hydroquinone “off” state was
not a reliable “off” state. It was found that hydroquinone can also be oxidized by
dissolved oxygen in the buffer solution. While the mechanism of this undesired oxidation
is still under investigation, we can certainly state that this chemical oxidation reaction is
pH dependent (with faster kinetics in basic media) and it is catalyzed by the presence of
amines. This unreliable off state causes a significant problem for the selectivity of the
capture reagents’ binding, as the reagent can bind to devices that are not supposed to be
121

turned on. A schematic diagram of this undesired oxidation followed by the binding of a
biomolecule is shown in Figure 3.19(a).
For this demonstration we have used a planar In
2
O
3
surface functionalized with an
Hq derivative. Hp gets oxidized by dissolved oxygen in the buffer and binds to an amino
terminated biotin. Biotin then binds a dye-labeled streptavidin, resulting in a bright red
surface under a fluorescent light. We could operate under rigorous conditions with
degassed solution and low pH but this solution may be non-practical when it comes to
prepare a large array of devices. We could also apply a reducing potential to all the
devices except the one undergoing functionalization, but, because of the way we prepare

a)
b)
P
O
O
O
H
3
CO
OC H
3
P
O
O
O
H
3
CO
OC H
3
P
O
O
O
O
O
P
O
O
O
HO
OH
NH
Bio tin
am in o-b iotin
Un d e sir e d
oxid ation
SA -d ye
P
O
O
O
H
3
CO
OCH
3
P
O
O
O
HO
OH
P
O
O
O
HO
OH
NH
Bio tin
NO
Un d e s ir e d
oxid ation
NO
b inding P
O
O
O
H
3
CO
OCH
3
NO
ca ptu re
Hq
DM P
Pq
PB S
bu ffe r
PBS
bu ffer
am in o -biotin
SA -d ye
Figure 3.19. (a) A planar indium oxide surface was functionalized with an Hq derivative. Dissolved
oxygen in the buffer oxidizes Hq to Pq. Hp then binds to an amino terminated biotin. Biotin acts as a
capture probe and binds a dye-labeled streptavidin, resulting in a bright red surface under a fluorescent
light (box on the right). (b) The off state (DMP) is truly off. An indium oxide planar surface coated with
DMP does not get oxidized by dissolved oxygen, does not bind biotin, and does not capture a dye-
labeled streptavidin. Thus it looks dark under fluorescence light (box on the right).
122

our array of devices, not every nanowire/nanotube is actually electrically connected to the
source and drain electrodes. For these non-connected nanowires/nanotubes, we do not
have control on the oxidation state of the coating hydroquinone groups, which can
undergo chemical oxidation and bind capture molecules. This undesired configuration
can be problematic when detection of analytes present in traces (single molecule, single
virus) is desired, since the analyte may bind to an “electrically dead” nanowire/nanotube.
Thus, the hydroquinone/para-quinone system is not suitable for electrochemical
activation.
Fortunately, we have recently found a solution for this problem.
33
If 2,5-
dimethoxyphenyl group (DMP) (Figure 3.20) is used in place of hydroquinone, the off
state (DMP) does not get oxidized by dissolved oxygen and thus shows no reaction
toward thiols, amines or any other functional group found in proteins. So, the off state for
a DMP derivative is truly off. We demonstrated this inertness of DMP with an
O
O
O
O
MeO
OMe
X
~1.0V
Molecule
attachment
OFF state ON state
-X
(i) (ii)
DMP Pq

Figure 3.20. Structure of DMP and electrochemical deprotection. DMP derivatives can be converted
to Pq on applying a working voltage ~1.0 V (i) and Hp can then be used to immobilize biomolecules to
the nanowire surfaces (ii).
123

experiment carried out in parallel to the Hq experiment. An indium oxide planar surface
coated with a DMP derivative does not get oxidized by dissolved oxygen when
submerged in buffer (Figure 3.19(b)) and, thus it does not bind biotin. Consequently it
does not capture a dye-labeled streptadivin, resulting in a dark surface under fluorescence
light.
On applying a voltage ~ 1.0 V, DMP is converted to para-quinone, Figure 3.20(i),
and will react rapidly with amines and thiols of biomolecules, resulting in the covalent
immobilization of these capture molecules on the nanowire surface, as schematically
shown in Figure 3.20(ii). DMP can be coated on the surface of nanowires by coupling it
to an anchoring functional group that can bind to the surface of metal oxide nanowires
(phosphonate). We have chosen to use 4-(2,5-dimethoxyphenyl)-butyl phosphonic acid
for In
2
O
3
nanowires. This bifunctional molecule is very inert in their resting form, but is
activated toward chemical reactions upon oxidation. Thus a given nanowire can be easily
“turned-on” for binding.

3.5 Conclusions
In this chapter I have discussed the surface modification of indium oxide
nanowires. Bare, unmodified indium oxide nanowires had been shown to perform well as
ultrasensitive gas sensors. However, their surface needs to be modified with capture
probes in order to function as biological sensors. I have performed the initial
investigation for the surface modification of In
2
O
3
for biological functionalization, and
found that linker bifunctional molecules based on phosphonate derivatives strongly bind
124

to indium oxide surfaces, such as ITO coated glass or nanowires. The chemical procedure
developed here has been used since 2004 (with some later improvements) to functionalize
In
2
O
3
surfaces for all the biosensing applications described in Chapter Five.
Moreover, I explored an electrochemical method for the selective
functionalization of In
2
O
3
based FETs. This method utilizes a hydroquinone derivative
that, after electrochemical oxidation to paraquinone, binds biomolecules terminated with
thiols, amines, azides, or cyclopentadienes. This method selectively attached thiol-
terminated DNA to one set of FETs while the other set of FETs was unaffected, as
confirmed by fluorescence studies.

125

3.6 Chapter 3 References
1. Li, C.; Zhang, D. H.; Han, S.; Liu, X. L.; Tang, T.; Zhou, C. W., Diameter-
controlled growth of single-crystalline In2O3 nanowires and their electronic
properties. Advanced Materials 2003, 15, (2), 143-146.
2. Zhang, D. H.; Li, C.; Han, S.; Liu, X. L.; Tang, T.; Jin, W.; Zhou, C. W.,
Electronic transport studies of single-crystalline In2O3 nanowires. Applied
Physics Letters 2003, 82, (1), 112-114.
3. Lei, B.; Li, C.; Zhang, D.; Tang, T.; Zhou, C., Tuning electronic properties of
In2O3 nanowires by doping control. Applied Physics a-Materials Science &
Processing 2004, 79, (3), 439-442.
4. Tang, T.; Han, S.; Jin, W.; Liu, X. L.; Li, C.; Zhang, D. H.; Zhou, C. W.; Chen,
B.; Han, J.; Meyyapan, M., Synthesis and characterization of single-crystal
indium nitride nanowires. Journal of Materials Research 2004, 19, (2), 423-426.
5. Jonkheijm, P.; Weinrich, D.; Schroder, H.; Niemeyer, C. M.; Waldmann, H.,
Chemical Strategies for Generating Protein Biochips. Angewandte Chemie-
International Edition 2008, 47, (50), 9618-9647.
6. Kim, K.; Hwang, J.; Seo, I.; Youn, T. H.; Kwak, J., Protein micropatterning based
on electrochemically switched immobilization of bioligand on electropolymerized
film of a dually electroactive monomer. Chemical Communications 2006, (45),
4723-4725.
7. Kim, K.; Yang, H.; Jon, S.; Kim, E.; Kwak, J., Protein patterning based on
electrochemical activation of bioinactive surfaces with hydroquinone-caged
biotin. Journal of the American Chemical Society 2004, 126, (47), 15368-15369.
8. Kim, K.; Jang, M.; Yang, H. S.; Kim, E.; Kim, Y. T.; Kwak, J., Electrochemically
induced and controlled one-step covalent coupling reaction on self-assembled
monolayers. Langmuir 2004, 20, (10), 3821-3823.
9. Kim, K.; Yang, H.; Kim, E.; Han, Y. B.; Kim, Y. T.; Kang, S. H.; Kwak, J.,
Electrochemical deprotection for site-selective immobilization of biomolecules.
Langmuir 2002, 18, (5), 1460-1462.
10. Yeo, W. S.; Yousaf, M. N.; Mrksich, M., Dynamic interfaces between cells and
surfaces: Electroactive substrates that sequentially release and attach cells.
Journal of the American Chemical Society 2003, 125, (49), 14994-14995.
126

11. Mrksich, M., What can surface chemistry do for cell biology? Current Opinion in
Chemical Biology 2002, 6, (6), 794-797.
12. Yousaf, M. N.; Houseman, B. T.; Mrksich, M., Using electroactive substrates to
pattern the attachment of two different cell populations. Proceedings of the
National Academy of Sciences of the United States of America 2001, 98, (11),
5992-5996.
13. Yousaf, M. N.; Houseman, B. T.; Mrksich, M., Turning on cell migration with
electroactive substrates. Angewandte Chemie-International Edition 2001, 40, (6),
1093-1096.
14. Yousaf, M. N.; Mrksich, M., Diels-Alder reaction for the selective immobilization
of protein to electroactive self-assembled monolayers. Journal of the American
Chemical Society 1999, 121, (17), 4286-4287.
15. Devaraj, N. K.; Dinolfo, P. H.; Chidsey, C. E. D.; Collman, J. P., Selective
functionalization of independently addressed microelectrodes by electrochemical
activation and deactivation of a coupling catalyst. Journal of the American
Chemical Society 2006, 128, (6), 1794-1795.
16. Bunimovich, Y. L.; Ge, G.; Beverly, K. C.; Ries, R. S.; Hood, L.; Heath, J. R.,
Electrochemically Programmed, Spatially Selective Biofunctionalization of
Silicon Wires. Langmuir 2004, 20, (24), 10630-10638.
17. Rohde, R. D.; Agnew, H. D.; Yeo, W. S.; Bailey, R. C.; Heath, J. R., A non-
oxidative approach toward chemically and electrochemically functionalizing
Si(111). Journal of the American Chemical Society 2006, 128, (29), 9518-9525.
18. Jung, Y. H.; Lee, K. B.; Kim, Y. G.; Choi, I. S., Proton-fueled, reversible
assembly of gold nanoparticles by controlled triplex formation. Angewandte
Chemie-International Edition 2006, 45, (36), 5960-5963.
19. Star, A.; Han, T. R.; Gabriel, J. C. P.; Bradley, K.; Gruner, G., Interaction of
aromatic compounds with carbon nanotubes: Correlation to the Hammett
parameter of the substituent and measured carbon nanotube FET response. Nano
Letters 2003, 3, (10), 1421-1423.
20. Besteman, K.; Lee, J. O.; Wiertz, F. G. M.; Heering, H. A.; Dekker, C., Enzyme-
coated carbon nanotubes as single-molecule biosensors. Nano Letters 2003, 3, (6),
727-730.
127

21. Kong, J.; Franklin, N. R.; Zhou, C. W.; Chapline, M. G.; Peng, S.; Cho, K. J.;
Dai, H. J., Nanotube molecular wires as chemical sensors. Science 2000, 287,
(5453), 622-625.
22. Chen, R. J.; Bangsaruntip, S.; Drouvalakis, K. A.; Kam, N. W. S.; Shim, M.; Li,
Y. M.; Kim, W.; Utz, P. J.; Dai, H. J., Noncovalent functionalization of carbon
nanotubes for highly specific electronic biosensors. Proceedings of the National
Academy of Sciences of the United States of America 2003, 100, (9), 4984-4989.
23. Cui, Y.; Wei, Q. Q.; Park, H. K.; Lieber, C. M., Nanowire nanosensors for highly
sensitive and selective detection of biological and chemical species. Science 2001,
293, (5533), 1289-1292.
24. Patolsky, F.; Zheng, G. F.; Hayden, O.; Lakadamyali, M.; Zhuang, X. W.; Lieber,
C. M., Electrical detection of single viruses. Proceedings of the National
Academy of Sciences of the United States of America 2004, 101, (39), 14017-
14022.
25. Li, C.; Lei, B.; Zhang, D. H.; Liu, X. L.; Han, S.; Tang, T.; Rouhanizadeh, M.;
Hsiai, T.; Zhou, C. W., Chemical gating of In2O3 nanowires by organic and
biomolecules. Applied Physics Letters 2003, 83, (19), 4014-4016.
26. Zhang, D. H.; Liu, Z. Q.; Li, C.; Tang, T.; Liu, X. L.; Han, S.; Lei, B.; Zhou, C.
W., Detection of NO2 down to ppb levels using individual and multiple In2O3
nanowire devices. Nano Letters 2004, 4, (10), 1919-1924.
27. Chan, E. W. L.; Yousaf, M. N.; Mrksich, M., Understanding the role of
adsorption in the reaction of cyclopentadiene with an immobilized dienophile.
Journal of Physical Chemistry A 2000, 104, (41), 9315-9320.
28. Sampson, N. S.; Mrksich, M.; Bertozzi, C. R., Surface molecular recognition.
Proceedings of the National Academy of Sciences of the United States of America
2001, 98, (23), 12870-12871.
29. Bunimovich, Y. L.; Ge, G. L.; Beverly, K. C.; Ries, R. S.; Hood, L.; Heath, J. R.,
Electrochemically programmed, spatially selective biofunctionalization of silicon
wires. Langmuir 2004, 20, (24), 10630-10638.
30. Gardner, T. J.; Frisbie, C. D.; Wrighton, M. S., Systems for Orthogonal Self-
Assembly of Electroactive Monolayers on Au and Ito - an Approach to Molecular
Electronics. Journal of the American Chemical Society 1995, 117, (26), 6927-
6933.
128

31. Bravo, B. G.; Mebrahtu, T.; Soriaga, M. P.; Zapien, D. C.; Hubbard, A. T.;
Stickney, J. L., Reversible Redox of 2,5-Dihydroxythiophenol Chemisorbed on
Gold and Platinum-Electrodes - Evidence for Substrate-Mediated Adsorbate
Adsorbate Interactions. Langmuir 1987, 3, (4), 595-597.
32. Curreli, M.; Li, C.; Sun, Y. H.; Lei, B.; Gundersen, M. A.; Thompson, M. E.;
Zhou, C. W., Selective functionalization of In2O3 nanowire mat devices for
biosensing applications. Journal of the American Chemical Society 2005, 127,
(19), 6922-6923.
33. Zhang, R.; Curreli, M.; Ishikawa, F.; Chen, P.; Zhou, C.; Thompson, M. E.,
Selective functionalization of In2O3 nanowire mat device: the true "OFF" state.
Manuscript in Preparation.

129

CHAPTER 4:
FACILITATING THE REAL TIME DETECTION OF
BIOMOLECULES
Chapter Outline
4.1. Improvements in the Work Process
4.2. Conclusions
4.3. Chapter 4 References

4.1 Improvements in the Work Process
The sequence of steps from producing the nanomaterials to manufacturing a
sensor device to carry out the real time detection of analytes is shown in Figure 4.1. In
this flow chart, the blue beveled boxes represent basic steps in the process, while the
green boxes indicate improvements to those steps as a response to an unfavorable
working condition. This flow chart illustrates for indium oxide nanowires, but it would
apply to carbon nanotube devices with minimal modifications. Improving procedures,
obtaining more instruments, and implementing software are all essential components in a
research project as they promote time efficient work. A significant amount of time was
spent implementing the overall process leading to the real time detection of biological
molecules.
130

Wiring the device into the
sample holder and mounting
the mixing cell
Device
fabrication
Device testing and data
analysis
Surface modification of nanowires
(attachment of the linker molecule)
Microfluidic chip
assembly
Preparation for step 4.9:
instrument connection and
liquid gate assembly
Data
analysis
Capture probe
configuration
Real time
detection
Equipment for controlled synthesis(trace O
2

analyzer, barometer, 4‐point probe) and
growth conditions/results database
Additional probe station, Agilent B1500 parameter analyzer,
Agilent E5250 switch matrix, sensor chip holders, data analysis
software, and device characteristics database
Furnace for
annealing the
linker molecules
New chip
assembly
methods
Data analysis software,
sensing curves database
Nanowire
synthesis
Wiring the device into the
sample holder
4.a 4.b 4.c 4.d
4.e.i
4.f
4.g
4.e.ii
4.h
4.i 4.j
Agilent B1500 parameter analyzer, Agilent E5250
switch matrix, and connector panel
Custom‐designed
sensor chip holders

Figure 4.1. Flow chart for the overall process leading to the real time detection of biomolecules. The blue beveled boxes indicate the basic steps
in the process, and the green boxes indicate improvements/additions to a particular operation. This process is illustrated for indium oxide
nanowires, but it would apply to carbon nanotube-based devices with little modification.

