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Surface functionalization of nanomaterials and the development of nanobiosensors
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Content
SURFACE FUNCTIONALIZATION OF NANOMATERIALS
AND THE DEVELOPMENT OF NANOBIOSENSORS
by
RUI ZHANG
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)
December 2012
Copyright 2012 RUI ZHANG
ii
All Hail to Alma Mater
To thy glory we sing;
All Hail to Southern California
Loud let thy praises ring;
Where Western sky meets Western sea
Our college stands in majesty;
Sing our love to Alma Mater,
Hail, all hail to thee!
iii
ACKNOWLEDGMENTS
I would like to express my deepest gratitude to my advisor, Professor Mark
Thompson, for his excellent guidance, caring, patience, and providing me with an
excellent atmosphere for doing research.
I would like to thank the collaborating professors during my PhD research, Prof.
Chongwu Zhou, Richard Cote, Ram Datar and Julio Camarero, for helping me develop
my knowledge in electrical engineering and biology.
I would like to thank all the former and current team members in biosensing team,
especially Marco Curreli, who has been a great mentor since my first day in this field.
In addition, I would like to extend my warmest thanks to all the group members in
Thompson Research Group and the collaborators that I have worked with on my research
projects.
Finally, I want to thank my parents and my boyfriend, Chao Wu. Your support
and love means everything to me.
iv
TABLE OF CONTENTS
Epigraph ii
Acknowledgments iii
List of Tables vi
List of Figures vii
Abstract x
Chapter One: Introduction
1.1 One-dimensional (1D) nanomaterials in biosensors 1
1.1.1 Point of care testing (POCT) devices 1
1.1.2 Synthesis of 1D nanomaterials 3
1.1.3 1D nanomaterial-based biosensors 4
1.2 Field-effect transistor (FET) nnaobiosensors 5
1.2.1 Operating mechanism 6
1.2.2 FET fabrication 8
1.2.3 Surface functionalization 10
1.2.4 Important factors during sensing 12
1.3 Current research trends and challenges in nanobiosensing 16
1.3.1 Mass production of 1D nanomaterials 16
1.3.2 Multiplexing 17
1.3.3 Sensing in physiological samples 19
1.4 Chapter one reference 20
Chapter Two: Selective, electrochemically activated biofunctionalization of In
2
O
3
nanowires using an air-stable surface modifier
2.1 Introduction 24
2.2 Results and discussion 27
2.2.1 Electrochemical behavior of DMP-PA 27
2.2.2 Stability test of DMP-PA in air 29
2.2.3 Selective functionalization of In
2
O
3
NW mat with DNA oligos31
2.3 Chapter conclusion 34
2.4 Experimental section 34
2.5 Chapter two reference 36
Chapter Three: Field-effect transistors based on ZnO nanomaterials synthesized from a
hydrothermal method
3.1 Introduction 39
3.2 Results and discussion 40
3.2.1 Synthesis and characterization of ZnO nanomaterials 40
3.2.2 FET fabrication and electrical performance 44
3.2.3 Statistical study of the effect of annealing on FET performance47
v
3.2.4 Device uniformity 48
3.2.5 Surface functionalization of ZnO products 50
3.3 Chapter conclusion 52
3.4 Experimental section 53
3.5 Chapter three reference 54
Chapter Four: Construction of nanobiosensors using cyclotides as capturing probes
4.1 Introduction 56
4.2 Results and discussion 58
4.2.1 Construction of nanobiosensors using MCoTI-II 58
4.2.2 Click chemistry condition 59
4.2.3 Surface click chemistry 62
4.2.4 Binding affinity of surface bound MCoTI-II towards trypsin 63
4.2.5 Real-time sensing of trypsin using MCoTI-II derivatized In2O3 NW
sensors 65
4.3 Chapter conclusion 67
4.4 Experimental section 68
4.5 Chapter three reference 70
Chapter Five: PolySilicon microribbon biosensors: surface functioanlization and sensing
applications
5.1 Introduction 72
5.2 Results and discussion 74
5.2.1 Device fabrication and performance 74
5.2.2 pH sensing 77
5.2.3 Surface functionalization of polysilicon with native oxide layer79
5.2.4 Biomarker sensing 83
5.2.5 Surface functionalization of polysilicon without native oxide layer
84
5.3 Chapter conclusion 86
5.4 Experimental section 87
5.5 Chapter three reference 90
Bibliography 91
vi
LIST OF TABLES
Table 3.1: Contact angle of ZnO nanomaterial films functionalized
with molecule A, B and C 50
Table 4.1: Reaction yield of molecule A and alkyne-terminated
ferrocene under different reaction conditions. 80
vii
LIST OF FIGURES
Figure 1.1: POCT device that consists of a bio-recognition layer on a
transducer attached to an analytical output 2
Figure 1.2: Size of several nanomaterials is compared to the size of
some biological molecules, such as nucleic acids, proteins,
virus, and cells 3
Figure 1.3: Typical structure of a FET sensor 6
Figure 1.4: Mechanism to modulate the conductance of a p-type
nanomaterial-based FET (holes as the main charge carriers) 8
Figure 1.5: Images of FETs fabricated in our research group 9
Figure 1.6: Chemical pathways used to anchor biological molecules to
different nanomaterial surfaces 11
Figure 1.7: Impact of Debye screening on streptavidin sensing using a
SiNW-FET 13
Figure 1.8: Reagent delivery systems 16
Figure 2.1: Oxidation process for HQ and para-dimethoxybenzene
derivatives 26
Figure 2.2: Electrochemical characterization of DMP-PA on ITO
electrodes 29
Figure 2.3: Stability test for DMP-PA 31
Figure 2.4: Selective functionalization of In
2
o
3
NW mat with DNA
oligos 33
Figure 3.1: Characterization of ZnO nanorods grown on Si substrate 41
Figure 3.2: Characterization of hydrothermal ZnO products 43
Figure 3.3: Characterization of hydrothermal ZnO products using TEM 44
Figure 3.4: FETs based on as-grown ZnO products 45
Figure 3.5: Measured characteristics of a typical FET made with
annealed ZnO products 47
viii
Figure 3.6: Statistics of 17 annealed devices and 13 unannealed ones 48
Figure 3.7: ZnO products synthesized without SDS. This batch has
fewer belts 49
Figure 3.8: Structure of molecule A, B and C 50
Figure 3.9: IR spectrum of ZnO nanomaterials functionalized with (a)
molecule B, (b) 3-phosphonopropionic acid and (c)
3-aminopropyl phosphonic acid. 51
Figure 4.1: Primary and tertiary structure of trypsin inhibitor (MCoTI-
II). Conserved cysteine residues are marked in yellow and
disulfide connectivities in red. The circular backbone
topology is shown with a blue line. 57
Figure 4.2: (a) Synthesis of bifunctional linker B. (b) Surface chemistry
strategy for immobilizing alkyne-terminated MCoTI-II onto
In2O3 NWs. 59
Figure 4.3: Reaction between molecule A and alkyne-terminated
ferrocene 60
Figure 4.4: HPLC traces of reaction mixtures with (a) and without (b)
TBTA. Reactants: 100 μM molecule A and 100 μM
MCoTI-II. Catalyst: 1 mM CuSO4 and 2 mM sodium
ascorbate. 61
Figure 4.5: (a) Surface functionalization of ITO-coated glass with
molecule B and ethynylferrocene. (b) CV traces for azide-
modified ITO (black), after coupled with ethynylferrocene
(red) and a bare ITO exposed to ethynylferrocene (blue). (c)
Surface functionalization of In2O3 NW mats with molecule
B and MCoTI-II. (d) and (e) are fluorescent images of the
top and bottom sample illustrated in (c), respectively. 63
Figure 4.6: (a) Surface derivatization strategy for Biacore CM5 sensor
chip. (b) Kinetic analysis sensorgrams for surface bound
MCoTI-II and trypsin. 65
Figure 4.7: (a) Schematic view of a MCoTI II derivatized In2O3 NW
sensor. (b) Schematic view of the sensing setup. (c) Real
time sensing result for trypsin using a MCoTI II derivatized
In2O3 NW sensor. 67
ix
Figure 5.1: (a)-(d) show the fabrication process of polysilicon
nanoribbon FETs. (e) and (f) is a SEM image of a group of
6 polysilicon nanoribbons and a photographic image of
hundreds of polysilicon nanoribbon sensors fabricated on a
3” wafer, respectively. 75
Figure 5.2: The IDS v.s. VDS and IDS v.s. VGS characteristics for
devices with 1×1017, 5×1017 and 1×1018 doping
concentration. 77
Figure 5.3: Real-time pH sensing using a polysilicon FET at pH range
of (a) pH 4 ~10 and (b) pH 7.2 ~ 8. 79
Figure 5.4: (a) Scheme of surface functionalization of polysilicon using
a biotin derivative. (b) XPS high-resolution spectra of the N
1s region after each step of functionalization. (c)
Fluorescent image of the biotin-functionalized polysilicon
after exposed to fluorescent labeled streptavidin. (d)
Fluorescent image of a polysilicon wafer as control. The
wafer was directly exposed to amine-PEG-biotin and then
fluorescent streptavidin right after pre-cleaning step. 82
Figure 5.5: Real-time sensing of CA-125 using polysilicon nanoribbon
biosensors. Signals from three sensors were monitored
simultaneously. A non-target protein, BSA, was added to
the sensing media at the end of the sensing experiment to
demonstrate the sensors’ selectivity. This part of sensing
signal is shown in inset. 84
Figure 5.6: (a) Scheme of surface functionalization strategy of
hydrogen-terminated polysilicon. (b) XPS survey spectra of
two hydrogen-terminated polysilicon samples after being
immersed in 5 hexenoic acid in N2 for 4 hours with (red)
and without (black) UV illumination. (c) and (d) are
fluorescent images of these two samples after being treated
with amine-PEG-biotin and fluorescently labeled
streptavidin. (c) is the image of the UV illuminated sample
and (d) is the image of the sample kept in dark 86
x
ABSTRACT
In the past 20 years, material scientists and engineers have progressively
miniaturized the materials that constitute the building blocks of various biomedical
devices. This progressive downscaling has led to the creation of materials with at least
one critical dimension falling within the 1-100 nm range. These nanomaterials have been
considered as ideal candidates for biosensing applications due to their high
surface-to-volume ratio and small sizes. A diversity of sensor architectures has been
designed and fabricated during the last decade that utilizes different nanomaterials as
sensing elements. Among them, sensors based on FETs have drawn increasing attention
because of their capability of performing rapid and label-free detections. Since the first
demonstration of FET-based biosensors in 2001, people have achieved detections of
proteins, oligonucleotides and viruses with high sensitivity and selectivity. However, to
facilitate the widespread adoption of nanobiosensing technology, researchers still need to
address a few challenges, including multiplexing, cost efficiency and signal
reproducibility. This dissertation tries to tackle these challenges by improving
nanosensor fabrication techniques and developing novel surface functionalization
methods.