131

Nanowire Synthesis (4.a). Indium oxide nanowires, unlike carbon nanotubes,
are not commercially available and need to be synthesized in our laboratory. The
synthesis of these nanowires is described in detail in Section 3.1. The initial setup for
nanowire synthesis is shown in Figure 4.2. This initial setup been shown to yield high
quality nanowires,
1
but results are often difficult to reproduce and also the rate of
production is slow. A major limitation of this setup is the uncontrolled O
2
concentration
in the furnace tube, which can often fluctuate, yielding low quality nanowires. The
improved setup, schematically shown in Figure 4.3, includes the addition of several
components used to maintain stable conditions during nanowire growth. Now, the source
of oxygen is a 500 ppm O
2
in Ar gas cylinder, which is blended with pure argon to vary
the O
2
concentration levels down to 1 ppm with an accuracy of 0.1 ppm. These oxygen
levels are monitored in real time by an oxygen trace analyzer. The pressure is monitored

Figure 4.2. Initial set up for the synthesis of indium oxide nanowires. The photographic image show a
side view of the furnace.
132

by a barometer. After nanowire growth, the conductivity of these nanowires is estimated
using a four-point-probe, and later will be correlated to the electronic properties of FET
devices fabricated using these nanowires. The purpose of this 4-point probe is to identify
the bad batches of nanowires before devices are actually fabricated. All the variables
relative to nanowire growth (oxygen concentration, temperature, pressure, laser intensity,
etc.), 4-point probe conductivity estimation, SEM images, and FET characteristics are
documented in a database and will always be available for consultation. Test FET devices
fabricated using nanowires grown under controlled conditions show an On/Off ratio of
10
3
-10
4
. Obtaining high quality, reproducible nanowires is crucial for advancing this
project since these nanowires constitute the sensing component of our devices.

Figure 4.3. New setup for the synthesis of nanowires. The top photographic images show the trace
oxygen analyzer used to monitor the oxygen (top) levels and the barometer (lower right corner).
133

Device Fabrication (4.b). Device fabrication includes nanowire dispersion on
the substrate, photolithography, metal deposition, silicon nitride deposition (optional),
and lifts off.

Device Testing and Data Analysis (4.c). Devices, often randomly chosen from a
sensor chip, are tested to verify the success of the fabrication. A probe station, shown in
Figure 4.4, was purchased and set up for this purpose. Probe manipulators are connected
to the Agilent B1500 semiconductor parameter analyzer via triaxial cables. The
availability of a probe station for this project can significantly speed up the testing of
devices.

For an in-depth analysis of device characteristics of the entire sensor array, we
have setup a system for automatic measurements. This system consists of an Agilent
B1500 semiconductor parameter analyzer, an Agilent E5250 switch matrix, a connector
panel, a sensor chip holder, data analysis software, and a device characteristics database.

Figure 4.4. The probe station and probe manipulators used to test FET devices.
134

These instruments/components/accessories are shown in Figures 4.5 and 4.6. A
sensor chip under analysis is placed in its designated spot on the printed circuit board
(PCB) (Figure 4.5 (a)). A pin probe connector card is assembled on top and immediately
establishes electrical connections between all the 24 devices in the sensor chip on one
hand and the printed circuit board on the other, 4.5(b)-(c). The PCB is then connected to
a connector panel via two computer data cables. This connector panel interfaces the
computer data cables and the triaxial cables from output ports of the Agilent E5250
switch matrix. The input ports of the switch matrix are connected to the four source-
measurements units (SMUs) of an Agilent B1500 semiconductor parameter analyzer. In a
typical device characteristic measurements require 3 SMUs to operate the device for the
source, drain, and gate electrodes. The switch matrix can rapidly shift the connection of
any FET device in the array with the 3 SMUs and this shifting is easily controlled by the
Desktop Easy Expert Plus software. With this software and setup we can take a desired
series of measurement in an automated fashion that does not require manual labor
reconnecting devices.
Using this setup, hundreds of curves can be quickly generated when the entire
array of devices is surveyed. Data analysis software was developed to rapidly reorganize
all the data generated in a spreadsheet and plot the curves of device characteristics. All
these curves are stored in a device characteristics database and will always be available
for consultation.
135

Surface Modification (4.d). The surface of the nanowires is then modified with
the linker bifunctional molecule that self assemble on the nanowire surface from aqueous
solutions.
2-4
Details of such surface modification procedures can be found in Chapter
Three and Five. After binding the linker molecule, the device used to be baked on a hot
plate for 20-30 minutes. This baking procedure was inefficient at ensuring robust binding

Figure 4.6. Instruments and essential components for the automated survey of devices.

Figure 4.5. Sample holder.

136

of the linkers. My colleague Rui Zhang found that a 12-hour baking step in nitrogen
atmosphere at 120
o
C ensures robust attachment of the linker molecules to the nanowires
surface. For this purpose, a furnace was purchased and set up. The device is then
prepared for the delivery of reagents/solutions either with a microfluidic chip (Section
4.e.i-ii) or with a simple mixing cell (Section 4.f).

Microfluidic Chip Assembly (4.e.i): Integrated Microfluidic Chip. Initially,
the sample reagent delivery system was just a Teflon cone-shape mixing cell.
5
The now-
complete microfluidic-integrated nanosensor platform is shown in Figure 4.8. The sensor
array is built from 24 independent field effect transistors (FETs), fabricated on silicon, as
shown in Figure 4.8(a). This platform is coupled with a microfluidic device 4.8(b), which
allows for accurate delivery of samples to each of the devices or a set of devices, for
multiple, simultaneous measurements, with only a few µL of sample solution. The
integrated platform is shown in Figure 4.8(c).

Figure 4.7. Furnace.
137

(a) Array of 24 devices (b) Microfluidc PDMS chip
(c) Integrated Platform

Figure 4.8. (a) Nanobiosensor chip has devices in center with electrial leads running to the edges of the
substrate. (b) The microfluidic chip is constructed of PDMS elastomer, establishing a network of
channels and valves to direct fluid flow. (c) Our integrated platform combines the microfluidic chip
with the nanosensor array.

All sensors are confined into a 3 mm x 6 mm space on the substrate (Figure 4.9a),
making the inter-device spacing compatible with PDMS based microfluidics. The
microfluidic system is designed to exactly match the layout of the FET array. The
microfluidic chambers of the elastomer chip perfectly align with the nanosensor array, as
clearly visible in Figure 4.9 (d)-(f). In this figure, (a) shows a photo image of the 24-
device array of nanosensors, (b) shows a magnified photo image of two sets of three
devices, and (c) shows a further magnified photo image of one device. Upon integration
of the microfluidic chip with the nanosensor array, sensor devices are positioned exactly
at the intersection of the sample and reagent channels, as shown in photo image (d). This
perfect overlap is clearly visible through the microfluidic chip in the magnified photo
image (e) for of two sets of three devices and in (f) for a single device. Thus, this
microfluidic chip can be used to deliver buffer solutions containing reagents, capture
probes, or analytes to the surface of the nanosensors.
138

Microfluidic Chip Assembly. A significant challenge we encountered was the
attachment of the microfluidic chip to the sensor array. The microfluidic chip is made of
PDMS, and this material does not spontaneously adhere to the silicon substrate.
Microfluidic operations require operating the chip with at least 5 PSI of difference
between the control channels and the reagent channels. Common pressures are 8-10 PSI
for the control channels and 3-5 PSI for the reagents. If the PDMS chip is not properly
attached to the sensor substrate, there will be leaking and uncontrolled flooding of the
channels. Coating the silicon substrate with a reactive siloxane derivative promotes full
adhesion.
Attachment of PDMS to silicon/silica surfaces has been achieved mostly via the
generation of highly reactive radical species on the PDMS surface via oxygen plasma

Figure 4.9. Details of the integrated platform. (a) Photo image of the 24-device array of nanosensors.
(b) Magnified photo image showing two sets of three devices. (c) Further magnified photo image
showing one device. (d) Photo image of microfluidics integrated on top of the device shown in (a): inset
wholebody of the device with microfluidics. (e) Magnified photo image of the devices with
microfluidic system in place. (f) Further magnified photo image showing a device, through the
microfluidic chip.
139

activation, which can react with OH group present on the silicon/silica surface. In order
to react, the OH groups on the silicon/silica must be vertically aligned with the radicals
on the PDMS surface. The larger the number of successful reactions, the stronger the
adhesion of PDMS to silicon/silica. Thus, it is important to have a large surface density
of OH groups on Si/SiO
2
and of radicals on the PDMS. The Si/SiO
2
chip at the end of
nanowire/nanotube growth process is highly dehydrated with a significantly reduced
number of OH groups on its surface. We have found that treating the Si/SiO
2
surface with
aminopropyltrimethoxysiloxane (APTMS) is very useful to covalently bind the surface of
silicon/silicon oxide to PDMS chips, as illustrated in Figure 4.10. By using any of the
above described linker molecules, one can accomplish a very robust attachment of PDMS
chip to a silicon/silica surface.

Figure 4.10. Attachment of the PDMS chip to an APTMS modified silicon oxide surface.
140

The procedure to attach the microfluidic chip (PDMS) to the sensor array
(silicon/silica) is outlined below and illustrated in Figure 4.11 for carbon nanotube
devices. The procedure will be similar for nanowire devices. As a first step, APTMS is
freshly distilled prior to its use, and a 5% solution in dry methanol is prepared. Then, the
silicon/silica surface is cleaned using oxygen plasma (60 Torr O
2
, 1 minute). Carbon
nanotubes or any other features present on the Si/SiO
2
surface that are sensitive to O
2

plasma can be protected with a layer of PMMA. The silicon/silica surface is submerged
in the silane-deriviative methanolic solution overnight. Unbound silane derivative is
washed away with methanol, and the protecting coating PMMA is washed away with hot
acetone or anisole. The silicon/silica chip is then baked for 1-2 hours at 120
o
C. At this
point, the Si/SiO
2
substrate is ready to adhere to the microfluidic chip. The PDMS chip is
activated with oxygen plasma (60 Torr O
2
, 10 minutes). The activated PDMS chip and
derivatized Si/SiO
2
chip are placed in contact and baked at 120
o
C for 2 hours.

Figure 4.11. Sequence of steps for the attachment of the PDMS chip to the sensor array.
141

Wiring the device (4.e.ii). Once the microfluidic chip has been assembled on the
sensor array, each FET device is connected to the PCB as described in Section 4.c.
Briefly, the microfluidic integrated sensor chip is placed on the PCB and the pin
connector card is used to connect the 24 devices with the corresponding pads on the PCB,
The pin connector card has a central aperture (window) that allows to the microfluid chip
to fit in, as shown in Figure 4.12 (a)-(b).

Wiring the device and mounting the mixing cell (4.f). An alternative approach
to microfluidics requires the use of a simple mixing Teflon cell to deliver solutions. The
same sample holder described in Section 4.3 is predisposed to incorporate this Teflon
cell, as schematically shown in Figure 4.12 (c).

Capture Probe Configuration (4.g). Once the device and delivery system are in
place and properly assembled on the PCB, the next step is to configure the device with
the appropriate capture probe, using either the microfluid or the mixing cell. The sample
holder can be easily transported into the fridge for a few days’ storage and many chips

Figure 4.12. Delivery systems—microfluidic (a) and (b) and mixing cell (c)—integrated with the
sensor arrays, mounted on the PCB, and wired using the pin connector card.
142

can be configured simultaneously since we have many holders. Details regarding the
capture probe configuration can be found in Chapter Five.

Instrument Connection and Liquid Gate Assembly (4.h). After the device has
been configured with the appropriate capture probe, the PCB is ultimately connected to
the semiconductor parameter analyzer as described in Section 4.c.
Gating the device is achieved by submerging an electrode in solution rather than
using the silicon substrate, as explained in Section 1.4.f. We initially used a simple
platinum wire submerged in (diluted) PBS buffer.
4
During real time measurements, we
noticed that after the addition of an aliquot of either pure buffer or buffered protein
solutions, the baseline signal was not stable but instead fluctuated, making measurements
very difficult and probably causing false signals. This baseline instability may have
resulted from protein binding to the surface of the Pt electrode
6
or exposing new Pt
surface upon the addition of more buffer. This can change the potential between the
source and the gate electrode, causing the baseline to shift.
6

We measured the open circuit voltage between two electrodes in solution as a
function of time and after adding buffer or protein solutions, replicating the experiment
reported by Minot et. al.
6
One of these electrodes was an encapsulated Ag/AgCl
reference electrode (ground), commonly used in electrochemistry; the other electrode was
a platinum wire, a gold wire, or another Ag/AgCl reference electrode. The experimental
setup is illustrated in Figure 4.13.
143

The open circuit voltage was monitored between the noble metal wire and the
Ag/AgCl electrode in 1 mL of 0.01x PBS. We noticed significant changes in the baseline
stability after adding to the mixing cell of aliquots of either buffer or streptavidin (SA)
solution, as shown in Figure 4.14. These additions occurred at the point in time indicated
in the figure. Notably, the baseline fluctuated by 10-15% even if no other aliquots were
added. Clearly, platinum and gold wires cannot be used to gate our sensor devices during
the real time detection of biomolecules.
On the other hand, when we repeated the same experiment using two
encapsulated Ag/AgCl electrodes, the baseline was significantly more stable, as shown in
Figure 4.15. The baseline fluctuations are short in duration--several seconds—and often
the change in potential is ~1% or less. This means that the addition of more buffer or
protein solution does not disturb the baseline, and that the encapsulated Ag/AgCl
electrode is a better candidate than noble metal wires to gate our devices for real time
detection experiments.

Mixing cell
with 0.01x
PBS buffer

Figure 4.13. Experimental setup for monitoring the potential between two electrodes immersed in
solution while more buffer or protein solutions are added to the mixing cell.
144

0 500 1000 1500 2000 2500 3000 3500 4000 4500 5000
85
90
95
100
10 uL Buffer
10 uL of 10 uM SA % Change
Time (Sec)
50 uL Buffer
0 500 1000 1500 2000 2500 3000 3500 4000 4500 5000
75
80
85
90
95
100
50 uL of
Buffer
% Change
Time (Sec)

Figure 4.14. Monitoring the potential between noble metal wires and the Ag/AgCl electrode over time
after adding either buffer or streptavidin (SA) solutions to the mixing cell. Notably, the baseline can
fluctuate—10-15%-- even if no other aliquots were added.

0 500 1000 1500 2000 2500 3000
90
92
94
96
98
100
102
104
106
108
110
Addition of
100 uL of
buffer
% Change
Time (sec)
0 500 1000 1500 2000 2500 3000 3500 4000
90
92
94
96
98
100
102
104
106
108
110
Addition of
100 uL of
150 uM BSA
Addition of
100 uL of
buffer
% Change
Time (sec)
Addition of
100 uL of
buffer

Figure 4.15. Monitoring the potential between two encapsulated Ag/AgCl electrodes over time after
adding either buffer or BSA solutions to the mixing cell. The baseline fluctuations are short in
duration--several seconds—and often the change in potential is ~1% or less.
145

Real time detection of biomolecules (4.i). The real time detection is then carried
out using the set up prepared as described in Section 4.h. Up to three devices can be
monitored simultaneously with the current set up. Our instrumentation would allow for
the addition of six more SMUs, so nine sensor devices could be monitored as long as they
all share the same gate electrode.

Data Analysis (4.j). The collected data points are sent to the data analysis
software and the graphs displayed in a database. This data will be available for posterity.

4.2 Conclusions
In this Chapter I discussed improving the overall process leading to the real time
detection of biomolecules. Initially, a significant amount of time was devoted to non-
science operations such as routine growth of nanowires (many batches were not
reproducible), waiting to use an instrument for routine device testing, plotting and
graphing data, etc. Many basic operations in the process were improved including
growing nanowires under controlled and reproducible conditions; and testing the device
in an automated fashion using a testing station consisting of our own semiconductor
parameter analyzer, switch matrix, connector panel, custom designed pin connector cards
and PCBs for fast device wiring, data analysis software, and measurements results
database for posterity. This equipment constitutes a significant improvement over our
initial working conditions and makes each step in the process much more time efficient.
146

4.3 Chapter 4 References
1. Li, C.; Zhang, D. H.; Han, S.; Liu, X. L.; Tang, T.; Zhou, C. W., Diameter-
controlled growth of single-crystalline In2O3 nanowires and their electronic
properties. Advanced Materials 2003, 15, (2), 143-146.
2. Curreli, M.; Li, C.; Sun, Y. H.; Lei, B.; Gundersen, M. A.; Thompson, M. E.;
Zhou, C. W., Selective functionalization of In2O3 nanowire mat devices for
biosensing applications. Journal of the American Chemical Society 2005, 127,
(19), 6922-6923.
3. Curreli, M.; Zhang, R.; Ishikawa, F. N.; Chang, H.-K.; Cote, R. J.; Zhou, C.;
Thompson, M. E., Real-Time, Label-Free Detection of Biological Entities Using
Nanowire-Based FETs. IEEE Transactions on Nanotechnology 2008, 7, (6), 651 -
667.
4. Ishikawa, F. N.; Chang, H.-K.; Curreli, M.; Hsiang-I Liao; Olson, C. A.; Chen, P.-
C.; Zhang, R.; Roberts, R. W.; Sun, R.; Cote, R. J.; Thompson, M. E.; Zhou, C.,
Label-Free, Electrical Detection of the SARS Virus N-Protein with Nanowire
Biosensors Utilizing Antibody Mimics as Capture Probes. ACS Nano 2009, 3, (5),
1219-1224.
5. Li, C.; Curreli, M.; Lin, H.; Lei, B.; Ishikawa, F. N.; Datar, R.; Cote, R. J.;
Thompson, M. E.; Zhou, C. W., Complementary detection of prostate-specific
antigen using ln(2)O(3) nanowires and carbon nanotubes. Journal of the
American Chemical Society 2005, 127, (36), 12484-12485.
6. Minot, E. D.; Janssens, A. M.; Heller, I.; Heering, H. A.; Dekker, C.; Lemay, S.
G., Carbon nanotube biosensors: The critical role of the reference electrode.
Applied Physics Letters 2007, 91, (9), 093507-093511.