Chapter 1 briefly introduces the fundamentals of nanobiosensors, including
nanomaterial synthesis, device fabrication and sensing setup, and discusses current
challenges in the nanobiosensing field.
Chapter 2 describes a newly designed electroactive surface modifier that can be
used in selective biofunctionalization of nanomaterials. This molecule offers promising
control over the surface reaction of each sensor in an array and can be considered as a
xi
key element in the fabrication of high-density biosensor arrays for multiplexed
biosensing in the future.
Chapter 3 focuses on sensing platforms based on solution grown ZnO
nanostructures. Mass production of ZnO nanowires and nanobelts are prepared using a
low cost, low temperature hydrothermal method, and used to fabricate back-gated FETs.
By applying a post-synthesis annealing step to the ZnO products, we adjust the doping
level of the nanowires and nanobelts, and thus significantly improve their electrical
properties. These FETs shows comparable performance with those based on ZnO
nanostructures synthesized via vapor-phase approaches.
Chapter 4 introduces a new class of small affinity binding agents, cyclotides, as
capturing probes in nanobiosensing. These backbone-cyclized polypeptides with a
disulfide-stabilized core are chemically more stable than conventional antibodies and can
be produced in relatively large quantities at low cost. A cyclotide, MCoTI-II, was
integrated with In
2
O
3
NW mats and sensors for trypsin sensing in this chapter.
In the end, chapter 5 studies top-down polysilicon nanoribbon sensors and their
sensing applications. Top-down sensors are believed to have better device uniformity
and thus can yield more reliable sensing signals. In this chapter, polysilicon nanoribbon
FETs are fabricated using a simple two-mask photolithography method on a wafer scale
and functionalized with and without the native oxide coating. The pH sensing and
biosensing performed with these sensors demonstrates their promising sensitivity and
selectivity.
1
CHAPTER ONE: Introduction
1.1 One-dimensional (1D) nanomaterials in biosensors
1.1.1 Point of care testing (POCT) devices
The existing health care system is focused on treating diseases rather than
preventing them. Patients are generally not tested until physiological symptoms are
present. When they do get tested, the results often take several days and can be
inconclusive if the disease is at an early stage. In order to facilitate the diagnostics
process and make tests more readily available for patients, the concept of “point of care
testing” (POCT) has been brought up and developed in recent years.
1,2
POCT is defined
as the medical testing at or near the site of patient care.
3
The major driving force behind
POCT is to provide convenient tests to patients and deliver timely results to the health
care team. This increases the chances that a disease is diagnosed at its early stage and
allows immediate clinical decisions to be made.
POCT is often accomplished through the use of low cost, portable, handheld
testing devices and kits. They are expected to make rapid and precise diagnosis using
only small amount of samples (e.g. a drop of blood for glucose test). A typical POCT
device normally contains three components (Figure 1.1):
4
(1) a biological recognition
element (e.g. an antibody), (2) a material or matrix that transduces the bio-recognition
events into detectable signals and (3) a detector. For a POCT device to be workable,
there are several requirements that need to be satisfied.
5
First, an easy-to-construct and
reliable interface is required to output reproducible signals. Second, the detection method
needs to be highly sensitive and specific. Third, the sensing element (transducing
material) is ideally to be cost-efficient and miniaturized. Finally, due to the complexity
2
of biological systems, especially the human body, clinical diagnosis usually requires
simultaneous detection of multiple disease markers in order to accurate results.
Therefore, the next generation of POCT devices will require the capability to be operated
in a multiplexed manner.
Among the various types of materials and platforms for current and potential
POCT devices, 1D nanomaterials are recognized as ideal candidates due to their unique
electronic, optical, chemical and mechanical properties, which are intrinsically associated
with their low dimensionality and the quantum confinement effect.
6
1D nanomaterials
have a high surface-to-volume ratio. 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 1D nanomaterials
highly sensitive in molecular sensing applications. The small size of these nanomaterials
is another important feature that makes them ideal candidates for POCT devices. They
are comparable in size with most biological entities, such as proteins, nucleic acids, cells,
Figure 1.1 POCT device that consists of a bio-recognition layer on a transducer
attached to an analytical output.
3
viruses, etc. (as shown in Figure 1.2), making them the ideal interface materials between
biological molecules and scientific instruments. Also, their extreme smallness would
allow packing of a huge number of sensing elements into a small chip of an array device,
which can be used in multiplexed sensing of a panel of disease markers.
1.1.2 Synthesis of 1D nanomaterials
Various methods and technologies have been developed to fabricate 1D
nanomaterials. These methods can be grouped into two main categories: top-down and
bottom-up. Top-down fabrication starts with bulk materials and reduce the materials
dimensions using various techniques to cut, pattern, etch and shape these materials into
the desired geometry and order.
7
These top-down techniques can either be e-beam
lithography, photolithography combined with size-reducing strategies, such as the self-
limiting oxidation, or a “nano-dimension” transfer method such as the super lattice NW
pattern transfer method. Top-down techniques offer an excellent control over the
dimension and orientation of 1D nanomaterials. Disadvantages include high costs and a
slow rate of production. In contrast, the bottom-up approach involves preparing 1D
Figure 1.2 Size of several nanomaterials is compared to the size of some biological
molecules, such as nucleic acids, proteins, virus, and cells.
4
nanomaterials from molecular precursors, rather than starting with the bulk materials.
For example, chemical vapor deposition (CVD) have been widely used to synthesize
carbon nanotubes (CNTs),
8,9
silicon NWs
10,11
and metal oxide NWs
12,13
on catalytically
patterned substrates. Hydrothermal and electrochemical methods have also been used to
produce NWs from precursor solutions. The bottom-up approach is playing an
increasingly important role because of its capability to make much smaller features
compared to the top-down approach. However, these nanomaterials usually grow with
random orientation and are characterized by a distribution of lengths and diameters.
7
The
variation in dimension can impose limits on bottom-up-based sensors because of poor
device uniformity and low fabrication yields.
1.1.3 1D nanomaterial-based Biosensors
In the past few decades, people have developed various types of biosensors based
on 1D nanomaterials. According to the detection methods, they can be classified into
three groups: optical, electrochemical and electronic sensors.
Optical sensors are mostly based on CNTs. Single walled carbon nanotubes
(SWNTs) can serve as optical materials in biosensing applications due to the unique
optical properties resulted from the sharp densities of electronic states at the van Hove
singularities.
14
SWNTs have been used as an optical label in Raman-based biosensing
and the detection limit was observed down at 1fM for protein analyte.