147

CHAPTER 5:
REAL TIME DETECTION OF BIOLOGICAL MOLECULES
Chapter Outline
5.1. Introduction to Biomarkers
5.2. Detection of Prostate Specific Antigen (PSA)
5.3. Detection of the SARS’s Nucleocapsid Protein
5.4. Conclusions
5.5. Chapter 5 References

5.1 Introduction to Biomarkers
During my doctoral studies at USC, I was involved in the development of
nanobiosensors, sensor devices capable of detecting the presence of particular biological
molecules from a sample under analysis. My research team demonstrated the detection of
two distinct classes of biological molecules: cancer biomarkers
1
and infectious agent
biomarkers.
2
Although the underlying technologies are almost identical, there is a
difference that makes the detection of cancer biomarkers and infectious agents dissimilar.
Cancer biomarkers. Cancer biomarkers are unique, quantifiable, molecular
signatures (mostly proteins) found in abnormal levels in biospecimens when a particular
tumor is growing. Biospecimens are biological samples/biorepositories such as blood,
urine, saliva, sputum, stool, other fluids, or tissue specimen in human systems where
biomarkers can be found. Certain common cancer biomarkers are listed in Table 5.1,
148

along with their biorepository source. Changes in a cell’s genetic material can result in a
change of certain biological function, sometimes causing the cell to become malignant. It
might take years for the cell to reach the stage at which the damage causes tumor
development. These changes produce biomarkers, which serve as a signal for some
particular cellular state that precede cancer and later for the presence of cancer itself.
These biomarkers are predominantly proteins already present in the human’s body but the
presence of cancer alters their concentration level.
The task of nanobiosensors is to reveal the abnormal concentrations of these
biomarkers, before the tumor symptoms appear. Sensors for cancer biomarkers need to be
able to discriminate between the concentrations of a biomarker in a healthy control versus
the concentration in a tumor patient. Many times these concentrations are quite similar,
differing only by a factor of 2, while other times they are far apart, differing by several
orders of magnitude. Thus, it might be difficult to interpret analytical results in order to
successfully discriminate between healthy individuals and cancer patients. As an
Cancer Site Biomarker(s) Biorepository Source
Prostate or>ISerum Serum
Breast CA-125; CA-27-29 Serum
Colorectal CEA Serum
Pancreatic and
Biliary Tract
CA 19-9 Serum
Table 5.1: Cancer biomarkers. Cancer biomarkers are listed for some common cancers along with the
biological source. PSA stands for prostate specific antigen, CA for cancer antigen, and CEA for
carcinoembryonic antigen.
149

EOC Serum
Marker
Serum Level of Ovarian Cancer Biomarkers
(ng/mL)
Healthy control Cancer patient Trend
CA-125 <35,000 >100,000 ↑
Prolactin 5.8-6.0 68-78 ↑
OPN 1.1-1.4 9.2-13.4 ↑
IGF-II 800-900 150-190 ↓
MIF 0.07-0.08 860-1180 ↑
Leptin 13.6-15.2 6.5-9 ↓
Table 5.2. Serum levels of epithelial ovarian cancer (EOC) biomarkers determined by a multiplexed
analysis in healthy controls and EOC patients at every stage of tumor. Ranges represent one standard
deviation of the median value. This panel of EOC biomarkers includes cancer antigen 125 (CA-125),
leptin, prolactin, osteopontin (OPN), insulin-like growth factor II (IGF-II), and microphage migratory
factor (MIF).
example, Table 5.2 shows the concentration range for a panel of ovarian cancer
biomarkers.
3, 4
Although there is no overlap between control and patient levels, the
concentration of MIF in a cancer patient is about four orders of magnitude higher than in
a healthy control, whereas the leptin concentration is barely decreased by half. A
clinically useful nanobiosensor will need extensive calibration before oncologists can
reliably use nanobiosensors to accurately measure the level of cancer biomarkers and
formulate a diagnosis/prognosis based on these results. Clearly, nanobiosensors
employed in a clinical setting will need to be able to measure the biomarker concentration
in the clinically relevant range. Ultrasensitive devices capable of detecting minimal traces
(femtomolar or below) of cancer biomarkers may not be useful because the biomarker is
present in the human system anyway.

150

Infectious Agents Biomarkers. On the other hand, infectious agents, such as
viruses, are completely alien to the human body. The minutest trace of biomarkers related
to infectious agents will be a sign of a possible, in progress infection. For instance, the
biomarker HIV-1 pg 41 is a glycoprotein released in the blood stream by the HIV-1 virus
during viral activity. The smallest trace of gp 41 would disclose viral activity and thus
the presence of the HIV-1 virus in the system. Therefore, biosensors for infectious agents
need to be ultrasensitive, with very low detection limits, and do not necessarily have to
discriminate between two ranges of concentrations to determine if a candidate is ill or not
as in the case of cancer biomarkers. Medical personnel could just accept a “positive” or
“negative” as an answer for a test for infectious diseases performed with nanobiosensors.

5.2 Detection of Prostate Specific Antigen (PSA)
Background. Prostate specific antigen (PSA) is an oncological biomarker for
prostate cancer. Prostate cancer counts for about 2-3% of all male deaths in the U.S. and
is the most frequently diagnosed cancer among men in the US
5, 6
with over 230,000 new
cases each year. Risk factors leading to the development of prostate cancer include age,
family history, high serum testosterone, and a high-fat diet. Although the tumor becomes
visible only at a terminal state, its presence can be detected and monitored since prostate
cancer patients overproduce the PSA protein. They usually test positive if PSA is present
in serum over 4.0 ng/mL.
5, 6

151

Achievements of This work. At the beginning of our efforts in developing
nanobiosensors (2004-2005), the Thompson/Zhou nanosensing team fabricated FET
sensor devices sensitive to the presence of PSA.
1
Although these devices were not
calibrated to carry out quantitative analysis, our work was highly innovative at that time.
In fact, we were the first to:
1) Publish the detection of a cancer biomarker using these nanowire- / nanotube-based
FET nanosensors. Detection of any cancer biomarker using nanowire or nanotube
FETs had never been reported before. Our work demonstrated the detection of PSA in
solution to a concentration level as low as 5 ng/mL, a level useful for the clinical
diagnosis of prostate cancer.
5, 6

2) Use metal oxide nanowires in biosensing applications. Prior to our publication, the
biosensing literature reported the use of both carbon nanotubes and silicon nanowires
as materials to fabricate nanobiosensors for a number of biological analytes.
7, 8
The
application of metal oxide nanowires in this field had never been investigated.
3) Report the detection of biomolecules in a complementary fashion, using both n-type
(indium oxide nanowires) and p-type (carbon nanotubes) semiconductors to generate
complementary response signals. The combination of nanotube and nanowire devices
to detect the same analyte could reduce false detection signals and lead to the
development of novel sensing strategies.
4) Demonstrate the biofunctionalization of metal oxide nanowire-based functional
devices. While several functionalization techniques have been developed to attach
antibodies to nanotubes and silicon nanowires,
7, 8
little was reported for the biological
152

surface modification of metal oxide nanowires (e.g., In
2
O
3
and SnO
2
), which are
traditionally the key materials for sensing.
9, 10
A novel approach has been developed
to covalently attach antibodies to In
2
O
3
nanowire surfaces via the onsite surface
synthesis of a succinimidyl linking molecule.
Results and Discussion. The device structure of nanowire / nanotube sensors is
schematically shown in Figure 5.1(a), where an active channel made up of nanowires or
nanotubes bridges the source / drain electrodes, and the silicon substrate can be used as
the gate. Both individual and mat nanowires/nanotubes were used as the active channel,
as described below. The key to selective detection of PSA is to functionalize the
nanochannel surface with anti-PSA monoclonal antibodies (PSA-AB).

Si
iv
(c)
HO O
P
O
O
O
In
2
O
3
NW
i
(b)
N
OO
P
O
O
O
O O
iii ii
O
P
O
O
HN O
v
O
O
O
O
N
O
HN
Au/Ti
SiO
2

PSA
PSA
Antibody
Linker NW / SWNT
(a)

Figure 5.1. (a) Schematic diagram of the nanosensor. PSA-ABs are anchored to the nanowire /
nanotube surface and function as specific recognition groups for PSA binding. (b) Reaction sequence
for the modification of In
2
O
3
nanowire: i, deposition of 3-phosphonopropionic acid; ii, DCC and N-
hydroxysuccinimide activation; iii, PSA-AB incubation (c) Reaction sequence for the modification of
nanotube: iv, deposition of 1-pyrenebutanoic acid succinimidyl ester; v, PSA-AB incubation.
153

The functionalization strategy adopted for In
2
O
3
nanowires is schematically
shown in Figure 5.1(b), and described in depth in the Methods Section. In
2
O
3
nanowire
devices were first submerged in a solution of 3-phosphonopropionic acid, which bound
the phosphonic acid to the indium oxide surface, leaving the COOH groups available for
further reaction. The COOH groups on the nanowire surface were subsequently converted
to a carboxylate succinimidyl ester via incubation in N,N'-Dicyclohexylcarbodiimide
(DCC) and N-hydroxysuccinimide, and treated with a buffered saline solution of PSA-
AB at 50 M concentration. Antibodies were thus anchored to the nanowire surface.
Nanotube devices were fabricated by a related procedure, illustrated in Figure 5.1(c) and
described in depth in the Methods Section. The nanotube surface is first functionalized
with 1-pyrenebutanoic acid succinimidyl ester,
11
then treated with the PSA-AB solution.
Our first step after anchoring PSA-AB to the nanowire and nanotube devices was
to investigate the chemical gating effect of PSA on the devices. We incubated devices
consisting of both individual nanowires and individual semiconducting nanotubes in a
PBS buffered solution containing PSA for ~15 hours, at a concentration of 1 g/mL. The
device surface was then thoroughly rinsed with de-ionized water and dried under a stream
of nitrogen.
The electrical properties of the devices, including both current-voltage (I-V
ds
) and
current-gate voltage (I-V
g
) characteristics, were measured in air before and after the PSA
incubation. We consistently observed enhanced conductance for nanowire devices and
reduced conductance for nanotube devices after PSA incubation, as shown in Figure
5.2(a) and (c), respectively. In addition to the complementary change in conductance, the
154

gate dependence of both the nanowire and nanotube devices also changed. As shown in
Figure 5.2(b) and (d), the threshold voltage (V
T
) of the nanowire device shifted from -
14V to -8V, in contrast to a shift from -22V to -25V for the nanotube device. This
complementary response in conductance can be understood since In
2
O
3
nanowires are n-
type and nanotubes are p-type semiconductor. The change of the device characteristics
originated from the chemical gating effect of PSA, which introduced carriers into In
2
O
3

nanowires. The introduction of these carriers led to enhanced conductance, while the PSA
binding decreased the carrier concentration in nanotubes, thus reducing the conductance.
Control samples for both kinds of devices went through incubation in buffer without
PSA, and showed little change in electrical properties before and after the incubation.

V
ds
= 0.1 V
V
ds
= 0.05 V
300
200
100
0
I (nA)
1.0 0.8 0.6 0.4 0.2 0.0
V
ds
(V)
5
4
3
2
1
0
I (
1.0 0.8 0.6 0.4 0.2 0.0
V
ds
(V)
1.0
0.8
0.6
0.4
0.2
0.0
I (nA)
-30 -20 -10 0
V
g
(V)
200
150
100
50
0
I (nA)
-20 0 20
V
g
(V)
(a)
(d)
(c)
(b)

Figure 5.2. I-V and I-V
g
curves of an In
2
O
3
nanowire device (a, b) and a nanotube device (c, d) before
and after PSA incubation. Red and blue curves represent measurements performed before and after
PSA incubation, respectively.
155

We have further performed real-time PSA detection in PBS solution with both
In
2
O
3
nanowire and carbon nanotube devices. Figure 5.3 insets display the device
images. We used nanotube mat devices in order to overcome the instability found in
individual nanotube devices. The antibody-functionalized nanosensors were submerged
in PBS buffer solution. The electrical currents through the nanowire and the nanotubes
devices were monitored as several solutions were then added to the solution above the
nanosensor. First, an additional aliquot of buffer solution was added to test the
nanosensor stability. Next a solution of non-target bovine serum albumin (BSA) was
added, followed by a solution of PSA. The resulting current versus time curves are
shown in Figure 5.3, with (a) for a single In
2
O
3
nanowire device (V
ds
= 100mV) and (b)
for a nanotube mat device (V
ds
= 5mV) (time points for each addition are indicated in
Figure 5.3).
The current readings from both devices displayed little change after the addition
of the buffer solution, thus attesting to the sufficiently high stability of the devices. Upon
the addition of 100 nM BSA in PBS, the readings still did not show any appreciable
change, indicating that there was no nonspecific binding of BSA.

In sharp contrast, the
current of the nanowire device increased rapidly after being exposed to 0.14 nM (5
ng/mL) PSA, while the current of the nanotube mat device decreased relatively slowly
and stabilized at lower values upon exposure to 1.4 nM (50 ng/mL) PSA. The amplitude
of the current change was about 1.3% for the nanowire device and 2% for the nanotube
device. We note that the signal-to-noise ratio is about 20 for the nanowire device
exposed to 5 ng/mL PSA (Figure 5.3(a)), indicating that the detection limit could
156

approach 250 pg/mL. Further optimization may lead to nanosensors with sensitivities
competitive to those offered by current analytical techniques,
12
while providing
additional advantages such as reduced cost, minimal blood sample volume, direct
electrical readout, and the ability to perform multiplexed detection for many biomarkers.

Buffer
BSA
PSA
(a)
390
385
380
I (nA)
500 400 300 200 100 0
Time (s)
2 µm
2.52
2.48
2.44
2.40
I (  
3000 2000 1000 0
Time (s)
BSA
PSA
Buffer
(b)
2 µm

Figure 5.3. Current recorded over time for an individual In
2
O
3
NW device (a) and a nanotube mat
device (b) when sequentially exposed buffer, BSA and PSA. Insets: SEM images of respective
devices.
157

5.2.a. Materials and Methods
Materials. 1-pyrenebutanoic acid succinimidyl ester was purchased from
Molecular Probe. Phosphate buffer saline (PBS, pH = 7.40, 0.14M NaCl) was purchased
from Mediatech Inc. Dry dichloromethane (DCM), dry dimethylformamide (DMF), and
dry acetonitrile were purchased from DriSolv (max water content: 50ppm); N-
hydroxysuccinimide and N,N'-Dicyclohexylcarbodiimide (DCC) were purchased from
Aldrich. 3-Phosphonopropionic acid was purchased from Lancaster. Bovine serum
albumin (BSA) was purchased from Sigma. Mouse monoclonal anti-human PSA
antibody and PSA (>99% purity) were obtained from Fitzgerald Industries International
(Concord, MA). Both the antibody and antigen were subjected to D-Salt Excellulose
desalting columns (Pierce, IL) and quantified by BCA protein assay (Pierce, IL) prior to
use for experiments.
In
2
O
3
nanowires synthesis and device fabrication. In
2
O
3
nanowires were
grown via a laser ablation method on Si/SiO
2
substrate.
13, 14
Au clusters with a diameter
of 10 nm worked as a catalyst, and the growth followed a vapor-solid-liquid mechanism.
The as-grown nanowires were sonicated into the IPA solution, and then the suspension
was spun onto a Si/SiO
2
substrate. Devices were made by photolithography, followed by
Ti (5nm)/Au (50nm) deposition. After lift-off, the device was carefully cleaned before
further modification.
Carbon nanotube synthesis and device fabrication: nanotubes were grown via
a CVD method at 900
o
C on Si/SiO
2
substrate. 1000 sccm CH
4
, 20 sccm C
2
H
4
and 600
sccm H
2
were used during the synthesis. Ferritin clusters with diameters around 2-5 nm
158

worked as the catalyst. Nanotube mat devices were directly made by photolithography
from the as-grown samples with high density nanotubes. Individual carbon nanotube
devices were made by patterning catalyst islands to grow nanotubes at desired locations
and then depositing the metal contacts.
Device cleaning procedure: In
2
O
3
nanowires devices were boiled for 5 minutes
each in trichloroethylene, acetone, and ethanol. The nanowires devices were then placed
in the ozone/UV chamber for 3 minutes. Nanotube devices were boiled for 3 minutes
each in both THF and isopropanol, followed by nitrogen gas drying.
Deposition of the linking molecules: Both nanowire and nanotube devices were
functionalized with a specific bifunctional molecule. Freshly cleaned In
2
O
3
nanowires
were submerged in a 0.1 mM aqueous solution of 3-Phosphonopropionic acid for 16
hours at room temperature, resulting in the binding of the phosphonic acid residue to the
surface of the In
2
O
3
nanowires. The carboxylic acid functional group of this linking
molecule was then activated by submerging the device first in a 0.05 mM solution of
DCC in dry DCM for 30 minutes and then in a 0.05 mM solution of N-
hydroxysuccinimide in dry acetonitrile for 1 hour. This resulted in the formation of
succinimidyl ester groups on the functionalized nanowires. The succinimidyl ester group
then served as anchoring point for anti-PSA monoclonalantibody. The device was
incubated in a PBS solution of anti-PSA monoclonal antibody for 12 hours.
Solutions/samples delivery: We have used a Teflon cell to deliver solutions of
interest to the surface of the nanobiosensors. Such delivery system is shown in Figure
159

5.4. Devices placed at the bottom of the cell are in contact with the solution via a micro-
sized aperture. Four screws guarantee sealing and prevent leaking of the solutions.

Control experiment of BSA on blank nanotube samples: We performed
control experiments with a blank carbon nanotube device, which was the same procedure
used for this nanotube mat sample but without surface functionalization and antibody
attachment. Figure 5.5 shows the result of real time measurement with V
ds
= 5mV. After
the device got stabilized in 0.5 ml PBS buffer environment, we added 0.02 mL PBS
buffer, and the device didn’t show any response. Then at 430s, we added 0.02 mL 100
nM BSA in PBS buffer, and the device exhibited a significant drop in conductance. This
change came from the nonspecific binding of BSA protein onto nanotube surface.
(a)
(b)

Figure 5.4. Solutions delivery system. We have used a Teflon cell to deliver solutions of interest to
the surface of the nanobiosensors. Devices placed at the bottom of the cell are in contact with the
solution via a micro-sized aperture. Four screws guarantee sealing and prevent leaking.
160

5.3 Detection of the SARS’ Nucleocapsid Protein
Background. Several years elapsed between the work on the detection of PSA
(Section 5.2) and the report of the detection of the nucleocapsid (N) protein, a biomarker
for the highly contagious severe acute respiratory syndrome (SARS). Meanwhile, the
field of nanobiosensors caught substantial momentum. It was demonstrated that these
FET-based nanobiosensors can detect a variety of clinical relevant biomarkers and DNA
sequences
1, 7, 8, 15-18
with highly selectivity and sensitivity. Substantial effort has also
been made to investigate the physical-chemical mechanism leading to the production of a

1.44
1.42
1.40
1.38
1.36
1.34
I ( 
500 400 300 200 100 0
Time (s)
Buffer
BSA

Figure 5.5. Current recorded over time for a blank nanotube mat device when sequentially exposed to
buffer and BSA.
161

sensing signal
19
(discussed in Section 1.4). Thanks to these discoveries, it is now possible
to design nanosensors with significantly improved performances.
It is generally accepted that the nature of the probe molecule plays a crucial role
in determining the device sensitivity, especially for detection signals generated by the
electrostatic doping
19
mechanism (Section 1.4.c). As mentioned in Section 1.4.c, the
intensity of the sensing signal is strictly correlated to the distance between the captured
molecule and the nanowire. The capturing agent should be smaller in size so the target
molecule is physically closer to the nanowire surface, thus exerting a stronger gating
effect. This means that the conventional antibodies might not be the most appropriate
capture probes. While the above referenced studies—whether experimental or theoretical/
computational--focused mostly on the widely popular silicon nanowires,
7
we
independently confirm that the sensing mechanism of our indium oxide-based devices is
dominated by electrostatic doping.
20
Thus, our nanowire devices should also reap the
advantage of using tiny capture probes.
Based on these recent discoveries about nanobiosensors, we opted to employ a
tiny capture probe as a capture agent for the N protein, instead of the traditional antibody.
This work was a proof of concept to demonstrate that an AMP engineered to target the
SARS’s biomarker can function as a capture probe on our devices based on In
2
O
3

nanowires and carbon nanotubes. Once this initial milestone has been achieved, these
AMPs will allow us to build nanobiosensors for virtually any biomolecule with high
sensitivity/selectivity.

162

Achievements of This Work. Some key innovations were introduced to the field of
FET nanobiosensors with this work. These innovations include:
1) The first application of antibody mimetics proteins (AMPs) to nanowire-/nanotube-
based biosensors.
2) A design of a novel immobilization strategy to anchor a protein-based capture probe
to the nanowires so it would retain an active configuration upon surface anchoring.
3) The application of interdigitated electrodes for nanowire-based FETs. Although
interdigitated electrodes were reported for carbon nanotubes, their application to
nanowires proved to be useful.
Application of Fibronectin-Based AMPs to Nanosensors. AMPs based on
fibronectins have been introduced in Section 1.4.c. Collaborators of our research team in
Professor Roberts’ research group have extensive experience in engineering and evolving
affinity binding reagents or antibody mimic proteins. AMPs are routinely designed and
implemented in Professor Roberts’ group using mRNA display, a powerful selection
technique performed entirely in vitro.
21, 22
This combinatorial library of proteins created
using this mRNA display technology is based on scaffolded proteins, specifically on the
tenth fibronectin type III domain of human fibronectin (10FnIII). Professor Robert’s
selection cycles yielded two protein structures exhibiting binding affinity for the SARS N
protein. Further affinity maturation procedures resulted in the creation of two high
affinity binders, fibronectins N17-602 and N22-460.
23
A ribbon representation of these
two affinity binders is shown in Figure 5.6. The evolved portion of these AMPs is shown
163

in red. The affinity of N17-602 and N22-460 to N protein was 3.3nM and 0.4nM,
respectively, as determined by SPR.
23

After successful evolution to achieve high affinity, the structure of the Fn was
also engineered to meet our strategies for capture probe immobilization on the
nanowire/nanotube surface. A single cystine, placed at the C-terminus of the peptide
chain, was incorporated in the final structure of this Fn based capture probe remote from
the binding site, as shown in Figure 5.6. This unique cystine was used as anchoring site
for the covalent attachment of Fn to the nanowire/nanotube surface.
2
Using this strategy,
Fn immobilization on the nanowires occurs away from the binding region so the probe
maintains an active configuration. Our Fn can also be configured with other functional
groups useful in bioconjugation such as an azide or a cyclopentadiene (See Section 3.3).
Engineered peptide
sequence for high
affinity binding (Red)
N terminus
C terminus
Unique anchoring point to
the NW via the only thiol
functional group present
in the whole Fibronectin
peptide sequence (from a
cystine residue (C)).
Peptide sequence after the
Fibronectin C terminus.
2 nm
4.3 nm

Figure 5.6. Ribbon structure of our engineered Fibronectin (Fn). The peptide sequence after the Fn C
terminus is “spelled out” to show the position of our selective attachment point to the NW surface.
164

Recent Achievements with In
2
O
3
Nanowire-Based Sensors. For indium oxide
nanowire devices, several critical issues were identified and addressed since the PSA
paper. These improvements were mainly the result of my colleague Fumiaki Ishikawa’s
diligent work. First, we have established a procedure to fabricate reliable In
2
O
3
nanowire
biosensors, based on interdigitated source and drain electrodes, as shown in Figure 5.7
(c). This fabrication procedure
2, 20
is very simple to employ, rapid, reliable, easily
scalable to a manufacturing environment, and yield uniform devices on a on a 3” wafer
scale, as shown in Figure 5.7. Second, we determined that electrostatic doping is the
dominant sensing mechanism in our devices.
20
Lastly, an annealing step added to the
procedure for the surface functionalization ensures robust binding between the
phosphonate-based linker and the nanowire surface, yielding a reliable method to anchor
capture probes to the sensor surface.

(a)
(b)
(c)
(e)
(d)

Figure 5.7. Nanobiosensor devices based on indium oxide nanowires. (a) Nanobiosensor chips as
fabricated on a 3 inches silicon wafer. We typically fabricate 9 chips with 2 different architectures per
silicon wafer. (b) One of the nanobiosensor chips. Distinguishable features include the gold pads along
the perimeter and an array of 24 sensors clustered at the center of the chip. (c) One of the 24
nanobiosensor devices. The interdigitated source and drain electrodes are clearly visible. (d) An SEM
image of the interdigitated source and drain interface showing several indium oxide nanowires. (e) A
single nanowire, the sensing element of our sensors.
165

Results and Discussions. A schematic illustration of our fibronectin-based
capture agent anchored to an In
2
O
3
nanowire field-effect transistor is shown in Figure 5.8
(a). We have measured the device characteristics utilizing a liquid gate electrode
24
while
using unmodified In
2
O
3
nanowire FET devices. Our devices exhibit excellent transistor
behavior in 0.01x phosphate buffered saline (PBS) solution. The linear behavior of the
source/drain current versus source/drain voltage (I
ds
-V
ds
) curves at V
ds,
 0.08 V (Figure
5.8 (b)) suggests ohmic contact between the nanowires and source/drain electrodes.
Strong gate dependence was also observed for a typical device, as shown by the I
ds
versus

V
g
=0.5V
b
0.4 V
0.3V
0.2V
0.1V
0.0 V

a Engineered peptide sequence
Fibronectin
Ti /Au
In
2
O
3
nanowire
S
Linker
molecule
Si/SiO
2
S
S
c

Figure 5.8. (a) Schematic diagram showing Fn immobilized on the surface of an In
2
O
3
nanowire FET
device. The regions of Fn with the engineered peptide sequence are highlighted in red. Fn was attached
to the NWs via the sulphydryl group of a cysteine near the C-terminus, remote from the binding site. (b)
A family of I
ds
-V
ds
curves and (c) a typical I
ds
-V
g
curve (plotted both in linear (red) and logarithmic
(blue)) obtained from one of our devices operating with the liquid gate configuration.
166

liquid gate voltage (I
ds
-V
g
) curves shown in linear (red curve) and logarithmic scale
(blue) in Figure 5.8 (c). The on/off ratio and transconductance were ~4.6 × 10
3
and ~3.6
µS, respectively. These results confirm the stability of our devices under active
measurement conditions.
Further surface modification of the nanowires conferred our devices with the
desired biological recognition properties. The carboxylic acid functional groups on the
nanowire surface were activated with EDC and the activated COOHs were allowed to
react with BMPH, resulting in the formation of a nanowire surface reactive toward the
unique thiol present on the Fibronectin probe molecule. The functionalized devices were
stored submerged in 1x PBS at 4 ºC.
The normalized electrical response of an Fn-modified nanowire device is shown
in Figure 5.9 (a)-(c), where we have plotted I
ds
divided by the I
ds
at t = 0 s, referred to as
I/I
0
. The device was operated at V
ds
= -200 mV and V
g
= -100 mV. Under such
experimental conditions, a baseline signal was quickly established in pure 0.01x PBS
buffer, as indicated in Figure 5.9 (a). A shift in the baseline level is often observed when
transitioning from pure buffer to protein-rich buffer, attributed to non-specific binding
interactions of proteins with the nanowire device. These non-specific binding
phenomena, if not adequately mitigated, may lead to false positive results. A technique
useful to minimize false positives during active measurements requires passivating
regions of the device that are subject to non-specific binding with a “blocking agent”
(typically BSA or Tween 20), as traditionally used in bioanalytical assays such as
ELISA.
25
We have employed BSA as “blocking agent” for our nanowire devices.
167

Aliquots of a 10 mg/mL solution of BSA were used to increase the protein concentration
of the buffer in contact with the nanowires. After each BSA addition, the baseline re-
equilibrated to a lower value of S-D current, as shown in Figure 5.9 (a) and (b).
Saturation of non-specific binding sites was achieved at 40µM concentration of BSA, as
indicated in Figure 5.9(b). The baseline stability in a protein-rich medium was then tested
by increasing the BSA concentration by 10% (4 µM) (shown with a black arrow in Figure
5.9(c)). The device showed no significant response, which confirmed that sites for non-
specific binding were blocked. The conductance of the device rapidly decreased (4%)
upon exposure the nanowire sensor to a solution containing 0.6 nM of N protein in 44
µM BSA. We further tested the response of our devices to higher N-protein
concentrations. N-protein solutions were prepared by successive additions of small
aliquots of a 100±30 nM stock solution of N-protein in 0.01x PBS containing 44 µM
BSA. When the N protein concentration was progressively increased to 2 nM, 5 nM, and
10 nM, we observed a consistent decrease in device conductance of 12%, 22%, and 31%,
respectively, relative to the baseline. Detecting the N protein in the nanomolar range can
also be achieved using current immunological clinical tests, however, our nanowire
sensors offer additional advantages such as label free detection and a comparatively short
response time.
168

BSA
addition
BSA
addition
a
N protein concentration
0.6 nM
2 nM
5 nM
10 nM
BSA Addition
BSA
N protein
Fn
b
BSA
addition
BSA
addition
BSA
addition
BSA
addition
c
Baseline stable in
pure 0.01x PBS Baseline stable in 0.01x
PBS with 40 μM BSA

Figure 5.9. Normalized electrical output (I/Io) versus time of a single operating device. (a)-(b)
Response curves to passivation upon addition of successive aliquots of BSA. Upon increasing the
concentration of BSA (from pure 0.01x PBS), the baseline re-equilibrates at lower values of S-D
current until stability is ultimately reach at 40 µM BSA, in 0.01x PBS. (c) Response for a nanowire
device functionalized with Fn. The red arrows indicate the times when the solution was raised to a
given concentration of N protein. Inset: configuration of our device during active sensing
measurements. BSA protein was used to block sites for non-specific binding. The Fn probe molecule
was then used to specifically capture the target N protein.
169

Three devices were tested simultaneously, and all the devices showed
quantitatively similar concentration dependence for their response. Plots of sensor
response versus N-protein concentration for these three devices are shown in Figure 5.10
(dots), confirming the reproducibility of the results. These plots were fitted using a
conventional Langmuir isotherm model
26,27
(solid line) and these fitting curves were used
to estimate the binding constant of Fn to the N protein. In applying this model, we
assumed that the response of the sensor is proportional to the number of captured
molecules on the sensor surface, such that I/I
0
 Fn surface coverage. Application of this
analytical model yields a binding constant of 4.9 ± 0.4 nM, which is close to the value of
the binding constant (K
D
= 3.3 nM) obtained from measurements of surface plasmon
resonance (SPR).
23
The close match of the binding constant illustrates the validity of our
assumption and the Langmuir isotherm model. The small difference may come from the
fact that the measurements were done in buffers with different ionic strengths (0.01x PBS
for nanobiosensor and 1x PBS for SPR).