15
SWNTs can also
be designed as signal transduction substrates in fluorescence biosensors utilizing their
capability of quenching a variety of fluorophores.
16,17
The second type of sensors,
electrochemical sensors, have traditionally received the major share of the attention in
5
POCT device development. Glucose testing device, one of the most successful POCT
devices in today’s market, is based on electrochemical sensing systems. In recent
researches, CNTs and NWs-based enzyme electrodes have been used to build
electrochemical sensors for glucose, cholesterols and DNAs.
18-20
As discussed above, the electrical properties of 1D nanomaterials are strongly
influenced by minor perturbations due to their high surface-to-volume ration and tunable
electron transport properties. Compared to 2D thin films where binding to the surface
leads to depletion or accumulation of charge carriers only on the surface, the charge
depletion or accumulation in the 1D nanomaterials takes place in the “bulk” of the
structure and thus gives rise to larger changes in the electrical properties. This advantage
of 1D nanomaterials provides a sensing modality for label-free and direct electrical
readout when used in electrical sensors. This part will be discussed in detail in section
1.2.
1.2 Field-effect transistor (FET) nanobiosensors
A diversity of sensor architectures has been designed and fabricated during the
past few decades that utilizes different nanomaterials as a sensing element (cantilevers,
quantum dots, nanotubes, NWs, nanobelts, nanogaps, and nanoscale films).
21-24
Some of
these sensing devices, such as those based on cantilevers and quantum dots, are highly
specific, ultrasensitive, 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 FET can
6
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, ultrasensitive, and label-free detection of the desired biomolecule.
25-28
1.2.1 Operating mechanism
An FET sensor has the structure of a common three-electrode transistor, where
the source and drain electrodes bridge the semiconductor channel and the gate electrode
modulates the channel conductance. The typical structure of an FET sensor is illustrated
in Figure 1.3. In the case of FET nanosensors, the semiconductor channel is made of a
nanomaterial and is used as the “sensing” component of the device. Semiconductor
channels can be fabricated using several nanomaterials, including CNTs and NWs. In
order to provide selectivity toward a unique analyte, a specific recognition group (also
called a receptor, ligand, or probe) is anchored to the surface of the semiconductor
channel. This receptor is typically chosen to recognize its target molecule (also called
analyte) with a high degree of both specificity and affinity.
7
The semiconductor channel has a uniform conductance determined by the main
carrier density in the nanomaterials (holes for a p-type semiconductor or electrons for an
n-type semiconductor). The carrier density is proportional to the conductance of the
channel, which can be determined from the source-drain current of the device. Any
change in the current can be related to a change in conductance of the channel. When a
charged analyte molecule binds to a receptor anchored on the nanomaterial, an electric
field created on the surface exerts an effect both inside and outside the semiconductor
channel.
29
If the bound analyte molecule carries a charge opposite to the main carriers in
the FET, then charge carriers will accumulate under the bound analyte, thus causing an
increase in the device conductivity. This mechanism is shown in Route A in Figure 1.4,
where a negatively charged molecule such as DNA binds to the p-type channel, causing a
buildup of hole carriers, thus resulting in an increase in conductivity. In contrast,
analytes with molecular charges same as that of the main carriers in the FET lead to
depletion of main carriers beneath the bound analyte, causing a decrease in conductivity.
The latter case is shown in Route B in Figure 1.4, where a positively charged molecule,
Figure 1.3 Typical structure of a FET sensor.
8
such as a protein below its isoelectric point, depletes the carriers upon binding to the
channel.
1.2.2 FET fabrication
Once the nanomaterials have been prepared, the source, drain, and gate electrodes
are deposited to complete the structure of the FET. Our research group has been using Si
substrate as the back gate electrode. In the case of bottom-up NWs, the NWs are
randomly dispersed on the substrate and metal source and drain electrodes are deposited
on the insulating layer (SiO
2
of 500 nm) on top of the NWs to define the channel length
and width of the FET (shown in Figure 1.5 a and b). It has been reported that the device
Figure 1.4 Mechanism to modulate the conductance of a p-type nanomaterial-based
FET (holes as the main charge carriers). The source-drain current of the channel is
monitored against time. In route A, when a negatively charged target binds to the
receptor anchored on the nanomaterial, the charge carriers will accumulate under the
bound analyte, thus causing an increase in the device conductivity and source-drain
current. In route B, the binding of a positively charged target leads to depletion of
charge carriers beneath the bound analyte, causing a decrease in conductivity and
source-drain current.
9
dimensionality directly affects the response time
30
and the sensitivity
31
of sensors. A
common channel length is on the order of 2-10 μm. In the case of top-down poly-silicon
microribbons, ribbons with leads are patterned with uniform width and length at
designated locations and metal electrodes are deposited on top of the leads for the
purpose of electrical connections (Shown in Figure 1.5 c and d).
A passivation layer on the source and drain metal electrodes is also an important
component of a FET for biosensing applications. It can help avoid complications during
measurements caused by corrosion, electrochemical reactions or change in metal work
function due to the non-specific binding of analytes. A widely used passivation approach
is to deposite a Si
3
N
4
layer on metal electrodes.
32-34
Figure 1.5 Images of FETs fabricated in our research group. (a) Optical image of an
In
2
O
3
NW FET with interdigitated electrodes. (b) Scanning electron microscope
(SEM) image of the same FET at a higher magnification. (c) Optical image of a group
of 6 poly-silicon microribbon FETs. (d) Optical image of poly-silicon microribbon
FETs at a higher magnification.
10
1.2.3 Surface functionalization
An as-fabricated FET device will not have the desired molecular recognition
properties. The surface of the sensing element (nanomaterial) needs to be modified so
that the device acquires specific recognition toward a desired analyte. This selectivity is
typically achieved by anchoring a specific recognition group to the surface of
nanomaterials. A bifunctional linker molecule with two chemically different termini is
used to help anchor the receptor molecules to the nanomaterial surface. In my doctoral
research, I focused on the surface functionalization of Si nanostructures and metal oxide
NWs.
In the case of Si nanostructures, the linker molecule of choice depends on whether
or not the wire has an oxide coating. A variety of linker molecules have been designed to
bind to the native SiO
2
layer on the surface. Alkoxysilane derivatives, such as
3-(trimethoxysilyl)propyl aldehyde (Figure 1.6a), 3-aminopropyltriethoxysilane and 3-
aminopropyldimethylethoxysilane are the most widely used linkers.
28,35-37
The
Si-methoxide or Si-ethoxide reacts with the surface OH group, anchoring the linker
molecule to the silicon oxide surface and creating a monolayer terminated with aldehyde
or amine groups. These groups can then react with amine or carboxylic acid groups that
are commonly present in biological capture probes. As for H-terminated Si surfaces, two
methods have been employed to functionalize the surface for further bioconjugation.
Several research groups use UV light to rapidly photo dissociate the Si-H bond to
generate radical species on the Si surface (Figure 1.6b). These radicals can subsequently
react with terminal olefin groups on linker molecules, thus forming stable Si-C bonds at
the Si surface.
34,38,39
The linker molecules usually carry a protected amine terminal,
11
which can be used to attach biological probes after deprotection. The other method,
developed by Nathan Lewis, uses a two-step chlorination/alkylation reaction to form Si-C
bond on the surface (Figure 1.6c).
40,41
The Si-H surface is first chlorinated to form Si-Cl
bond and then the surface was treated by an allyl Grignard. The resulted allyl surface can
be used for further bioconjugation.
42
In the case of metal oxide nanomaterials, a good choice for a linker molecule is
the one that is terminated with a group capable of forming a nonhydrolizable conjugate,
such as siloxides or phosphonates. Phosphonic acid was found to bind strongly to In
2
O
3
OH
Si
O
O
O
O
OH
OH
Si
O
O
O
HN
Si
O
O
O
O
H H H
NH
O
O
8
NH
O
O
8 NH
2
8
HN
8
O
HN
8
NH
4
H H H Cl Cl Cl
H H H
N N
N
n
PCl
5
(a)
(b)
(c)
Figure 1.6 Chemical pathways used to anchor biological molecules to different
nanomaterial surfaces. (a) Si surface coated with native oxide. (b) H-terminated Si
surface functionalized with an Olefin. (c) H-terminated Si surface functionalized with
the chlorination/alkylation method.
12
NWs and ZnO NWs.
43-46
Silane and carboxylic acid have been employed in the surface
functionalization of ZnO nanomateirals.
47-49
Once anchored on the surface, these linker
molecules can be used to attach biological capture probes.
1.2.4 Important factors in sensing
A FET device can now be used as a biosensor after the surface functionalization
with capturing probes. Besides the FET’s electrical performance, we need to pay
attention to a few factors that can significantly affect the sensing results: the choice of
sensing media (buffers), the non-specific binding of background proteins and the reagent
delivery setup.
Buffer: 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 a good model for human serum, which, like PBS, has
a pH value of 7.40 and 0.15 M electrolyte.
An important parameter that influences the device performance is the Debye
length ( ). The Debye length is defined as the maximum distance at which an external
charge can influence the NW carrier concentration.
50
The value of Debye length (in
nanometers) in water can be calculated with good approximation using the formula
, where is the ionic strength of the buffer solution.
51
In aqueous media,
the Debye length decreases rapidly with an increase in the ionic strength.
52
When
performing sensing 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.,
13
5-10 nm). Therefore, at such a short , the electrolytes present in the buffer screen the
charges carried by the analyte, resulting in a smaller NW conductance change. When
sensing is conducted at a lower ionic strength, a longer Debye length allows the detection
of analyte binding happened several nanometers away from the NW surface, resulting in
a larger NW conductance change. This effect has been demonstrated by Mark Reed and
coworkers using the well-studied biotin-streptavidin (SA) couple.
36
As shown in Figure
1.7, the sensor response was monitored at different buffer ionic strengths when a biotin
functionalized Si NW FET was exposed to the same SA solution. The binding signal
decreased with an increase in ionic strength. 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.
52
An alternative approach is to use smaller sized
capturing probes that can bring the analyte binding closer to the NW surface.
Figure 1.7 Impact of Debye screening on streptavidin sensing using a SiNW-FET.
27
(a) Schematic (not to scale) showing from the sensor surface at different buffer
ionic strength. The purple and red symbols represent receptors and targets,
respectively. (b) Biotin-functionalized sensor response to varying buffer ionic
concentrations with (red) and without (black) streptavidin addition at time = 0.
14
Non-specific binding: The analyte samples in clinical diagnosis are normally
physiological samples such as blood or urine, which contains not only the target molecule
but other high abundant proteins. These background proteins tend to non-specificly
absorb onto the NW surface and block the binding site on capturing probes. Driving
forces for protein adsorption have demonstrated the importance of hydrophobic
interaction, electrostatic attraction, van der Waals and hydrogen bonding.
53
Elimination
of protein adsorption requires the suppression of all these attractive forces between
proteins and surface. A common approach for blocking the adsorption of proteins is to
immobilize polymers in the form of well-solvated brushes on the surface. The polymer
layer shields the surface, introducing a high activation barrier for the proteins to adsorb.
54
Polyethylene glycols (PEGs) have been successfully employed to minimize
unspecific interaction between proteins and inorganic surfaces due to its excellent protein
repellent properties.
55-57
A range of methods have been employed for the immobilization
of PEG onto surfaces and these can be broadly classified as either physisorptive or
chemisorptive.
55,56
In our research, we use tween-20, a branched molecule with 20 PEG
segments, to non-covalently passivate In
2
O
3
NW surface. Using this molecule, we were
able to significantly decrease the non-specific binding of background proteins and
improve the sensor performance.
33
Reagent delivery: Another important factor to be taken into consideration is the
system used to deliver the analyte solution. 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,
15
two main methods have been utilized for such a delivery: microfluidic
channels
7,10,28,34,35,37
and mixing cells,
36,38,39,43
each having its advantages and
disadvantages.
A microfluidic channel is usually made of molded elastomer such as
polydimethylsiloxane (PDMS) with injection and drain channels, as shown in Figure
1.8(a) and (b). The microfluidic devices are placed on the top of the nanosensor so that
the solution can be directed over the NWs. 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. However, Laminar microfluidic flow can partially restricts the ability of
molecules to reach the nanosensor surface, 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.
31
Another disadvantage of PDMS
channels is caused by the highly hydrophobic sidewalls that can absorb biomolecules.
Another popular delivery method, shown schematically in Figure 1.8(c), utilizes a
mixing cell (also called solution chamber). This cell, typically made of Teflon, 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 1.8(d), has been designed
by Stern et al.
38
In this setup, injection of the solution tangential to the NW-FET sensor
significantly decreased the detection response times compared to those observed in
NW-FETs that used microchannels for the detection of similar target molecules.
34
16
1.3 Current research trends and challenges in nanobiosensing
Despite the rapid progress in biosensor development, clinical applications of
biosensors are still rare, with glucose monitor as an exception. This is in sharp contrast to
the urgent need in small clinics and POCT devices. For biosensors based on 1D
nanomaterials, a few challenges need to be addressed to facilitate the widespread
adoption of nanobiosensing technology.
1.3.1 Mass production of 1D nanomaterials
In order to commercialize 1D nanomaterial-based biosensors as POCT devices, it
is crucial to be able to produce these sensors massively at a relatively low price. The first
step is to scale up the synthesis of 1D nanomaterials and cut the cost at the same time.
Currently, 1D nanomaterials, including Si NWs, metal oxide NWs and CNTs, are
Figure 1.8 Reagent delivery systems. (a) Microfluidic channels can be used to
precisely deliver a small volume of analyte above the NW sensor. (b) View along the
flow direction. (c) Side-view of a mixing cell placed over a nanosensor chip. Analytes
are added to the buffer solution inside the cell and allowed to diffuse to the sensor
surface. (d) a system with injection and drain valves facilitate the analyte transport.
17
commonly produced by vapor-phase synthesis methods.
8-13
These methods usually
require high temperature and complicated instrumentation, but give comparatively low
product yields. In recent years, people have turned their interest to solution-phase
synthesis techniques including hydrothermal method and electrochemical deposition.
58,59
These approaches are attractive because of the low growth temperature and potential of
producing high density of nanomaterials. However, one major drawback of
solution-phase synthesized nanomaterials is that they usually have poor electrical
properties compared to the conventional gas-phase synthesized ones due to the higher
amount of defects and impurities introduced during the synthesis process. It is highly
desirable to improve the electrical properties of solution-phase synthesized nanomaterials
by modifying synthesis methods or using post-synthesis treatments. One of my research
projects is to enhance the conductance and device uniformity of hydrothermal
synthesized ZnO NWs-based FETs using a post-synthesis annealing step.
1.3.2 Multiplexing
Due to the complexity of biological systems, especially the human body, a single
biomarker alone is not effective enough for accurate diagnosis. Medical decision based
on single biomarker usually has a high possibility of false positive and false negative.
Recent research shows that combination of multiple biomarkers generates improved
accuracy compared to single biomarker.
60,61
This fact brings up the importance of
multiplexing assay of biomarkers. An ideal biosensing technology should be capable of
simultaneous detection of a combination of biomarkers.
18
To construct sensor arrays for multiplexed biosensing, the sensors must be
selectively functionalized with different capturing probes against their designated
analytes. Efforts have been made to achieve the selective functionalization of
nanomaterial-based devices, by using microfluidic chips, microspotting techniques
28,62
and electroactive monolayers.
44,63
Lieber and coworkers have demonstrated the highest
level of multiplexing to date for NW-FET sensors in a simultaneous assay for three
cancer markers with a detection of 0.9 pg/mL in desalted but undiluted serum samples.
28
Monoclonal antibodies specific for each of the targets were spotted onto different
NW-FETs. Samples solutions were delivered through microfluidic channels and the
electrical signal from each FET was monitored in real time during exposure to each of the
targets. Compared to the microspotting technique, the use of electroactive monolayers
possesses a unique advantage because it is only limited by the ability to electronically
address the individual sensors.
63
In this method, the key step is to design a bifunctional
molecule, which bears a NW-anchoring group on one end and an electroactive moiety on
the other. After being covalently linked to the NWs, the molecule can be activated from
the chemically inert “OFF” state to its “ON” state by applying an external voltage to
device electrodes. In the “ON” state the electroactive moiety reacts with the desired
capture probe to covalently anchor it to the NW surface. Several groups including
ourselves are currently working on developing multiplexed sensor arrays using this
technique.
19
1.3.3 Sensing in physiological samples
With today’s nanosensors, researchers claim that they are able to detect proteins
and DNAs down to femtomolar or even attomolar range with good selectivity.
7,28,37,38,43
However, these detections are performed in purified buffers with very low ionic
strengths. When it comes to clinical diagnosis, the sensitivity and selectivity of a
nanosensor will be significantly suppressed due to the complexity of the sample
composition. Efforts have been made to address this problem by sample purifications
and novel surface modification approaches.
Mark Reed and coworkers have reported biomarker detection from whole blood
samples purified by a microfluidic purification chip (MPC).
64
The biomarkers spiked in a
whole blood sample were captured by the antibody-modified MPC and the
antibody/antigen complexes were released into 0.01X PBS buffer. The complex solution
was then delivered to SiNW-based sensors functionalized with a secondary antibody to
perform sensing. This study marks the first use of label-free nanosensors with
physiological solutions.