Figure 5.10. Normalized response from three devices versus concentration of N protein (dots). These
plots can be fitted using a Langmuir isotherm model (solid line).
170

We conducted further experiments to confirm the role of Fn as a selective capture
probe. A nanowire surface, previously activated for bioconjugation, was treated with 2-
mercaptoethanol, prior to Fn. The Fn capture probe is not expected to bind to the
nanowire surface coated with 2-mercaptoethanol moieties, and thus this device should not
specifically recognize the N protein. A baseline was established for this device after
saturation of any site for non-specific binding with a 40 µM solution of BSA, as with the
device with Fn (Section 5.3.e). This device was then sequentially exposed to a 2, 5, and
10 nM solution of N protein, while still in the presence of 40 µM BSA. (Figure 5.11) We
did not observe any significant responses, in sharp contrast to the response observed
when we used a device functionalized with Fn, confirming that our Fn based capture
probe can selectively capture the N protein.
BSA
BSA
N protein
N protein concentration

2 nM 5 nM 10 nM

Figure 5.11. A control device without the Fn capture probe does not respond to the presence of the
SARS N protein.
171

5.3.a. Materials and Methods
Materials. Phosphate buffer saline (PBS, pH = 7.40, 10 mM phosphates, 137 mM
NaCl, 2 mM KCl) was purchased from Mediatech Inc. Water was of high purity (HPLC
grade). [(2-N-morpholino) ethanesulfonic acid] buffer (MES buffer, 100 mM, pH =5.2)
was purchased from Fluka. (1-Ethyl-3-[ 3-Dimethylaminopropyl]carbodiimide
hydrochloride) (EDC) was purchased from Aldrich. (N-[ β-Maleimidopropionic acid]
hydrazide, trifluoroacetic acid salt) (BMPH) was purchased from Pierce. 6-
Phosphohexanoic acid was purchased from Aldrich. Bovine serum albumin (BSA) was
purchased from Sigma. Aqueous solution of 0.01% Au nanoparticles (mean diameter: 9.6
nm) were purchased from BBInternational. Indium arsenide was purchased from Alfa
Aesar. Stock solutions of Fibronectin (1 µM in 1x PBS buffer) and N protein (200 nM in
1x PBS buffer) were provided by Prof. Ren Sun’s research group. The N protein solution
was buffer exchanged to 0.01x PBS before use utilizing a NAP-5 column (Amersham
Scientific). We note that during buffer exchange the concentration of the N protein is
lowered due to about two fold dilution. We estimate the concentration of N protein in
0.01x PBS to be 100 nM with an error bar of ± 30%.
Device Fabrication. In
2
O
3
nanowires were grown via a laser ablation CVD on a
Si/SiO
2
substrate, following a well established procedure in our laboratory.
1, 28
Briefly, a
stock aqueous solution of Au nanoparticles, with a diameter of about 10 nm, was
obtained by a 1:10 dilution of the purchase solution in isopropanol. A veil of this diluted
Au nanoparticle solution was allowed to dry on a Si/SiO
2
surface (typical size: 1-2 cm
2
),
resulting in a uniform surface coating of Au nanoparticles. These nanoparticles were then
172

employed as a catalyst for the laser ablation promoted growth of indium oxide nanowires,
using InAs as a source of In. Details of the nanowire growth are described elsewhere.
13, 29,
30
The as-grown nanowires were suspended in isopropanol by sonication and this
suspension was deposited onto another Si/SiO
2
substrate (typically on a 3” Si wafer with
a surface coverage of 1 nanowire per 100 μm
2
). The position of the source and drain (S-
D) electrodes was defined by photolithography. The S-D electrodes were designed to
have an interdigitated interface at the semiconductor channel, leading to FETs with
channel length and width of 2.5 µm and 780 µm, respectively. Metal deposition (5 nm of
Ti and 50 nm of Au) on the pre-patterned surface followed by lift-off completed the
device fabrication.
Surface Functionalization. In
2
O
3
nanowire devices were submerged for 5
minutes each in boiling tetrachloroethylene, acetone, ethanol, and water. The nanowire
devices were rinsed with ethanol, dried with nitrogen, placed in a ozone/UV chamber for
8-10 minutes, and then submerged in hot water (80-90 ºC) for 5 minutes.
The attachment of the Fibronectin probe molecules is schematically illustrated in
Figure 5.12. Immediately after cleaning, the nanowire devices were submerged in a 0.1
mM aqueous solution of 6-phosphonohexanoic acid for 16 hours at room temperature,
resulting in the binding of the phosphonic acid residue to the surface of the In
2
O
3

nanowires (i). These devices were baked at 125 ºC under nitrogen for 2 hr. These devices
were then wired on a custom-made printed circuit board and a mixing cell was assembled
on the device chip (Figure 5.13). This mixing cell was used to deliver and handle all the
chemical reagents necessary for surface modification and all the buffer solutions during
173

active measurements. The carboxylic acid functional groups on the nanowire surface
were activated by placing in the mixing cell a 50 mM solution of EDC in MES buffer and
allowed the activated COOHs to react with BMPH (present in the same solution at 5 mM
concentration) for 60-90 minutes (ii). Unreacted, EDC-activated carboxyl groups were
quenched with a 50 mM solution of ethanolamine (ii). This resulted in the formation of a
nanowire surface reactive toward the unique thiol present on the Fibronectin probe
molecule. This EDC/BMPH solution was removed from the mixing cell by progressive
dilution with PBS buffer. A solution containing about 200 nM of Fn in 1x PBS (along
with 1 equivalent of TCPE) was allowed to react with the nanowire surface, overnight, at
4 ºC (iii). The unreacted maleimide groups were quenched with the addition of 10 uL of a
50 mM solution of 2-mercaptoethanol and allowed to react for 10-15 minutes (iii). This
Fn solution was replaced with 1xPBS by progressive dilutions and the functionalized
devices were stored submerged in 1x PBS at 4 ºC.
174

Deposition of 6‐
phosphonohexanoic acid
1) EDC/BMPH in MES
buffer
2) Disactivation with
ethanolamine

1) Fibronectin (1 Eq.
TCEP)
2) Disactivation with 2‐
mercaptoethanol

Freshly cleaned In
2
O
3
NW
surface
(i)
(ii)
(iii)

Figure 5.12. Surface modification of our In
2
O
3
nanowire devices resulting in the covalent
immobilization of the Fibronectin probe on the nanowires.
175

Experimental setup diagram. The setup we used is shown in Figure 5.13 and
explained in the figure legend.

Figure 5.13. Details and components are indicated in the figures. (A) Cross-section schematic diagram
of the setup used for real time N protein sensing. This diagram is not on scale; the nanosensor device is
shown enlarged for clarity with respect to the rest of the set up. (B) A side view of such setup. (C) A
top-view photograph of the setup showing the Teflon cell atop a circuit board.
176

Absolute responses for N protein sensing. Figure 5.14 shows the absolute
response of the three devices used to produce Figure 5.9. Clearly, the response shows
large device-to-device variation, before normalization by each device’s initial
conductance.

Fitting using Langmuir isotherm and estimation of binding constant. The
following equation was used to describe the concentration dependence of our sensor
response:
n
n
A I I
 

  


1
/
0

where A is a constant coefficient that converts surface coverage into electrical response,
α is a constant, and n is the concentration of N protein. The values of A and α were
determined using least-squares method to give the best fit to the experimental values. The
binding constant was estimated by calculating the solution of n in the following equation:

Figure 5.14. Absolute response of the three nanowire device used to detect the N protein. These are the
same devices shown as relative response in Figure 5.9
177

5 . 0
1
1
/
0

 

   
n
n
A
I I



where θ is the percentage coverage of the surface, since the definition of binding constant
is an analyte concentration where half of the sites are occupied.

Baseline for the control experiment to confirm the role of Fn. A device was
functionalized according to steps (i), (ii), and (iii) outlined Figure 5.15. The maleimide-
surface-rich nanowire device was then treated with 2-mercaptoethanol prior to Fn
immobilization (step (iv)). With all the binding sites blocked with 2-mercaptoethanol, the
Fn capture probe is not expected to bind to the nanowire surface, and thus this device
should not specifically recognize the N protein. A stable baseline was established for this
device after saturation of any site for non-specific binding with a 40 µM solution of BSA
(Figure 5.15).
BSA
Addition
Baseline stable in a
protein‐rich
environment (40 μM
BSA) in 0.01x PBS
Baseline stable in
0.01x PBS
BSA
Addition

Figure 5.15. Establishing baseline in a protein-rich environment for the device passivated with 2-
mercaptoethanol prior to Fn.
178

5.4 Conclusions
This chapter highlights my successful surface modification of nanomaterials to
obtain functional devices. My teammates and I demonstrate the functionality of these
devices by carrying out the real time detection of biological molecules. In particular, two
successful detection experiments proved very innovative in the field of nanobiosensors.
The experiment demonstrated the complementary detection of PSA using an
individual In
2
O
3
nanowire-based and carbon nanotube network-based devices. This
experiment yielded the first biofunctionalization of a metal oxide nanowire. We achieved
specificity via proper surface functionalization, including a novel approach developed to
covalently attach PSA antibodies to In
2
O
3
nanowire surfaces. Electronic characterization
revealed enhanced conduction for In
2
O
3
nanowire devices and suppressed conduction for
nanotube devices upon PSA exposure. The more sensitive In
2
O
3
nanowire devices
showed sensitivity down to 5 ng/mL in 1xPBS; however, these results were difficult to
reproduce due to the difficulty of fabricating individual nanowire devices.
We performed the second detection experiment a few years later using sensor
devices that were a significant improvement over those in the first detection experiment.
This time, the new design of interdigitated electrodes yielded a steady supply of uniform
and highly reproducible devices. We configured these devices with an AMP capture
probe, specifically an engineered fibronectin that can selectively recognize and bind to
the N protein, a biomarker associated with the SARS coronavirus. This was the first
application of AMPs to FET biosensors. Our platform can detect the N protein at sub-
nanomolar concentrations, with a background of 44 µM bovine serum albumin (BSA).
179

This sensitivity, while comparable to current immunological detection methods, can be
obtained in a relatively short time and without the aid of any signal amplifier, such as
fluorescence labeled reagents. Ultimately, we show that our platform can also be used to
accurately determine the binding constant of the N protein and Fn by applying a
conventional Langmuir model to the concentration-dependent sensing response.
These two detection experiments are original work and represent significant
advances in this field. Notably, we are the only research team to have carried out
biomarker detection work using two separate platforms: indium oxide nanowires and
carbon nanotubes. Moreover, we are the only research team that has ever published the
application of metal oxide nanowires to biological sensing. These achievements
constitute a strong foundation for the development of diagnostic tools which can serve as
a cost-effective, rapid, portable system with health care and biomedical applications.

180

5.5 Chapter 5 References
1. Li, C.; Curreli, M.; Lin, H.; Lei, B.; Ishikawa, F. N.; Datar, R.; Cote, R. J.;
Thompson, M. E.; Zhou, C. W., Complementary detection of prostate-specific
antigen using ln(2)O(3) nanowires and carbon nanotubes. Journal of the
American Chemical Society 2005, 127, (36), 12484-12485.
2. Ishikawa, F. N.; Chang, H.-K.; Curreli, M.; Hsiang-I Liao; Olson, C. A.; Chen, P.-
C.; Zhang, R.; Roberts, R. W.; Sun, R.; Cote, R. J.; Thompson, M. E.; Zhou, C.,
Label-Free, Electrical Detection of the SARS Virus N-Protein with Nanowire
Biosensors Utilizing Antibody Mimics as Capture Probes. ACS Nano 2009, 3, (5),
1219-1224.
3. Mor, G.; Visintin, I.; Feng, Z. D.; Longton, G.; Ward, D. C.; Alvero, A. B.; Lai,
Y. L.; Tenthorey, J.; Leiser, A.; Flores, R.; Yu, H.; Azori, M.; Rutherford, T.;
Schwartz, P. E., Diagnostic markers for early detection of ovarian cancer. Cancer
Biomarkers 2008, 4, (3), 190-191.
4. Visintin, I.; Feng, Z.; Longton, G.; Ward, D. C.; Alvero, A. B.; Lai, Y. L.;
Tenthorey, J.; Leiser, A.; Flores-Saaib, R.; Yu, H.; Azori, M.; Rutherford, T.;
Schwartz, P. E.; Mor, G., Diagnostic markers for early detection of ovarian
cancer. Clinical Cancer Research 2008, 14, (4), 1065-1072.
5. Li, X.; Zhang, Y. P.; Kim, H. S.; Bae, K. H.; Stantz, K. M.; Lee, S. J.; Jung, C.
Y.; Jimenez, J. A.; Gardner, T. A.; Jeng, M. H.; Kao, C. H., Gene therapy for
prostate cancer by controlling adenovirus E1a and E4 gene expression with PSES
enhancer. Cancer Research 2005, 65, (5), 1941-1951.
6. Kelloff, G. J.; Coffey, D. S.; Chabner, B. A.; Dicker, A. P.; Guyton, K. Z.; Nisen,
P. D.; Soule, H. R.; D'Amico, A. V., Prostate-specific antigen doubling time as a
surrogate marker for evaluation of oncologic drugs to treat prostate cancer.
Clinical Cancer Research 2004, 10, (11), 3927-3933.
7. Curreli, M.; Zhang, R.; Ishikawa, F. N.; Chang, H.-K.; Cote, R. J.; Zhou, C.;
Thompson, M. E., Real-Time, Label-Free Detection of Biological Entities Using
Nanowire-Based FETs. IEEE Transactions on Nanotechnology 2008, 7, (6), 651 -
667.
8. Allen, B. L.; Kichambare, P. D.; Star, A., Carbon nanotube field-effect-transistor-
based biosensors. Advanced Materials 2007, 19, (11), 1439-1451.
181

9. Li, C.; Zhang, D. H.; Lei, B.; Han, S.; Liu, X. L.; Zhou, C. W., Surface treatment
and doping dependence of In2O3 nanowires as ammonia sensors. Journal of
Physical Chemistry B 2003, 107, (45), 12451-12455.
10. Zhang, D. H.; Liu, Z. Q.; Li, C.; Tang, T.; Liu, X. L.; Han, S.; Lei, B.; Zhou, C.
W., Detection of NO2 down to ppb levels using individual and multiple In2O3
nanowire devices. Nano Letters 2004, 4, (10), 1919-1924.
11. Chen, R. J.; Bangsaruntip, S.; Drouvalakis, K. A.; Kam, N. W. S.; Shim, M.; Li,
Y. M.; Kim, W.; Utz, P. J.; Dai, H. J., Noncovalent functionalization of carbon
nanotubes for highly specific electronic biosensors. Proceedings of the National
Academy of Sciences of the United States of America 2003, 100, (9), 4984-4989.
12. Acevedo, B.; Perera, Y.; Ruiz, M.; Rojas, G.; Benitez, J.; Ayala, M.; Gavilondo,
J., Development and validation of a quantitative ELISA for the measurement of
PSA concentration. Clinica Chimica Acta 2002, 317, (1-2), 55-63.
13. Li, C.; Zhang, D.; Han, S.; Liu, X.; Tang, T.; Lei, B.; Liu, Z.; Zhou, C., Synthesis,
electronic properties, and applications of indium oxide nanowires. Molecular
Electronics Iii 2003, 1006, 104-121.
14. Li, C.; Zhang, D. H.; Han, S.; Liu, X. L.; Tang, T.; Zhou, C. W., Diameter-
controlled growth of single-crystalline In2O3 nanowires and their electronic
properties. Advanced Materials 2003, 15, (2), 143-146.
15. Patolsky, F.; Zheng, G.; Lieber, C. M., Nanowire sensors for medicine and the life
sciences. Nanomedicine 2006, 1, (1), 51-65.
16. Stern, E.; Klemic, J. F.; Routenberg, D. A.; Wyrembak, P. N.; Turner-Evans, D.
B.; Hamilton, A. D.; LaVan, D. A.; Fahmy, T. M.; Reed, M. A., Label-free
immunodetection with CMOS-compatible semiconducting nanowires. Nature
2007, 445, (7127), 519-522.
17. Gruner, G., Carbon nanotube transistors for biosensing applications. Analytical
and Bioanalytical Chemistry 2006, 384, (2), 322-335.
18. Bunimovich, Y. L.; Shin, Y. S.; Yeo, W. S.; Amori, M.; Kwong, G.; Heath, J. R.,
Quantitative real-time measurements of DNA hybridization with alkylated
nonoxidized silicon nanowires in electrolyte solution. Journal of the American
Chemical Society 2006, 128, (50), 16323-16331.
182

19. Zhang, G.-J.; Zhang, G.; Chua, J. H.; Chee, R.-E.; Wong, E. H.; Agarwal, A.;
Buddharaju, K. D.; Singh, N.; Gao, Z.; Balasubramanian, N., DNA Sensing by
Silicon Nanowire: Charge Layer Distance Dependence. Nano Letters 2008, 8, (4),
1066-1070.
20. Ishikawa, F. N.; Curreli, M.; Chen, P. C.; Zhang, R.; Cote, R. J.; Thompson, M.
E.; Zhou, C., A Calibration Method for Nanowire Biosensors to Suppress Device-
to-device Variation. ACS Nano 2009, 3, (12), 3969-3976.
21. Liu, R.; Barrick, J.; Szostak, J. W.; Roberts, R. W., Optimized Synthesis of RNA-
Protein Fusions for In Vitro Protein Selection. Methods in Enzymology 2000, 317,
268-293.
22. Roberts, R. W.; Szostak, J. W., RNA-peptide fusions for the in vitro selection of
peptides and proteins. Proceedings of the National Academy of Sciences of the
United States of America 1997, 94, 12297-12302.
23. Liao, H. I.; Olson, C. A.; Hwang, S. M.; Deng, H. Y.; Wong, E.; Baric, R. S.;
Roberts, R. W.; Sun, R., mRNA Display Design of Fibronectin-based Intrabodies
That Detect and Inhibit Severe Acute Respiratory Syndrome Coronavirus
Nucleocapsid Protein. Journal of Biological Chemistry 2009, 284, (26), 17512-
17520.
24. Rosenblatt, S.; Yaish, Y.; Park, J.; Gore, J.; Sazonova, V.; McEuen, P. L., High
performance electrolyte gated carbon nanotube transistors. Nano Letters 2002, 2,
(8), 869-872.
25. Stadtherr, K.; Wolf, H.; Lindner, P., An aptamer-based protein biochip. Analytical
Chemistry 2005, 77, (11), 3437-3443.
26. Langmuir, I., The constitution and fundamental properties of solids and liquids
Part I Solids. Journal of the American Chemical Society 1916, 38, 2221-2295.
27. Halperin, A.; Buhot, A.; Zhulina, E. B., On the hybridization isotherms of DNA
microarrays: the Langmuir model and its extensions. Journal of Physics-
Condensed Matter 2006, 18, (18), S463-S490.
28. Curreli, M.; Li, C.; Sun, Y. H.; Lei, B.; Gundersen, M. A.; Thompson, M. E.;
Zhou, C. W., Selective functionalization of In2O3 nanowire mat devices for
biosensing applications. Journal of the American Chemical Society 2005, 127,
(19), 6922-6923.
183

29. Liu, F.; Bao, M.; Wang, K. L.; Li, C.; Lei, B.; Zhou, C., One-dimensional
transport of In2O3 nanowires. Applied Physics Letters 2005, 86, (21), 213101-
213104.
30. Lei, B.; Li, C.; Zhang, D.; Tang, T.; Zhou, C., Tuning electronic properties of
In2O3 nanowires by doping control. Applied Physics a-Materials Science &
Processing 2004, 79, (3), 439-442.
184

CHAPTER 6:
SUMMARY AND CONCLUDING REMARKS

Chapter Outline
6.1. Summary and Concluding Remarks

6.1 Summary and Concluding Remarks
Chapter 1. In this chapter, I introduced nanobiosensors and discussed several
parameters influencing the sensing curves in real time detection experiments, such as
sensitivity, selectivity, and settling time. The sensitivity is mainly affected by nanowire
and nanotube dimensions, doping levels, device geometry, gating method (back gate or
liquid gate) ionic strength of the buffer, size of the capture probe, and applied gate
voltage affect. The selectivity of these devices is directly related from the binding
affinity of the capture probe for the analyte. The settling time, the time it takes to capture
the analyte and produce a binding signal, is mainly affected by the type of delivery
system used (microfluidic or mixing cell).
This deep understanding of nanobiosensing physics can only help the field,
resulting in the design and fabrication of highly performing devices.