In order to overcome the complication caused by physiological samples, our
group developed a faster approach without requiring extra process for device
fabrication.
33
By passivating the In
2
O
3
NW surface with tween-20, we successfully
blocked the signal induced by nonspecific binding when performing active measurement
in whole blood. The detection limit of tween-20 passivated sensors for biomarkers in
whole blood was enhanced to the level similar to the detection limit for the same analyte
in purified buffer solutions at the same ionic strength, suggesting minimal decrease in
device performance in the complex media.
20
Other factors including fabrication cost and device uniformity also limit the wide
application of nanobiosensors in real clinical diagnosis and POCT. In our research team,
we try to tackle some of the problems listed above by improving nanosensor fabrication
techniques, developing novel surface functionalization methods and using engineered
capturing probes. The next four chapters summarize the research projects that I have been
working on during my doctoral studies.
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24
CHAPTER TWO: Selective, electrochemically activated biofunctionalization of
In
2
O
3
nanowires using an air-stable surface modifier
2.1 Introduction
There is growing interest in detecting biological materials in a multiplexed
fashion, which are demonstrated to be essential to the diagnosis and treatment of a range
of diseases and the identification of infectious agents.
1-4
Among various techniques for
multiplexed sensing, nanoscale devices utilizing NWs as active channels are of particular
interest due to the high sensitivity and biocompatibility.
5
In the past few years, these
devices have been employed in the detection of not only individual biomolecules but also
a combination of multiple targets.
6-8
To construct sensor arrays for multiplexed
biosensing, the NWs must be selectively functionalized with different capturing probes
against their designated analytes.
Efforts have been made to achieve the selective functionalization of NW-based
devices, by using microfluidic chips, microspotting techniques
7
and electroactive
monolayers.
9,10
Among these approaches, the use of electroactive monolayers possesses
a number of unique advantages including low cost and simplicity of instrumentation. In
this method, the key step is to design a bifunctional molecule, which bears a NW-
anchoring group on one end and an electroactive moiety on the other. A monolayer of
this molecule is assembled on NWs to create a controllable interface between devices and
capture probes. After being covalently linked to the NWs, the molecule can be activated
from the chemically inert “OFF” state to its “ON” state by applying an external voltage to
device electrodes. In the “ON” state the electroactive moiety reacts with the desired
capture probe to covalently anchor it to the NW surface. This method holds great
25
potential in fabricating high-density sensor arrays for multiplexed biosensing because it is
only limited by the ability to electronically address the individual sensors.
10
By
controlling the potential applied to each sensor, selected devices within an array can be
functionalized with a desired biomolecule, without contaminating the other devices
within the array.
Derivatives of 1,4-hydroquinone (HQ) have been employed in electrochemically
activated surface functionalization of gold, by Mrksich and co-workers.
11
Since that
initial report, this family of molecules has been used in selective functionalization of
various surfaces including In
2
O
3
NWs and Si NWs.
9-13
The oxidized form, BQ (the
“ON” state) readily reacts with a variety of functional groups such as thiols, primary
amines and azides, while HQ (the “OFF” state) is inactive towards these functional
groups. Thus, one can choose a particular device in an array for surface coating by
electrochemically activating it for probe binding, while leaving the other devices in the
array inert. The conversion from HQ derivatives to BQ takes place at relatively low
potentials (ca. 200 mV vs. Ag/AgCl) and can be done in physiological solutions such as
phosphate buffered saline (PBS). Another advantage of HQ derivatives is their
compatibility with biomolecules. However, a major drawback of these molecules is that
the HQ moiety can be gradually oxidized to BQ under aerobic conditions (Figure 2.1).
14
As a result, during the functionalization of a large number of devices, HQ will be
unintentionally converted to BQ (the “ON” state) over time without applying any external
voltage, eliminating the selectivity of this method. Therefore, in order to fabricate high-
density sensor arrays for multiplexed biosensing, it is desirable to design an electrically
26
activated bioconjugate group that possess the advantages of HQ derivatives, and are
stable under aerobic conditions.
Here we report that para-dimethoxybenzene derivatives can serve as reliable, air-
stable, electroactive surface modifiers in selective functionalization of nano-structured
surfaces. In
2
O
3
NWs were chosen in this work due to its simple and well studied surface
derivatization methods. Also, In
2
O
3
NWs are recognized as good candidates in
biosensing,
15,16
because the surface of In
2
O
3
NWs does not possess an insulating, native
oxide layer (e.g., SiO
2
on Si nanowires) that may decrease the nanowire sensitivity.
17
A
para-dimethoxybenzene derivative, 4-(2,5-dimethoxyphenyl)butyl-phosphonic acid
(DMP-PA), was synthesized and preliminarily studies were carried out on indium-tin
oxide (ITO)-coated glass in order to understand the electrochemical behavior of this
molecule. Next, a monolayer of this molecule was formed on an In
2
O
3
NW mat and a
thiol-terminated DNA oligonucleotide was selectively coupled to the region where the
para-dimethoxybenzene moiety was oxidized. We also compared DMP-PA to its HQ
analog, the conventional “OFF” state used in previous reports. The results show that the
OH HO O O
O O O O
+V
O
2
+V
H
3
C
CH
3
O
2
HQ
para-dimethoxybenzene
Figure 2.1 Oxidation process for HQ and para-dimethoxybenzene derivatives.
27
former can largely enhance the selectivity during the functionalization of both ITO and
In
2
O
3
NWs.
2.2 Results and Discussion
2.2.1 Electrochemical behavior of DMP-PA
In aqueous solutions, the para-dimethoxybenzene moiety can be
electrochemically oxidized to produces BQ
18
(Figure 2.1), which serves as the “ON” state,
reacting to form covalent linkages with molecules containing a range of functional groups.
In order to incorporate this group onto In
2
O
3
NWs, we synthesized DMP-PA, a para-
dimethoxybenzene derivative with phosphonic acid terminus, which covalently binds to
metal oxides and metal oxide based NWs.
6
The phosphonic acid group is also known to
bind to the common transparent conducting ITO, which has a surface composition very
similar to that of In
2
O
3
NWs.
19
Therefore, preliminary studies were carried out on ITO-
coated glass slides in order to understand the electrochemical behavior of the molecule.
Freshly-cleaned ITOs were submerged into a 1mM ethanol solution of DMP-PA for 16
hours to form a self-assembled monolayer of DMP-PA. They were then baked at 120°C
for 12 hours in a nitrogen atmosphere to promote the binding of the monolayer to the
substrate. The resultant monolayer binds to ITO surface predominantly via
bidentate/tridentate binding that involves P-O-In bonds.
19
In our cyclic voltammetry (CV)
studies of the DMP-PA derivatized ITO samples (Figure 2.2a), we observed that the
DMP-PA molecule was irreversibly oxidized to BQ-PA at 950 mV, and the BQ-PA was
then reversibly converted to HQ-PA in the following scans, with the oxidation and
reduction potentials centered at +220 mV and -80 mV, respectively. The corresponding
28
reaction pathway of DMP-PA is shown in Figure 2.2b. The mechanism of reaction i in
Figure 2.2b is still under investigation. Judging from the CV traces, the peak area at
+950 mV is much larger than the one for HQ/BQ conversion, which is known to be a
two-electron process. This suggests that the oxidation of para-dimethoxybenzene group
doesn’t follow the two-electron mechanism proposed for the oxidation of HQ diesters, a
group of molecules with similar structures.
20
The surface coverage of BQ-PA/HQ-PA
monolayer on ITO was determined by chronocoulometry (Figure 2.2a, inset). The DMP-
PA coating on an ITO with a fixed area (0.64 cm
2
) was first converted to HQ-PA by
applying an oxidizing potential of 950 mV and subsequent reduction at -80 mV. The
HQ-PA monolayer was then held at -300 mV for 5 seconds and raised to 500 mV and
held for 10 seconds to ensure the complete oxidation to BQ-PA. A charge of 30µC was
consumed for the oxidation process and the calculated molecular coverage was
69Å
2
/molecule or 2.4×10
-10
mol/cm
2
, which is approximately half of the coverage of a
monolayer directly assembled from HQ-PA molecules.
9
The relative low surface
coverage of BQ-PA/HQ-PA can be attributed to the low yield of anodic oxidation of
para-dimethoxybenzene group.
18
29
2.2.2 Stability test of DMP-PA in air
In selective surface functionalization, the “OFF” state should be completely inert
toward the functional groups present in biomolecules, which will be used to attach the
target biomolecule to the surface. Before we were able to perform selective
functionalization using DMP-PA, we first compared it to HQ-PA, the previously used
“OFF” state, and studied their stabilities when they are exposed to thiols in air. Thiol is a
common group present in antibodies and is readily incorporated into oligonucleotides.
Nucleophiles such as thiols and amines react with BQ derivatives by Michael addition,
and the addition typically causes a negative shift of the redox potential due to the
-0.5 0.0 0.5 1.0 1.5
-20
0
20
40
Current ( A)
Potential vs. Ag/AgCl (V)
First scan
Second scan
-5 0 5 10
0
10
20
30
Q ( C)
Time (s)
(a)
(b)
DMP-PA BQ-PA HQ-PA
OMe MeO
P
O
O O
O O
P
O
O O
i
P
O
O O
ii
OH HO
iii
Figure 2.2. Electrochemical characterization of DMP-PA on ITO electrodes. (a) CV
characterization of a DMP-PA derivatized ITO sample. Inset: A chronocoulometry
trace showing the amount of charge necessary to oxidize a pre-defined area of HQ-PA
to BQ-PA. (b) The corresponding reaction pathway of the electrochemical process
shown in (a).
30
electron-donating effect.
21-23
The effect of thiol addition to BQ-PA monolayer is shown
by the CV traces in Figure 2.3a. A DMP-PA derivatized ITO sample was first
electrochemically oxidized, and the resulting BQ-PA monolayer was continuously
scanned by CV in presence of 10 µM 3-mercaptopropanol (in PBS, pH 7.4). During 10
scans, the redox potential of the monolayer gradually shifted in the negative direction,
which corresponds to the addition of 3-mercaptopropanol to the BQ head group. Based
on this result, we can use the negative shift in redox potential as an indicator of thiol
addition. In order to study the stability of DMP-PA when exposed to thiols
electrochemically, a DMP-PA derivatized ITO sample was submerged to 3-
mercaptopropanol solution for 30 minutes and then washed extensively with PBS to
remove any unbound thiols. The sample was then subject to an oxidative potential at 950
mV, followed by CV scans (Figure 2.3b). The CV trace of this sample (red) showed
good alignment with the black trace collected from a pure BQ-PA monolayer on ITO.
This indicates that DMP-PA is stable under aerobic conditions and is unreactive toward
thiols. In parallel, a HQ-PA-ITO sample was also treated with 3-mercaptopropanol in the
same manner and examined by CV (blue trace in Figure 2.3b). It is clear that the redox
potential of this monolayer showed a negative shift compared to the pure BQ-PA
monolayer. As previously mentioned, the HQ group is prone to aerobic oxidation to BQ
under aerobic conditions. We attribute the potential shift to the fact that some HQ-PA
molecules were oxidized to BQ-PA during the thiol-incubation and reacted with the thiol.
From this experiment, we can conclude that DMP-PA has better stability compared to
HQ-PA and can be used as a reliable “OFF” state in selective functionalization methods.
31
2.2.3 Selective functionalization of In
2
O
3
NW mat with DNA oligos
After we demonstrated the good stability of DMP-PA on ITO electrodes, we
performed selective functionalization of In
2
O
3
NW mats with thiolated-DNA. Single-
crystalline In
2
O
3
NWs (average diameter of 20 nm and length of 5-10 µm) were grown
on a Si/SiO
2
substrate with a SiO
2
layer of 500 nm using a laser ablation technique. The
-0.2 -0.1 0.0 0.1 0.2 0.3
-20
-10
0
10
Current (uA)
Potential v.s. Ag/AgCl (V)
BQ-PA
D MP-PA expo se d to thiol
HQ-PA exposed to thiol
(a)
-0.4 -0.2 0.0 0.2 0.4 0.6 0.8
-20
-15
-10
-5
0
5
Current ( A)
Potential vs. Ag/AgCl (V)
-0.2 -0.1
-15
-10
-5
Figure 2.3. Stability test for DMP-PA. (a) The first 10 CV scans of a BQ-PA
monolayer in presence of 10 µM 3-mercaptopropanol (in PBS, pH 7.4). The peak
current decreases during the scans. Inset: The region of the reduction peak. The arrow
indicates the shift of the peak current during the 10 scans. (b) CV characterization of
ITO samples with BQ-PA monolayer (black), DMP-PA monolayer after exposed to
thiols (red) and HQ-PA monolayer after exposed to thiols (blue). The DMP-PA
monolayer was first subject to an oxidative potential at 950 mV before CV
measurement.
32
detailed synthesis method and the characterization of these NWs can be found
elsewhere.
24,25
Metal electrodes (Ti/Au/Ni, 5 nm/40 nm/5 nm) were patterned on as-
grown NW mats by photolithography and metal deposition. Figure 2.4a shows an optical
image of four groups of electrodes deposited on In
2
O
3
NW mats. Scanning electron
microscopy (SEM) images of NW mat and electrodes are shown in Figure 2.4b and 2.4c.
The mat sample was submerged in 1mM solution of DMP-PA overnight and baked at
120 °C for 12 hours in a nitrogen atmosphere. The chip was then soaked in an ethanol
solution of dodecane-1-thiol overnight to passivate the metal electrodes. An oxidizing
potential of 950 mV was applied to the first group of electrodes, 1, converting the NW
surface coating of DMP-PA to BQ-PA. The second group, 2, was subjected to an
oxidizing potential of 950 mV, followed by a reducing potential of -80 mV, converting
the DMP-PA coating to a HQ-PA coating. The third group, 3, has the original DMP-PA
coating. The entire chip was then soaked in a solution of a thiol-terminated probe DNA
for 2 hours. The BQ-PA coating on 1 is expected to bind the thiol-DNA efficiently,
while 2 and 3 are not expected to bind the DNA. After extensive rinsing with PBS buffer,
the chip was treated with a solution of complementary DNA labeled with a fluorescence
dye, and again washed with PBS buffer. The three groups of electrodes were examined
by fluorescence microscopy and are shown in Figure 2.4 (d), (e) and (f), showing 1, 2 and
3, respectively. In these figures the gold electrodes appear as dark stripes and any bound
fluorescently labeled DNA appears as a bright network. As expected, the NWs in 1 show
intense fluorescence, indicating that the BQ-PA surface did bind the thiol-DNA and then
the labeled compliment. In contrast, 3, which went through the same DNA treatment, did
not show any fluorescence, as seen in Figure 2.4(f), and thus is inert under these
33
conditions. The NWs in 2, however, show some fluorescence after the DNA treatment,
albeit at a brightness lower than that seen for 1, but nonetheless there was binding of the
thiol-DNA to the HQ-PA surface. This is in agreement with the results shown in Fig
2.3(b), indicating that HQ-PA reacts with thiols due to the aerobic oxidation of HQ
moiety.
Figure 2.4. Selective functionalization of In
2
o
3
NW mat with DNA oligos. (a) An
optical image of In
2
O
3
NW mats with four groups of electrode deposited on top. (b) A
typical SEM image taken in the center region of one group of electrodes. (c) The same
image at higher magnification. Figure (d)-(f) are fluorescence images of In
2
O
3
NWs
near three groups of electrodes. NWs were first functionalized with DMP-PA. An
oxidizing potential of 950 mV was applied to the first group (d), converting the DMP-
PA coating to BQ-PA. The second group (e) was subjected to an oxidizing potential of
950 mV, followed by a reducing potential of -80 mV, converting the DMP-PA coating
to a HQ-PA. The third group (f) has the original DMP-PA coating. The three groups
were treated with thiolated single strand DNA, and then its complementary DNA
labeled with a fluorescent dye. After extensive rinsing with PBS buffer, the NWs near
the three groups of electrodes were examined under fluorescent microscope.
34
2.3 Chapter Conclusions
In this work we demonstrate that a para-dimethoxybenzene derivative, DMP-PA,
can serve as an air-stable, reliable, electroactive surface modifier in selective
functionalization of In
2
O
3
NWs. By tailoring the molecular structure, the para-
dimethoxybenzene group shows improved stability under aerobic conditions and
significantly enhanced selectivity in surface functionalization compared to the
conventional HQ moiety. The approach described herein allows for the functionalization
of a single In
2
O
3
NW device within an array with a desired capture probe, without
contamination of the other devices/sensors in the array. While this work is focused on
In
2
O
3
NW based devices, the same approach can be used for selective, electrochemically
driven functionalization of devices within an array for nearly any semiconductor NW
based device. This can be considered a key step for the future fabrication of high-density
biosensor arrays for multiplexed biosensing.
2.4 Experimental section
Materials and instruments. 4-(2,5-dimethoxyphenyl)butyl-phosphonic acid
(DMP-PA) was synthesized according to a reported procedure.
9
Phosphate Buffered
Saline (PBS), dodecanethiol and 3-mercaptopropanol was purchased from Sigma-Aldrich.
Tris (2-carboxyethyl) phosphine hydrochloride (TCEP) was purchased from Strem
Chemicals. Both the probe and the target DNA oligonucleotide were purchased from
Integrated DNA Technologies, Inc. The electrochemistry was performed with
Potentiostat/Galvanostat Model 263A (Princeton Applied Research). The fluorescence
images were captured using a Nikon Eclipse LV-100D-U fluorescence microscope.