185

Chapter 2. In this Chapter, I discussed the investigation of novel methods for the
surface modification of carbon nanotubes for applications in FET devices, such as
nanobiosensors. Carbon nanotubes are a very promising material for sensing applications
since every atom in the material is at the surface, but their use is limited due to drawbacks
such as the presence of metallic nanotubes and challenging surface chemistry and doping.
I proposed and explored two surface modification methods that address the above
mentioned problems.
The first method I proposed and investigated is a chemical method that results in
the coordination of Cp*-Ru to the fused, six-member carbon rings of the nanotube. This
method provides a technique to covalently modify the nanotube sidewalls without
breaking any of the nanotube C=C (thus preserving the conjugation system), and at the
same time n-dope the semiconducting nanotubes via electron donation from the
coordinated Cp*-Ru. This project was highly innovative since metal sandwich
compounds using nanotubes have never been reported before. Two synthetic routes were
investigated to make the Cp*Ru-nanotube complex. Both routes yielded a product whose
UV-Vis spectrum has a typical shapeline of Cp*Ru coordinated to fused-rings arenes. IR
spectroscopy confirmed the presence of Cp*Ru in both products. However, IR
spectroscopy is also raising questions about the product electronic structure and nature of
the counter ion to [Cp*Ru]
+
. In both IR spectra, especially for route 2, the band of
absorption for the P-F bond (from PF
6
-
) is relatively weak, indicating low abundance of
this anion in the products. This low abundance of the PF
6
might be explained by a
“negative” six-member

carbon ring

η
6
coordinated to Cp*-Ru. If this is the case, the
186

reaction product Cp*-Ru-nanotube is neutral, and no counter ions such as Cl
-
or PF
6
-

are
necessary to balance to the positive Cp*-Ru
(II)
moiety. This structure is highly probable
for route 2 since this synthetic procedure uses reduced nanotubes. However, the actual
product might also be a mixture of the two structures (positive and neutral Cp*-Ru-
nanotube), with a predominance of the second structure (neutral). The proposed product
of route 1 synthesis is [Cp*Ru-nanotube][PF
6
] with a neutral carbon nanotube, a cationic
Cp*Ru, and PF
6
as a counter ion.
The Cp*Ru coordination to nanotubes was recently resumed using nanotubes of
higher purity. Individual nanotube suspensions were achieved with the aid of an
ultrasonicator. Cp*Ru coordination was accomplished according to route 1 and the
products characterized by UV-Vis-nIR and Raman spectroscopy. Both characterization
techniques confirmed the coordination of Cp*Ru to the nanotubes.
A second method to modify the nanotube surface utilizes the electrochemical-
mediated addition of arene radicals (generated in situ from reduction of diazonium salt
precursors) to the nanotube C=C. This method was investigated for two reasons. First,
provides us to destroy metallic pathways in the nanotube network in the FET channel
after drastic derivatization using radicals generated from precursors such as para-
diazonium salt of nitrobenzene. Second, this method offers an attachment points for
bioconjugation after controlled derivatization using a precursor such as the para-
diazonium salt of benzoic acid. Nanotube-based biosensors modified as described above
have never been reported before. The addition of diazonium salts was only demonstrated
on buckypapers.
187

A related project involved investigating an electrochemical method for the
selective functionalization of dense arrays of nanotube-based FET devices. A
hydroquinone derivative of pyrene was designed and synthesized, inspired by
achievements in nanowire surface modification (see Chapter Three). The functionality of
this switchable molecule was demonstrated on buckypapers.

Chapter 3. In this Chapter I discussed the surface modification of indium oxide
nanowires. Bare, unmodified indium oxide nanowires had been shown to perform well as
ultrasensitive gas sensors. However, their surface needs to be modified with capture
probes in order to function as biological sensors. I have performed the initial
investigation for the surface modification of In
2
O
3
for biological functionalization, and
found that linker bifunctional molecules based on phosphonate derivatives strongly bind
to indium oxide surfaces, such as ITO coated glass or nanowires. The chemical procedure
developed here has been used since 2004 (with some later improvements) to functionalize
In
2
O
3
surfaces for all the biosensing applications described in Chapter Five.
Moreover, I explored an electrochemical method for the selective
functionalization of In
2
O
3
based FETs. This method utilizes a hydroquinone derivative
that, after electrochemical oxidation to paraquinone, binds biomolecules terminated with
thiols, amines, azides, or cyclopentadienes. This method selectively attached thiol-
terminated DNA to one set of FETs while the other set of FETs was unaffected, as
confirmed by fluorescence studies.

188

Chapter 4. In this Chapter I discussed improving the overall process leading to
the real time detection of biomolecules. Initially, a significant amount of time was
devoted to non-science operations such as routine growth of nanowires (many batches
were not reproducible), waiting to use an instrument for routine device testing, plotting
and graphing data, etc. Many basic operations in the process were improved including
growing nanowires under controlled and reproducible conditions; and testing the device
in an automated fashion using a testing station consisting of our own semiconductor
parameter analyzer, switch matrix, connector panel, custom designed pin connector cards
and PCBs for fast device wiring, data analysis software, and measurements results
database for posterity. This equipment constitutes a significant improvement over our
initial working conditions and makes each step in the process much more time efficient.

Chapter 5. This Chapter discussed the real time detection of biological
molecules, specifically the detection of biomarkers for two classes of malignancy: cancer
and infectious diseases, and highlights my successful surface modification of
nanomaterials to obtain functional devices. In particular, two successful detection
experiments proved very innovative in the field of nanobiosensors.
The experiment demonstrated the complementary detection of PSA using an
individual In
2
O
3
nanowire-based and carbon nanotube network-based devices. In fact,
electronic characterization revealed enhanced conduction for In
2
O
3
nanowire devices and
suppressed conduction for nanotube devices upon PSA exposure. The more sensitive
In
2
O
3
nanowire devices showed sensitivity down to 5 ng/mL in 1xPBS.
189

The following innovations resulted from the detection work on PSA:
1) Published the detection of a cancer biomarker using these nanowire- / nanotube-based
FET nanosensors. Detection of any cancer biomarker using nanowire or nanotube
FETs had never been reported before.
2) Used metal oxide nanowires in biosensing applications. Prior to our publication, the
biosensing literature reported the use of both carbon nanotubes and silicon nanowires
as materials to fabricate nanobiosensors for a number of biological analytes. The
application of metal oxide nanowires in this field had never been investigated.
3) Reported the detection of biomolecules in a complementary fashion, using both n-
type (indium oxide nanowires) and p-type (carbon nanotubes) semiconductors to
generate complementary response signals. The combination of nanotube and
nanowire devices to detect the same analyte could reduce false detection signals and
lead to the development of novel sensing strategies.
4) Demonstrated the biofunctionalization of metal oxide nanowire-based functional
devices. While several functionalization techniques have been developed to attach
antibodies to nanotubes and silicon nanowires, little was reported for the biological
surface modification of metal oxide nanowires (e.g., In
2
O
3
and SnO
2
), which are
traditionally the key materials for sensing. A novel approach has been developed to
covalently attach antibodies to In
2
O
3
nanowire surfaces via the onsite surface
synthesis of a succinimidyl linking molecule.
We performed the second detection experiment a few years later using sensor
devices that were a significant improvement over those in the first detection experiment.
190

This time, the new design of interdigitated electrodes yielded a steady supply of uniform
and highly reproducible devices. We configured these devices with an AMP capture
probe, specifically an engineered fibronectin that can selectively recognize and bind to
the N protein, a biomarker associated with the SARS coronavirus. This was the first
application of AMPs to FET biosensors. Our platform can detect the N protein at sub-
nanomolar concentrations, with a background of 44 µM bovine serum albumin (BSA).
This sensitivity, while comparable to current immunological detection methods, can be
obtained in a relatively short time and without the aid of any signal amplifier, such as
fluorescence labeled reagents. Ultimately, we show that our platform can also be used to
accurately determine the binding constant of the N protein and Fn by applying a
conventional Langmuir model to the concentration-dependent sensing response.
The following innovations resulted from the detection work on SARS’s N protein:
1) The application of antibody mimetics to nanowire/nanotube detection. These AMPs
can allow us to build nanobiosensors for virtually any biomolecule with high
sensitivity/selectivity.
2) A novel immobilization strategy to anchor a protein-based capture probe to the
nanowires so it would retain an active configuration upon surface anchoring.
3) The application of interdigitated electrodes for nanowire-based FETs. Although
interdigitated electrodes were reported for carbon nanotubes, their application to
nanowires proved to be useful.
These two detection experiments are original work and represent significant
advances in this field. Notably, we are the only research team to have carried out
191

biomarker detection work using two separate platforms: indium oxide nanowires and
carbon nanotubes. Moreover, we are the only research team that has ever published the
application of metal oxide nanowires to biological sensing. These achievements
constitute a strong foundation for the development of diagnostic tools which can serve as
a cost-effective, rapid, portable system with health care and biomedical applications.
192

BIBLIOGRAPHY
Acevedo, B.; Perera, Y.; Ruiz, M.; Rojas, G.; Benitez, J.; Ayala, M.; Gavilondo, J.,
Development and validation of a quantitative ELISA for the measurement of PSA
concentration. Clinica Chimica Acta 2002, 317, (1-2), 55-63.
Allen, B. L.; Kichambare, P. D.; Star, A., Carbon nanotube field-effect-transistor-based
biosensors. Advanced Materials 2007, 19, (11), 1439-1451.
Anantram, M. P.; Leonard, F., Physics of carbon nanotube electronic devices. Reports on
Progress in Physics 2006, 69, (3), 507-561.
Angelici, R. J.; Zhu, B.; Fedi, S.; Laschi, F.; Zanello, P., Electrochemical and EPR
studies of the corannulene ruthenium(II) sandwich complex [(eta(6)-
C6Me6)Ru(eta(6)-C20H10)](SbF6)(2). Inorganic Chemistry 2007, 46, (25),
10901-10906.
Avouris, P., Molecular electronics with carbon nanotubes. Accounts of Chemical
Research 2002, 35, (12), 1026-1034.
Avouris, P.; Chen, J., Nanotube electronics and optoelectronics. Materials Today 2006, 9,
(10), 46-54.
Balasubramanian, K.; Burghard, M., Biosensors based on carbon nanotubes. Analytical
and Bioanalytical Chemistry 2006, 385, (3), 452-468.
Balasubramanian, K.; Burghard, M.; Kern, K., Effect of the electronic structure of carbon
nanotubes on the selectivity of electrochemical functionalization. Physical
Chemistry Chemical Physics 2008, 10, (16), 2256-2262.
Banerjee, S.; Hemraj-Benny, T.; Wong, S. S., Covalent surface chemistry of single-
walled carbon nanotubes. Advanced Materials 2005, 17, (1), 17-29.
Barrick, J. E.; Takahashi, T. T.; Ren, J.; Xia, T.; Roberts, R. W., Large Libraries Reveal
Diverse Solutions to an RNA Recognition Problem. Proceedings of the National
Academy of Sciences of the United States of America 2001, 98, 12374-12378.
Besteman, K.; Lee, J. O.; Wiertz, F. G. M.; Heering, H. A.; Dekker, C., Enzyme-coated
carbon nanotubes as single-molecule biosensors. Nano Letters 2003, 3, (6), 727-
730.
193

Bogunia-Kubik, K.; Sugisaka, M., From molecular biology to nanotechnology and
nanomedicine. Biosystems 2002, 65, (2-3), 123-138.
Bravo, B. G.; Mebrahtu, T.; Soriaga, M. P.; Zapien, D. C.; Hubbard, A. T.; Stickney, J.
L., Reversible Redox of 2,5-Dihydroxythiophenol Chemisorbed on Gold and
Platinum-Electrodes - Evidence for Substrate-Mediated Adsorbate Adsorbate
Interactions. Langmuir 1987, 3, (4), 595-597.
Bunimovich, Y. L.; Ge, G.; Beverly, K. C.; Ries, R. S.; Hood, L.; Heath, J. R.,
Electrochemically Programmed, Spatially Selective Biofunctionalization of
Silicon Wires. Langmuir 2004, 20, (24), 10630-10638.
Bunimovich, Y. L.; Ge, G. L.; Beverly, K. C.; Ries, R. S.; Hood, L.; Heath, J. R.,
Electrochemically programmed, spatially selective biofunctionalization of silicon
wires. Langmuir 2004, 20, (24), 10630-10638.
Bunimovich, Y. L.; Shin, Y. S.; Yeo, W. S.; Amori, M.; Kwong, G.; Heath, J. R.,
Quantitative real-time measurements of DNA hybridization with alkylated
nonoxidized silicon nanowires in electrolyte solution. Journal of the American
Chemical Society 2006, 128, (50), 16323-16331.
Byon, H. R.; Kim, S.; Choi, H. C., Label-Free Biomolecular Detection Using Carbon
Nanotube Field Effect Transistors. Nano 2008, 3, (6), 415-431.
Chan, E. W. L.; Yousaf, M. N.; Mrksich, M., Understanding the role of adsorption in the
reaction of cyclopentadiene with an immobilized dienophile. Journal of Physical
Chemistry A 2000, 104, (41), 9315-9320.
Chaturvedi, P.; Verma, P.; Singh, A.; Chaudhary, P. K.; Harsh; Basu, P. K., Carbon
nanotube - Purification and sorting protocols. Defence Science Journal 2008, 58,
(5), 591-599.
Chen, R. J.; Bangsaruntip, S.; Drouvalakis, K. A.; Kam, N. W. S.; Shim, M.; Li, Y. M.;
Kim, W.; Utz, P. J.; Dai, H. J., Noncovalent functionalization of carbon nanotubes
for highly specific electronic biosensors. Proceedings of the National Academy of
Sciences of the United States of America 2003, 100, (9), 4984-4989.
Chen, R. J.; Choi, H. C.; Bangsaruntip, S.; Yenilmez, E.; Tang, X. W.; Wang, Q.; Chang,
Y. L.; Dai, H. J., An investigation of the mechanisms of electronic sensing of
protein adsorption on carbon nanotube devices. Journal of the American
Chemical Society 2004, 126, (5), 1563-1568.
194

Chen, Y.; Wang, X.; Hong, M. K.; Erramilli, S.; Mohanty, P.; Rosenberg, C., Nanoscale
field effect transistor for biomolecular signal amplification. Applied Physics
Letters 2007, 91, (24), 243511-243514.
Cui, Y.; Wei, Q. Q.; Park, H. K.; Lieber, C. M., Nanowire nanosensors for highly
sensitive and selective detection of biological and chemical species. Science 2001,
293, (5533), 1289-1292.
Curreli, M.; Li, C.; Sun, Y. H.; Lei, B.; Gundersen, M. A.; Thompson, M. E.; Zhou, C.
W., Selective functionalization of In2O3 nanowire mat devices for biosensing
applications. Journal of the American Chemical Society 2005, 127, (19), 6922-
6923.
Curreli, M.; Zhang, R.; Ishikawa, F. N.; Chang, H.-K.; Cote, R. J.; Zhou, C.; Thompson,
M. E., Real-Time, Label-Free Detection of Biological Entities Using Nanowire-
Based FETs. IEEE Transactions on Nanotechnology 2008, 7, (6), 651 - 667.
Devaraj, N. K.; Dinolfo, P. H.; Chidsey, C. E. D.; Collman, J. P., Selective
functionalization of independently addressed microelectrodes by electrochemical
activation and deactivation of a coupling catalyst. Journal of the American
Chemical Society 2006, 128, (6), 1794-1795.
Dubin, R. A.; Callegari, G. C.; Kohn, J.; Neimark, A. V., Carbon nanotube fibers are
compatible with mammalian cells and neurons. IEEE Transactions on
Nanobioscience 2008, 7, (1), 11-14.
Elfstrom, N.; Juhasz, R.; Sychugov, I.; Engfeldt, T.; Karlstrom, A. E.; Linnros, J.,
Surface charge sensitivity of silicon nanowires: Size dependence. Nano Letters
2007, 7, (9), 2608-2612.
Endo, M.; Hayashi, T.; Kim, Y. A.; Muramatsu, H., Development and application of
carbon nanotubes. Japanese Journal of Applied Physics Part 1-Regular Papers
Brief Communications & Review Papers 2006, 45, (6A), 4883-4892.
Englebienne, P.; Van Hoonacker, A., Bionanotechnology: The science of revealing life
with nanostructures. Current Nanoscience 2005, 1, (2), 97-106.
Ferrari, M., Cancer nanotechnology: Opportunities and challenges. Nature Reviews
Cancer 2005, 5, (3), 161-171.
Gao, J. H.; Xu, B., Applications of nanomaterials inside cells. Nano Today 2009, 4, (1),
37-51.
195