35
In
2
O
3
nanowire growth and electrode deposition. In
2
O
3
nanowires were
synthesized via laser ablation on Si/SiO
2
substrates.
18
Metal electrodes were fabricated
on as-grown nanowire samples by photolithography, followed by Ti (5 nm)/Au (40
nm)/Ni (5 nm) deposition. After lift-off, the chip was carefully cleaned before further
modification.
Formation of monolayer coatings on ITO-coated glass slides and In
2
O
3
nanowire mats. ITO slides were boiled for 5 minutes each in trichloroethylene, acetone,
and finally, ethanol. The slides were then placed in an ozone/UV chamber for 10 minutes.
The NW mats were cleaned by the same procedure as ITO, but placed in the ozone/UV
chamber for 2 minutes. The pre-cleaned ITO slides or NW mats were soaked into a
solution of DMP-PA in ethanol (1mM) for 16 hours, and then washed extensively with
ethanol and dried under nitrogen. The ITO slides or NW mats were then annealed at
120°C in nitrogen atmosphere for 12 hours. For NW mats, a further step was taken to
passivate the gold electrodes. The sample was soaked in a 1mM solution of
dodecanethiol in hexane overnight.
Electrochemistry. Electrochemistry of the DMP-PA monolayer on both ITO and
In
2
O
3
NWs were performed in an electrochemical cell filled with PBS buffer (pH 7.4),
with Pt wire as the counter electrode and Ag/AgCl as the reference electrode. All
electrochemical procedures were carried out in the air. The determination of the BQ-PA
monolayer coverage was carried out by chronocoulometry.
Preliminary experiments on ITO slides. For the thiol addition experiment, the
DMP-PA monolayer on ITO was first converted to BQ-PA. Then the sample was
continuously scanned by CV in a solution of 3-mercaptopropanol (60 µM) in PBS. For
36
the stability test experiment, a DMP-PA-ITO sample was incubated in 3-
mercaptopropanol solution for 30 minutes and then washed extensively by PBS buffer.
The sample was then oxidized to BQ-PA before CV measurements. Another DMP-PA-
ITO sample was first converted to HQ-PA and then treated with 3-mercaptopropanol
solution in the same manner.
Selective functionalization of In
2
O
3
NWs. The probe DNA oligonucleotide used
for NW mats functionalization was a 20-base DNA with the sequence of [5’-XGCT TTG
AGG TGC GTG TTT GT-3’], where X was a 5’ thiol modifier C6. To reduce the thiol
modifier, the probe DNA (7 nmol) was first incubated with TCEP (10.5 nmol) in PBS
buffer at room temperature for 2 hours. The excess TCEP was then removed by passing
the mixture through a desalting column. The electrochemically treated NW mats were
immediately immersed into a solution of the probe DNA (~15 μM in PBS) and stored in a
humidity chamber for 2 hours. The devices were then carefully rinsed with PBS to
remove any unbound DNA. For the DNA hybridization, the NW mats were exposed to a
solution of the target DNA (5’-RhoR-XN/ACA AAC ACG CAC CTC AAA GC-3’) for
20 minutes, followed by extensive washing with PBS. The NW mats were then
examined under a fluorescence microscope and the images were captured using NIS-
elements software.
2.5 Chapter 2 References
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82, 136-144.
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R. J.; Thompson, M. E.; Zhou, C. W. Acs Nano 2009, 3, 3969-3976.
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P. C.; Zhang, R.; Roberts, R. W.; Sun, R.; Cote, R. J.; Thompson, M. E.; Zhou, C. W. Acs
Nano 2009, 3, 1219-1224.
(17) Bunimovich, Y. L.; Shin, Y. S.; Yeo, W. S.; Amori, M.; Kwong, G.; Heath,
J. R. Journal of the American Chemical Society 2006, 128, 16323-16331.
(18) Fichter, F.; Dietrich, W. Helvetica Chimica Acta 1924, 7, 131-142.
(19) Paramonov, P. B.; Paniagua, S. A.; Hotchkiss, P. J.; Jones, S. C.;
Armstrong, N. R.; Marder, S. R.; Bredas, J. L. Chemistry of Materials 2008, 20, 5131-
5133.
(20) Meier, E. P.; Chambers, J. Q.; Chambers, C. A.; Eggins, B. R.; Liao, C. S.
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Electroanalytical Chemistry 1992, 326, 197-212.
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47, 4351-4356.
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39
CHAPTER THREE: Field-effect transistors based on ZnO nanomaterials
synthesized from a hydrothermal method
3.1 Introduction
Zinc oxide (ZnO) is a wide bandgap (3.37 eV) semiconductor having a high
electron-hole binding energy (60 meV) and important applications in electronics, optics,
optoelectronics, laser and light-emitting diode.
1
It is also a green material that is
bio-compatible, biodegradable and bio-safe for medical applications and environmental
science.
2,3
The study of 1D ZnO nanomaterials was first inspired by the discovery of
ZnO nanobelts by Z. L. Wang’s research group in 2001.
4
In the past decade, ZnO 1D
nanostructures have demonstrated promising potential as building blocks for nanoscale
electronics. As an example, FETs using ZnO nanowires or nanorods as the active
channel have been intensively studied, and their sensing properties to gas,
5,6
chemical
7
and biological species
8
have been investigated.
There have been many existing preparative techniques for this material. Among
them, a vapor-phase transport process with the assistance of noble metal catalysts or
thermal evaporation are the two major vapor methods to fabricate one-dimensional ZnO
nanostructures with controllable diameters.
4,9-16
However, these methods normally
require high growth temperature and complicated instrumentation while giving low
product yields. In recent years, people have turned their interest to solution-phase
synthesis techniques including hydrothermal method and electrochemical deposition.
17,18
These approaches are attractive because of the low growth temperature and potential of
producing high density of nanomaterials. A major drawback of solution-phase
synthesized nanomaterials is that they usually have poor electrical properties compared to
40
the conventional vapor-phase synthesized ones due to higher amount of defects and
impurities introduced during the synthesis process. FETs fabricated with these ZnO
nanostructures typically show much poorer electrical performance compared to those
fabricated with vapor-phase synthesized materials, characterized by their low
conductance and on/off ratio.
19-22
This chapter describes a simple annealing method that effectively improves the
electrical properties of ZnO nanomaterials (nanowires and nanobelts) prepared by a
hydrothermal method. The electrical properties of ZnO nanomaterials were measured in
the structure of FETs. By applying a post-synthesis annealing step to these ZnO products,
the performance of FETs was significantly enhanced in terms of both conductance and
on/off ratio. The so-obtained FETs shows comparable performance with those based on
ZnO nanostructures synthesized via vapor-phase approaches.
3.2 Results and Discussion
3.2.1 Synthesis and characterization of ZnO nanomaterials
Hydrothermal growth of ZnO 1D nanomaterials is normally performed by
hydrolyzing Zn
2+
salts in a basic solution, with or without the aid of seeds or templates.
23
In this project we started with a seeded growth method which was first reported by P. D.
Yang in 2003.
24
This method involves thermal decomposition of Zn(NO
3
)
2
at ~ 90 C in
an aqueous solution containing equal molar of hexamethylenetetramine. A substrate is
pre-seeded with ZnO nanoparticles and suspended in the solution upside down during the
growth. A surfactant, polyethylenimine, is usually added to the growth solution in order
to increase the length-to-diameter ratio of the ZnO products. Figure 3.1a and 3.1b shows
41
the scanning electron microscopy (SEM) images of a typical sample after 7.5 hours
growth. The diameter of the as-grown ZnO nanorods is in the range of 40~80 nm and
their length is about 2.5 μm. The X-ray diffraction (XRD) pattern shows only the 002
peak in Figure 3.1e, indicating that the ZnO nanorods grow off the substrate along the
c-axis direction. In order to obtain longer ZnO nanorods for FET fabrication, we
increased growth time from 7.5 hours to 14.5 hours. However, SEM images (Figure 3.1c
and 3.1d) show that the average length of these ZnO nanorods only increased to 3 μm
while the diameter almost doubled.
Figure 3.1. Characterization of ZnO nanorods grown on Si substrate. (a) and (b) are
SEM images of the top-view and side-view of nanorods from 7.5 hours growth,
respectively. (c) and (d) are the top-view and side-view of nanorods from 14.5 hours
growth, respectively. (e) is the XRD pattern of ZnO nanorods from 7.5 hours growth.
42
We then switched to a substrate-free hydrothermal method in which the growth is
conducted by hydrolyzing ZnCl
2
in an aqueous solution of Na
2
CO
3
at 140 C.