Gao, Z.; Agarwal, A.; Trigg, A. D.; Singh, N.; Fang, C.; Tung, C. H.; Fan, Y.;
Buddharaju, K. D.; Kong, J., Silicon Nanowire Arrays for Label-Free Detection
of DNA. Analitical Chemisty 2007, 79, (9), 3291-3297.
Gardner, T. J.; Frisbie, C. D.; Wrighton, M. S., Systems for Orthogonal Self-Assembly of
Electroactive Monolayers on Au and Ito - an Approach to Molecular Electronics.
Journal of the American Chemical Society 1995, 117, (26), 6927-6933.
Gomez, L. M.; Kumar, A.; Zhang, Y.; Ryu, K.; Badmaev, A.; Zhou, C. W., Scalable
Light-Induced Metal to Semiconductor Conversion of Carbon Nanotubes. Nano
Letters 2009, 9, (10), 3592-3598.
Gruner, G., Carbon nanotube transistors for biosensing applications. Analytical and
Bioanalytical Chemistry 2006, 384, (2), 322-335.
Hahm, J.; Lieber, C. M., Direct ultrasensitive electrical detection of DNA and DNA
sequence variations using nanowire nanosensors. Nano Letters 2004, 4, (1), 51-
54.
Halperin, A.; Buhot, A.; Zhulina, E. B., On the hybridization isotherms of DNA
microarrays: the Langmuir model and its extensions. Journal of Physics-
Condensed Matter 2006, 18, (18), S463-S490.
Han, Y.; Offenhausser, A.; Ingebrandt, S., Detection of DNA hybridization by a field-
effect transistor with covalently attached catcher molecules. Surface and Interface
Analysis 2006, 38, (4), 176-181.
Harutyunyan, A. R.; Chen, G. G.; Paronyan, T. M.; Pigos, E. M.; Kuznetsov, O. A.;
Hewaparakrama, K.; Kim, S. M.; Zakharov, D.; Stach, E. A.; Sumanasekera, G.
U., Preferential Growth of Single-Walled Carbon Nanotubes with Metallic
Conductivity. Science 2009, 326, (5949), 116-120.
Heath, J. R.; Davis, M. E., Nanotechnology and cancer. Annual Review of Medicine 2008,
59, 251-265.
Heitzinger, C.; Klimeck, G., Computational aspects of the three-dimensional feature-
scale simulation of silicon-nanowire field-effect sensor for DNA detaction.
Journal of Computational Electronics 2007, 6, 387-390.
Heller, I.; Janssens, A. M.; Mannik, J.; Minot, E. D.; Lemay, S. G.; Dekker, C.,
Identifying the mechanism of biosensing with carbon nanotube transistors. Nano
Letters 2008, 8, (2), 591-595.
196

Holleman, A.; Cheok, M. H.; den Boer, M. L.; Yang, W. J.; Veerman, A. J. P.; Kazemier,
K. M.; Pei, D. Q.; Cheng, C.; Pui, C. H.; Relling, M. V.; Janka-Schaub, G. E.;
Pieters, R.; Evans, W. E., Gene-expression patterns in drug-resistant acute
lymphoblastic leukemia cells and response to treatment. New England Journal of
Medicine 2004, 351, (6), 533-542.
Hu, Y. H.; Shenderova, O. A.; Hu, Z.; Padgett, C. W.; Brenner, D. W., Carbon
nanostructures for advanced composites. Reports on Progress in Physics 2006,
69, (6), 1847-1895.
Huang, X. J.; Choi, Y. K., Chemical sensors based on nanostructured materials. Sensors
and Actuators B-Chemical 2007, 122, (2), 659-671.
Hussain, M. A.; Kabir, M. A.; Sood, A. K., On the cytotoxicity of carbon nanotubes.
Current Science 2009, 96, (5), 664-673.
Iijima, S., Helical Microtubules of Graphitic Carbon. Nature 1991, 354, (6348), 56-58.
Ingebrandt, S.; Offenhausser, A., Label-free detection of DNA using field-effect
transistors. Physica Status Solidi a-Applications and Materials Science 2006, 203,
(14), 3399-3411.
Ishikawa, F. N.; Chang, H. K.; Curreli, M.; Liao, H. I.; Olson, C. A.; Chen, P. C.; Zhang,
R.; Roberts, R. W.; Sun, R.; Cote, R. J.; Thompson, M. E.; Zhou, C., Label-Free,
Electrical Detection of the SARS Virus N-Protein with Nanowire Biosensors
Utilizing Antibody Mimics as Capture Probes. ACS Nano 2009, 3, (5), 1219-
1224.
Ishikawa, F. N.; Chang, H.-K.; Curreli, M.; Chen, P.; Zhang, R.; Cote, R. J.; Thompson,
M. E.; Zhou, C., Enhanced Sensitivity Observed in Biosensors Based on Carbon
Nanotube FETs Fabricated Using a Semiconducting Network of Nanotubes.
Manuscript in Preparation 2010.
Ishikawa, F. N.; Curreli, M.; Chen, P. C.; Zhang, R.; Cote, R. J.; Thompson, M. E.; Zhou,
C., A Calibration Method for Nanowire Biosensors to Suppress Device-to-device
Variation. ACS Nano 2009, 3, (12), 3969-3976.
Jin, W. J.; Sun, X. F.; Wang, Y., Solubilization and functionalization of carbon
nanotubes. New Carbon Materials 2004, 19, (4), 312-318.
Jonkheijm, P.; Weinrich, D.; Schroder, H.; Niemeyer, C. M.; Waldmann, H., Chemical
Strategies for Generating Protein Biochips. Angewandte Chemie-International
Edition 2008, 47, (50), 9618-9647.
197

Jung, Y. H.; Lee, K. B.; Kim, Y. G.; Choi, I. S., Proton-fueled, reversible assembly of
gold nanoparticles by controlled triplex formation. Angewandte Chemie-
International Edition 2006, 45, (36), 5960-5963.
Kang, B. S.; Wang, H. T.; Lele, T. P.; Tseng, Y.; Ren, F.; Pearton, S. J.; Johnson, J. W.;
Rajagopal, P.; Roberts, J. C.; Piner, E. L.; Linthicum, K. J., Prostate specific
antigen detection using AlGaN/GaN high electron mobility transistors. Applied
Physics Letters 2007, 91, (11), 112106-112109.
Kelloff, G. J.; Coffey, D. S.; Chabner, B. A.; Dicker, A. P.; Guyton, K. Z.; Nisen, P. D.;
Soule, H. R.; D'Amico, A. V., Prostate-specific antigen doubling time as a
surrogate marker for evaluation of oncologic drugs to treat prostate cancer.
Clinical Cancer Research 2004, 10, (11), 3927-3933.
Kim, A.; Ah, C. S.; Yu, H. Y.; Yang, J. H.; Baek, I. B.; Ahn, C. G.; Park, C. W.; Jun, M.
S.; Lee, S., Ultrasensitive, label-free, and real-time immunodetection using silicon
field-effect transistors. Applied Physics Letters 2007, 91, (10), 103901-103904.
Kim, K.; Hwang, J.; Seo, I.; Youn, T. H.; Kwak, J., Protein micropatterning based on
electrochemically switched immobilization of bioligand on electropolymerized
film of a dually electroactive monomer. Chemical Communications 2006, (45),
4723-4725.
Kim, K.; Jang, M.; Yang, H. S.; Kim, E.; Kim, Y. T.; Kwak, J., Electrochemically
induced and controlled one-step covalent coupling reaction on self-assembled
monolayers. Langmuir 2004, 20, (10), 3821-3823.
Kim, K.; Yang, H.; Jon, S.; Kim, E.; Kwak, J., Protein patterning based on
electrochemical activation of bioinactive surfaces with hydroquinone-caged
biotin. Journal of the American Chemical Society 2004, 126, (47), 15368-15369.
Kim, K.; Yang, H.; Kim, E.; Han, Y. B.; Kim, Y. T.; Kang, S. H.; Kwak, J.,
Electrochemical deprotection for site-selective immobilization of biomolecules.
Langmuir 2002, 18, (5), 1460-1462.
Kim, S. N.; Rusling, J. F.; Papadimitrakopoulos, F., Carbon nanotubes for electronic and
electrochemical detection of biomolecules. Advanced Materials 2007, 19, (20),
3214-3228.
Kong, J.; Franklin, N. R.; Zhou, C. W.; Chapline, M. G.; Peng, S.; Cho, K. J.; Dai, H. J.,
Nanotube molecular wires as chemical sensors. Science 2000, 287, (5453), 622-
625.
198

Kooi, S. E.; Schlecht, U.; Burghard, M.; Kern, K., Electrochemical modification of single
carbon nanotubes. Angewandte Chemie-International Edition 2002, 41, (8), 1353-
1355.
Kumar, M. J.; Reed, M. A.; Amaratunga, G. A. J.; Cohen, G. M.; Janes, D. B.; Lieber, C.
M.; Meyyappan, M.; Wernersson, L. E.; Wang, K. L.; Chau, R. S.; Kamins, T. I.;
Lundstrom, M.; Yu, B.; Zhou, C. W., Special Issue on Nanowire Transistors:
Modeling, Device Design, and Technology. IEEE Transactions on Electron
Devices 2008, 55, (11), 2813-2819.
Lacerda, L.; Bianco, A.; Prato, M.; Kostarelos, K., Carbon nanotubes as nanomedicines:
From toxicology to pharmacology. Advanced Drug Delivery Reviews 2006, 58,
(14), 1460-1470.
Lacerda, L.; Raffa, S.; Prato, M.; Bianco, A.; Kostarelos, K., Cell-penetrating CNTs for
delivery of therapeutics. Nano Today 2007, 2, (6), 38-43.
Lam, C. W.; James, J. T.; McCluskey, R.; Arepalli, S.; Hunter, R. L., A review of carbon
nanotube toxicity and assessment of potential occupational and environmental
health risks. Critical Reviews in Toxicology 2006, 36, (3), 189-217.
Langmuir, I., The constitution and fundamental properties of solids and liquids Part I
Solids. Journal of the American Chemical Society 1916, 38, 2221-2295.
Lee, M.; Baik, K. Y.; Noah, M.; Kwon, Y. K.; Lee, J. O.; Hong, S., Nanowire and
nanotube transistors for lab-on-a-chip applications. Lab on a Chip 2009, 9, (16),
2267-2280.
Lei, B.; Li, C.; Zhang, D.; Tang, T.; Zhou, C., Tuning electronic properties of In2O3
nanowires by doping control. Applied Physics a-Materials Science & Processing
2004, 79, (3), 439-442.
Li, C.; Curreli, M.; Lin, H.; Lei, B.; Ishikawa, F. N.; Datar, R.; Cote, R. J.; Thompson, M.
E.; Zhou, C. W., Complementary detection of prostate-specific antigen using
ln(2)O(3) nanowires and carbon nanotubes. Journal of the American Chemical
Society 2005, 127, (36), 12484-12485.
Li, C.; Lei, B.; Zhang, D. H.; Liu, X. L.; Han, S.; Tang, T.; Rouhanizadeh, M.; Hsiai, T.;
Zhou, C. W., Chemical gating of In2O3 nanowires by organic and biomolecules.
Applied Physics Letters 2003, 83, (19), 4014-4016.
199

Li, C.; Zhang, D.; Han, S.; Liu, X.; Tang, T.; Lei, B.; Liu, Z.; Zhou, C., Synthesis,
electronic properties, and applications of indium oxide nanowires. Molecular
Electronics Iii 2003, 1006, 104-121.
Li, C.; Zhang, D. H.; Han, S.; Liu, X. L.; Tang, T.; Zhou, C. W., Diameter-controlled
growth of single-crystalline In2O3 nanowires and their electronic properties.
Advanced Materials 2003, 15, (2), 143-146.
Li, C.; Zhang, D. H.; Lei, B.; Han, S.; Liu, X. L.; Zhou, C. W., Surface treatment and
doping dependence of In2O3 nanowires as ammonia sensors. Journal of Physical
Chemistry B 2003, 107, (45), 12451-12455.
Li, X.; Zhang, Y. P.; Kim, H. S.; Bae, K. H.; Stantz, K. M.; Lee, S. J.; Jung, C. Y.;
Jimenez, J. A.; Gardner, T. A.; Jeng, M. H.; Kao, C. H., Gene therapy for prostate
cancer by controlling adenovirus E1a and E4 gene expression with PSES
enhancer. Cancer Research 2005, 65, (5), 1941-1951.
Liang, X. G.; Chou, S. Y., Nanogap detector inside nanofluidic channel for fast real-time
label-free DNA analysis. Nano Letters 2008, 8, (5), 1472-1476.
Liao, H. I.; Olson, C. A.; Hwang, S. M.; Deng, H. Y.; Wong, E.; Baric, R. S.; Roberts, R.
W.; Sun, R., mRNA Display Design of Fibronectin-based Intrabodies That Detect
and Inhibit Severe Acute Respiratory Syndrome Coronavirus Nucleocapsid
Protein. Journal of Biological Chemistry 2009, 284, (26), 17512-17520.
Lieber, C. M.; Wang, Z. L., Functional nanowires. Mrs Bulletin 2007, 32, (2), 99-108.
Liu, F.; Bao, M.; Wang, K. L.; Li, C.; Lei, B.; Zhou, C., One-dimensional transport of
In2O3 nanowires. Applied Physics Letters 2005, 86, (21), 213101-213104.
Liu, R.; Barrick, J.; Szostak, J. W.; Roberts, R. W., Optimized Synthesis of RNA-Protein
Fusions for In Vitro Protein Selection. Methods in Enzymology 2000, 317, 268-
293.
Lud, S. Q.; Nikolaides, M. G.; Haase, I.; Fischer, M.; Bausch, A. R., Field effect of
screened charges: Electrical detection of peptides and proteins by a thin-film
resistor. Chemphyschem 2006, 7, (2), 379-384.
Maehashi, K.; Katsura, T.; Kerman, K.; Takamura, Y.; Matsumoto, K.; Tamiya, E.,
Label-free protein biosensor based on aptamer-modified carbon nanotube field-
effect transistors. Analytical Chemistry 2007, 79, (2), 782-787.
200

Mansfield, E.; Kar, A.; Hooker, S. A., Applications of TGA in quality control of
SWCNTs. Analytical and Bioanalytical Chemistry 2010, Article in Press, DOI
10.1007/s00216-009-3319-2.
Mattson, M. P.; Haddon, R. C.; Rao, A. M., Molecular functionalization of carbon
nanotubes and use as substrates for neuronal growth. Journal of Molecular
Neuroscience 2000, 14, (3), 175-182.
Minot, E. D.; Janssens, A. M.; Heller, I.; Heering, H. A.; Dekker, C.; Lemay, S. G.,
Carbon nanotube biosensors: The critical role of the reference electrode. Applied
Physics Letters 2007, 91, (9), 093507-093511.
Moonoosawmy, K. R.; Kruse, P., To Dope or Not To Dope: The Effect of Sonicating
Single-Wall Carbon Nanotubes in Common Laboratory Solvents on Their
Electronic Structure. Journal of the American Chemical Society 2008, 130, (40),
13417-13424.
Mor, G.; Visintin, I.; Feng, Z. D.; Longton, G.; Ward, D. C.; Alvero, A. B.; Lai, Y. L.;
Tenthorey, J.; Leiser, A.; Flores, R.; Yu, H.; Azori, M.; Rutherford, T.; Schwartz,
P. E., Diagnostic markers for early detection of ovarian cancer. Cancer
Biomarkers 2008, 4, (3), 190-191.
Moulton, S. E.; Minett, A. I.; Wallace, G. G., Carbon nanotube based electronic and
electrochemical sensors. Sensor Letters 2005, 3, (3), 183-193.
Mrksich, M., What can surface chemistry do for cell biology? Current Opinion in
Chemical Biology 2002, 6, (6), 794-797.
Nair, P. R.; Alam, M. A., Performance limits of nanobiosensors. Applied Physics Letters
2006, 88, (23), 233120-233123.
Nair, P. R.; Alam, M. A., Screening-limited response of nanobiosensors. . Los Alamos
National Laboratory, Preprint Archive, Condensed Matter 2007, 1-7.
Nair, P. R.; Alam, M. A., Design Considerations of Silicon nanowire biosensors. IEEE
Transactions on Electron Devices 2007, 54, 3400-3407.
Neff, P. A.; Serr, A.; Wunderlich, B. K.; Bausch, A. R., Label-free electrical
determination of trypsin activity by a silicon-on-insulator based thin film resistor.
Chemphyschem 2007, 8, (14), 2133-2137.
201

Neff, P. A.; Wunderlich, B. K.; Lud, S. Q.; Bausch, A. R., Silicon-on-insulator based thin
film resistors for quantitative biosensing applications. Physica Status Solidi a-
Applications and Materials Science 2006, 203, (14), 3417-3423.
Olson, C. A.; Liao, H. I.; Sun, R.; Roberts, R. W., nRna display selection of a high-
affinity, modification-specific phospho-IkBalpha-binding fibronectin. ACS
Chemical Biology 2008, 3, (3), 480-485.
Park, I. Y.; Li, Z. Y.; Li, X. M.; Pisano, A. P.; Williams, R. S., Towards the silicon
nanowire-based sensor for intracellular biochemical detection. Biosensors &
Bioelectronics 2007, 22, (9-10), 2065-2070.
Patolsky, F.; Timko, B. P.; Yu, G. H.; Fang, Y.; Greytak, A. B.; Zheng, G. F.; Lieber, C.
M., Detection, stimulation, and inhibition of neuronal signals with high-density
nanowire transistor arrays. Science 2006, 313, (5790), 1100-1104.
Patolsky, F.; Timko, B. P.; Zheng, G. F.; Lieber, C. M., Nanowire-based nanoelectronic
devices in the life sciences. Mrs Bulletin 2007, 32, (2), 142-149.
Patolsky, F.; Zheng, G.; Lieber, C. M., Nanowire sensors for medicine and the life
sciences. Nanomedicine 2006, 1, (1), 51-65.
Patolsky, F.; Zheng, G. F.; Hayden, O.; Lakadamyali, M.; Zhuang, X. W.; Lieber, C. M.,
Electrical detection of single viruses. Proceedings of the National Academy of
Sciences of the United States of America 2004, 101, (39), 14017-14022.
Patolsky, F.; Zheng, G. F.; Lieber, C. M., Nanowire-based biosensors. Analytical
Chemistry 2006, 78, (13), 4260-4269.
Patra, M. K.; Manzoor, K.; Manoth, M.; Negi, S. C.; Vadera, S. R.; Kumar, N.,
Nanotechnology applications for chemical and biological sensors. Defence
Science Journal 2008, 58, (5), 636-649.
Peng, X. H.; Wong, S. S., Functional Covalent Chemistry of Carbon Nanotube Surfaces.
Advanced Materials 2009, 21, (6), 625-642.
Pillai, S. K.; Ray, S. S.; Moodley, M., Purification of single-walled carbon nanotubes.
Journal of Nanoscience and Nanotechnology 2007, 7, (9), 3011-3047.
Poghossian, A.; Ingebrandt, S.; Abouzar, M. H.; Schoning, M. J., Label-free detection of
charged macromolecules by using a field-effect-based sensor platform:
Experiments and possible mechanisms of signal generation. Applied Physics a-
Materials Science & Processing 2007, 87, (3), 517-524.
202

Popov, A. M.; Lozovik, Y. E.; Fiorito, S.; Yahia, L., Biocompatibility and applications of
carbon nanotubes in medical nano robots. International Journal of Nanomedicine
2007, 2, (3), 361-372.
Raymo, F. M.; Yildiz, I., Luminescent chemosensors based on semiconductor quantum
dots. Physical Chemistry Chemical Physics 2007, 9, (17), 2036-2043.
Roberts, R. W.; Szostak, J. W., RNA-peptide fusions for the in vitro selection of peptides
and proteins. Proceedings of the National Academy of Sciences of the United
States of America 1997, 94, 12297-12302.
Rohde, R. D.; Agnew, H. D.; Yeo, W. S.; Bailey, R. C.; Heath, J. R., A non-oxidative
approach toward chemically and electrochemically functionalizing Si(111).
Journal of the American Chemical Society 2006, 128, (29), 9518-9525.
Rosenblatt, S.; Yaish, Y.; Park, J.; Gore, J.; Sazonova, V.; McEuen, P. L., High
performance electrolyte gated carbon nanotube transistors. Nano Letters 2002, 2,
(8), 869-872.
Rosi, N. L.; Mirkin, C. A., Nanostructures in biodiagnostics. Chemical Reviews 2005,
105, (4), 1547-1562.
Salvetat, J. P.; Bhattacharyya, S.; Pipes, R. B., Progress on mechanics of carbon
nanotubes and derived materials. Journal of Nanoscience and Nanotechnology
2006, 6, (7), 1857-1882.
Sampson, N. S.; Mrksich, M.; Bertozzi, C. R., Surface molecular recognition.
Proceedings of the National Academy of Sciences of the United States of America
2001, 98, (23), 12870-12871.
Schrenk, J. L.; Mann, K. R., Photochemical Studies of [(Eta-5-C5r5)M(Eta-6-Cot)]Pf6
and [(Eta-5-C5r5)M(Eta-4-1,5-Cot)L]Pf6 Complexes of Iron and Ruthenium.
Inorganic Chemistry 1986, 25, (11), 1906-1908.
Sheehan, P. E.; Whitman, L. J., Detection limits for nanoscale biosensors. Nano Letters
2005, 5, (4), 803-807.
Singh, S.; Kruse, P., Carbon nanotube surface science. International Journal of
Nanotechnology 2008, 5, (9-12), 900-929.
Sinha, N.; Ma, J. Z.; Yeow, J. T. W., Carbon nanotube-based sensors. Journal of
Nanoscience and Nanotechnology 2006, 6, (3), 573-590.
203

Stadtherr, K.; Wolf, H.; Lindner, P., An aptamer-based protein biochip. Analytical
Chemistry 2005, 77, (11), 3437-3443.
Star, A.; Gabriel, J. C. P.; Bradley, K.; Gruner, G., Electronic detection of specific
protein binding using nanotube FET devices. Nano Letters 2003, 3, (4), 459-463.
Star, A.; Han, T. R.; Gabriel, J. C. P.; Bradley, K.; Gruner, G., Interaction of aromatic
compounds with carbon nanotubes: Correlation to the Hammett parameter of the
substituent and measured carbon nanotube FET response. Nano Letters 2003, 3,
(10), 1421-1423.
Stern, E.; Klemic, J. F.; Routenberg, D. A.; Wyrembak, P. N.; Turner-Evans, D. B.;
Hamilton, A. D.; LaVan, D. A.; Fahmy, T. M.; Reed, M. A., Label-free
immunodetection with CMOS-compatible semiconducting nanowires. Nature
2007, 445, (7127), 519-522.
Stern, E.; Vacic, A.; Reed, M. A., Semiconducting Nanowire Field-Effect Transistor
Biomolecular Sensors. Ieee Transactions on Electron Devices 2008, 55, (11),
3119-3130.
Stern, E.; Wagner, R.; Sigworth, F. J.; Breaker, R.; Fahmy, T. M.; Reed, M. A.,
Importance of the Debye Screening Length on Nanowire Field Effect Transistor
Sensors. Nano Letters 2007, 7, (11), 3405-3409.
Takahashi, T. T.; Austin, R. J.; Roberts, R. W., mRNA display: ligand discovery,
interaction analysis and beyond. Trends in Biochemical Sciences 2003, 28, (3),
159-165.
Tamerler, C.; Sarikaya, M., Molecular biomimetics: nanotechnology and
bionanotechnology using genetically engineered peptides. Philosophical
Transactions of the Royal Society a-Mathematical Physical and Engineering
Sciences 2009, 367, (1894), 1705-1726.
Tang, T.; Han, S.; Jin, W.; Liu, X. L.; Li, C.; Zhang, D. H.; Zhou, C. W.; Chen, B.; Han,
J.; Meyyapan, M., Synthesis and characterization of single-crystal indium nitride
nanowires. Journal of Materials Research 2004, 19, (2), 423-426.
Tang, X. W.; Bansaruntip, S.; Nakayama, N.; Yenilmez, E.; Chang, Y. L.; Wang, Q.,
Carbon nanotube DNA sensor and sensing mechanism. Nano Letters 2006, 6, (8),
1632-1636.
Tans, S. J.; Verschueren, A. R. M.; Dekker, C., Room-temperature transistor based on a
single carbon nanotube. Nature 1998, 393, (6680), 49-52.
204

Tansil, N. C.; Gao, Z. Q., Nanoparticles in biomolecular detection. Nano Today 2006, 1,
(1), 28-37.
Tasis, D.; Tagmatarchis, N.; Georgakilas, V.; Prato, M., Soluble carbon nanotubes.
Chemistry-a European Journal 2003, 9, (17), 4001-4008.
Terrones, M.; Souza, A. G.; Rao, A. M., Doped carbon nanotubes: Synthesis,
characterization and applications. Carbon Nanotubes 2008, 111, 531-566.
Timko, B. P.; Cohen-Karni, T.; Yu, G. H.; Qing, Q.; Tian, B. Z.; Lieber, C. M., Electrical
Recording from Hearts with Flexible Nanowire Device Arrays. Nano Letters
2009, 9, (2), 914-918.
Trojanowicz, M., Analytical applications of carbon nanotubes: a review. Trac-Trends in
Analytical Chemistry 2006, 25, (5), 480-489.
Vecchi, P. A.; Alvarez, C. M.; Ellern, A.; Angelici, R. J.; Sygula, A.; Sygula, R.;
Rabideau, P. W., Synthesis and structure of a dimetallated buckybowl:
Coordination of one {Cp*Ru}(+) unit to each side of corannulene. Angewandte
Chemie-International Edition 2004, 43, (34), 4497-4500.
Vecchi, P. A.; Alvarez, C. M.; Ellern, A.; Angelici, R. J.; Sygula, A.; Sygula, R.;
Rabideau, P. W., Flattening of a curved-surface buckybowl (corannulene) by
eta(6) coordination to {Cp*Ru}(+). Organometallics 2005, 24, (19), 4543-4552.
Visintin, I.; Feng, Z.; Longton, G.; Ward, D. C.; Alvero, A. B.; Lai, Y. L.; Tenthorey, J.;
Leiser, A.; Flores-Saaib, R.; Yu, H.; Azori, M.; Rutherford, T.; Schwartz, P. E.;
Mor, G., Diagnostic markers for early detection of ovarian cancer. Clinical
Cancer Research 2008, 14, (4), 1065-1072.
Wang, W. U.; Chen, C.; Lin, K. H.; Fang, Y.; Lieber, C. M., Label-free detection of
small-molecule-protein interactions by using nanowire nanosensors. Proceedings
of the National Academy of Sciences of the United States of America 2005, 102,
(9), 3208-3212.
Yang, W. R.; Thordarson, P.; Gooding, J. J.; Ringer, S. P.; Braet, F., Carbon nanotubes
for biological and biomedical applications. Nanotechnology 2007, 18, (41),
412001-412013.
Yeo, W. S.; Yousaf, M. N.; Mrksich, M., Dynamic interfaces between cells and surfaces:
Electroactive substrates that sequentially release and attach cells. Journal of the
American Chemical Society 2003, 125, (49), 14994-14995.
205

Yousaf, M. N.; Houseman, B. T.; Mrksich, M., Using electroactive substrates to pattern
the attachment of two different cell populations. Proceedings of the National
Academy of Sciences of the United States of America 2001, 98, (11), 5992-5996.
Yousaf, M. N.; Houseman, B. T.; Mrksich, M., Turning on cell migration with
electroactive substrates. Angewandte Chemie-International Edition 2001, 40, (6),
1093-1096.
Yousaf, M. N.; Mrksich, M., Diels-Alder reaction for the selective immobilization of
protein to electroactive self-assembled monolayers. Journal of the American
Chemical Society 1999, 121, (17), 4286-4287.
Zakhartchouk, A. N.; Viswanathan, S.; Mahony, J. B.; Gauldie, J.; Babiuk, L. A., Severe
acute respiratory syndrome coronavirus nucleocapsid protein expressed by an
adenovirus vector is phosphorylated and immunogenic in mice. Journal of
General Virology 2005, 86, 211-215.
Zhang, D. H.; Li, C.; Han, S.; Liu, X. L.; Tang, T.; Jin, W.; Zhou, C. W., Electronic
transport studies of single-crystalline In2O3 nanowires. Applied Physics Letters
2003, 82, (1), 112-114.
Zhang, D. H.; Liu, Z. Q.; Li, C.; Tang, T.; Liu, X. L.; Han, S.; Lei, B.; Zhou, C. W.,
Detection of NO2 down to ppb levels using individual and multiple In2O3
nanowire devices. Nano Letters 2004, 4, (10), 1919-1924.
Zhang, G.-J.; Zhang, G.; Chua, J. H.; Chee, R.-E.; Wong, E. H.; Agarwal, A.;
Buddharaju, K. D.; Singh, N.; Gao, Z.; Balasubramanian, N., DNA Sensing by
Silicon Nanowire: Charge Layer Distance Dependence. Nano Letters 2008, 8, (4),
1066-1070.
Zhang, R.; Curreli, M.; Ishikawa, F.; Chen, P.; Zhou, C.; Thompson, M. E., Selective
functionalization of In2O3 nanowire mat device: the true "OFF" state. Manuscript
in Preparation 2010.
Zheng, G. F.; Patolsky, F.; Cui, Y.; Wang, W. U.; Lieber, C. M., Multiplexed electrical
detection of cancer markers with nanowire sensor arrays. Nature Biotechnology
2005, 23, (10), 1294-1301.
Zhu, B. L.; Ellern, A.; Sygula, A.; Sygula, R.; Angelici, R. J., eta(6)-coordination of the
curved carbon surface of corannulene (C20H10) to (eta(6)-arene)M2+ (M = Ru,
Os). Organometallics 2007, 26, (7), 1721-1728.

Abstract (if available)

Abstract Semiconducting nanomaterials are strong candidates for sensing applications because of their high surface to volume ratio. During my doctorate studies, I contributed to the development of nanobiosensors, sensor devices based on field effect transistors that utilize these nanomaterials as the sensing element in the device. I was particularly involved in the surface modification of such nanomaterials.

Linked assets

University of Southern California Dissertations and Theses

Conceptually similar

PDF

Surface functionalization of nanomaterials and development of field effect transistor nanobiosensors

PDF

Surface functionalization of nanomaterials and the development of nanobiosensors

PDF

One-dimensional nanomaterials: synthesis and applications

PDF

Applications of one-dimensional structured nanomaterials as biosensors and transparent electronics

PDF

Surface modification of parylene C and indium tin oxide for retinal and cortical prostheses

PDF

Synthesis and application of one-dimensional nanomaterials

PDF

Raman spectroscopy of carbon nanotubes under axial strain and surface-enhanced Raman spectroscopy of individual carbon nanotubes

PDF

One-dimensional nanomaterials for electronic and sensing applications

PDF

Graphene and carbon nanotubes: synthesis, characterization and applications for beyond silicon electronics

PDF

Nanomaterials for macroelectronics and energy storage device

PDF

The influence of CdSe surface ligands on hybrid inorganic/organic solar cell performance

PDF

Expanding the library of surface ligands for semiconductor nanocrystal synthesis and photovoltaic applications

PDF

Studies of nanosecond pulsed power for modifications of biomaterials and nanomaterials (SWCNT)

PDF

Novel nanomaterials for electronics, optoelectronics and sensing applications

PDF

Nanocrystal surface engineering as a route towards improved photovoltaics

PDF

Electrical and electrochemical properties of molecular and conducting polymer coated nanoscale field effect transistors

PDF

Semiconducting metal oxide nanostructures for scalable sensing applications

PDF

Ultrafast spectroscopic interrogation and simulation of excited states and reactive surfaces

PDF

Theoretical, experimental, device fabrication, and degradation studies of materials for optoelectronic devices

PDF

Development of novel technology platforms for blood based diagnosis and cellular characterization in cancer

Asset Metadata

Creator Curreli, Marco (author)

Core Title Surface modification of nanomaterials and developments of nanobiosensors

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Chemistry

Publication Date 02/12/2012

Defense Date 01/29/2010

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag cancer nanotechnology,nanomaterial surface modification,nanotube biosensors,nanowire biosensors,OAI-PMH Harvest

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Thompson, Mark E. (committee chair), Brutchey, Richard L. (committee member), Mansfeld, Florian B. (committee member)

Creator Email curreli@usc.edu,marcocurreli@gmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-m2848

Unique identifier UC1170282

Identifier etd-Curreli-3473 (filename),usctheses-m40 (legacy collection record id),usctheses-c127-296288 (legacy record id),usctheses-m2848 (legacy record id)

Legacy Identifier etd-Curreli-3473.pdf

Dmrecord 296288

Document Type Dissertation

Rights Curreli, Marco

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Repository Name Libraries, University of Southern California

Repository Location Los Angeles, California

Repository Email cisadmin@lib.usc.edu