25
The
reaction is performed in an autoclave for 12 hours and the products are then filtered from
the solution and dried. Approximately 25 mg ZnO products can be obtained from 80 mg
ZnCl
2
in each synthesis, while the chemical vapor deposition (CVD) method yields only
~0.5 mg each time on a 1cm × 1cm substrate (shown in Figure 3.2a).
26
The ZnO
products were examined by SEM and a low-magnification image is shown in Figure 3.2b.
At higher magnification (Figure 3.2c), we can clearly see that the as-prepared ZnO
products are composed of wire and belt-like nanostructures, which is consistent with the
reported results. The diameters of ZnO nanowires and nanobelts are 60~90 nm and
100~250 nm, respectively, and the average length is 5.5 µm with a standard deviation of
1.5 µm. A typical XRD pattern is shown in Figure 3.2d, in which all the diffraction
peaks can be well indexed to the wurtzite structure of ZnO. The high intensity and
narrow width of the peaks indicates that the as-prepared ZnO products are of high
crystalline quality.
43
Further structural analyses of the ZnO products were performed using
transmission electron microscopy (TEM). The TEM image presented in Figure 3.3a
reveals the shape of a nanobelt and a nanowire with diameters of 60 nm and 120 nm,
respectively. In the nanobelt, we can clearly see the ripple-like contrast that is due to the
strain resulting from the bending of the nanobelts.
27
On the other hand, the nanowire
shows much less contrast since it is more resistant to the bending. The high-resolution
TEM (HRTEM) image in Figure 3.3b shows clear lattice fringes, which indicate no
defect or dislocation in the ZnO products. The average distance of 0.26 nm between the
adjacent lattice planes corresponds to the (0002) plane lattice spacing of wurtzite
structured ZnO. The inset in the top-left-hand corner of Figure 3.3b is a typical selected
10 µm
30 35 40 45 50 55 60
0.0
5.0x10
2
1.0x10
3
1.5x10
3
2.0x10
3
2.5x10
3
Intensity
2 theta (degree)
100
101
002
102
110
1 µm
belt
wire
(a)
(c)
(d)
(b) (a)
Figure 3.2. Characterization of hydrothermal ZnO products. (a) A picture showing the
amount of ZnO products made from two different methods. On the left side is a vial of
25 mg ZnO products obtained from one run of hydrothermal synthesis. On the right
side is a film of ZnO nanowires grown on a 1cm × 1cm quartz substrate by CVD
method (~0.5 mg). (b) and (c) is the low-magnification and high-magnification SEM
images of the ZnO products, respectively. (d) XRD pattern of the as-prepared ZnO
products.
44
area electron diffraction pattern (SAED) of the ZnO product, which indicates the single
crystalline character and the growth direction along the c-axis.
3.2.2 FET fabrication and electrical performance
To fabricate FETs, the as-grown ZnO products were first suspended in isopropyl
alcohol (IPA) by ultra-sonication and then dispersed onto a 3-inch Si/SiO
2
wafer with a
SiO
2
layer of 50 nm. This step was repeated multiple times until a desired surface
density (2~3 nanowires/100 μm
2
) was achieved, which was verified by optical microscopy
and SEM. Electrodes were patterned by photolithography followed by e-beam deposition
of 5 nm Ti and 45 nm Au. FETs were thus obtained, with metal contacts functioning as
source and drain electrodes and the Si substrate as a back gate (Figure 3.4a). A SEM
image (Figure 3.4a, inset) shows a single ZnO nanowire confined between the
source/drain electrodes (channel length 2 µm).
The electrical properties of these ZnO nanowires and nanobelts were studied in
the structure of FETs. The drain current versus gate voltage (I
ds
vs. V
gs
) characteristics of
a typical FET at V
ds
(source-drain voltage) = 0.1 V is shown in Figure 3.4b. The
transistor shows low conductance and nearly no gate dependence. This poor electrical
(a)
(b)
Figure 3.3. Characterization of hydrothermal ZnO products using TEM. (a) TEM
image of a ZnO nanowire and a ZnO nanobelt. (b) HRTEM image taken from an
individual ZnO nanowire. The insert is the corresponding SAED pattern.
45
performance is probably due to the fact that solution-grown ZnO nanomaterials normally
contain more impurities compared to vapor-phase synthesized ones and also have a
relatively low doping level. It is well known that ZnO is intrinsically an n-type
semiconductor and the doping concentration is largely determined by the density of
oxygen vacancies.
28
The number of oxygen vacancies in the ZnO lattice is relatively low
when the growth takes place in solutions instead of an oxygen-deficient vapor
environment. Based on our experience with other metal oxide nanowires,
29
a
post-synthesis annealing step in vacuum can modify the condition of oxygen vacancies in
nanowires and results in improved device performance.
We applied a post-synthesis annealing step to the ZnO products in order to
enhance the device performance. The ZnO products were annealed under vacuum at
200
o
C for 80 minutes after being dispersed on Si/SiO
2
wafers, but before the metal
contacts were added. Figure 3.5a illustrates the I
ds
vs. V
gs
characteristics of a typical FET
made with annealed ZnO products. At V
ds
= 0.1V, the device exhibits a threshold voltage
of -5 V and an on-current higher than 10
-7
A. The semilog-scale plot clearly shows that
-20 -10 0 10 20
20
25
30
35
Ids (nA)
Vgs (V)
(a)
1 μm
(b)
Figure 3.4. FETs based on as-grown ZnO products. (a) Schematic view of a ZnO FET
structure, with Ti/Au as source and drain electrodes and Si as a back gate. Inset: a SEM
image of a ZnO FET. (b) I
ds
-V
gs
characteristics of a FET made with as-grown ZnO
products at V
ds
= 0.1V.
46
the on/off ratio of this device is on the order of 10
5
. For the same device, I
ds
vs. V
ds
characteristics were measured under different V
gs
varying from -10 V to 10 V, and the
obtained curves are shown in Figure 3.5b. The device exhibits typical
enhancement-mode n-type semiconductor transistor behavior with an on-current of 2 µA,
while V
gs
= 10 V and V
ds
= 2 V. The calculated transconductance g
m
( ) at V
ds
=
0.1V, along with a Gaussian fit to the data, is plotted versus V
gs
in Figure 3.5c. The g
m
peaks at 1.3× 10
-8
S, at V
gs
= 7.5V, and falls off with increasing gate voltage. The field
effect mobility in a typical cylindrical nanowire system with radius r can be expressed as
follows
30
(1)
where is the number of nanowires confined between two electrodes, L the
channel length, the thickness of the dielectric layer, and the dielectric constant of the
SiO
2
. Although a nanobelt does not have a cylindrical geometry, it is reasonable to
estimate the capacitance by replacing radius with width as a first order
approximation.
31
The calculated mobility for the device mentioned above is 64
cm
2
V
-1
s
-1
. This value is much higher than previously reported data of FETs made with
hydrothermal grown ZnO nanomaterials.
47
3.2.3 Statistical study of the effect of annealing on FET performance
In order to study the effect of the annealing step statistically, we fabricated two
groups of FETs in parallel. Group 1 contains 17 devices made with annealed ZnO
products while group 2 contains 13 ones made with unannealed ZnO. The two groups of
FETs were characterized and the statistics of on-current and on/off ratio is shown in
Figure 3.6a and 3.6b, respectively. In Figure 3.6a, the devices from group 2 show a wide
distribution of on-current over the range of 10
-12
A to 10
-7
A at V
ds
= 0.1V. For group 1,
upon annealing, the ZnO nanowires and nanobelts produced devices with higher
on-current in a narrower range of 10
-8
A to 10
-6
A. Note that the average on-current of
-2 -1 012
-4
-2
0
2
Ids ( A)
Vds (V)
Vg = -10V
Vg = -5V
Vg = 0V
Vg = 5V
Vg = 10V
-20 -10 0 10 20
0
50
100
150
200
250
Ids (nA)
Vgs (V)
10
-12
10
-11
10
-10
10
-9
10
-8
10
-7
10
-6
Log(Ids)
1 µm
(c)
-20 -10 0 10 20
0.00
0.01
0.02
0.03
g
m
GaussFit of g
m
g
m
( S)
Vgs (V)
(b) (a)
Figure 3.5. Measured characteristics of a typical FET made with annealed ZnO
products. (a) I
ds
-V
gs
characteristics at V
ds
= 0.1V. The black and red curve correspond
to the linear and semilog-scale plot of the I
ds
-V
gs
. (b) I
ds
-V
ds
family curves when V
gs
varies from -10 V to 10 V with a step of 5 V. (c) Measured g
m
at V
ds
= 0.1 V, along
with the Gaussian fit to the data.
48
10
-7
A exceeds that of the previously reported FETs based on 1D ZnO nanostructures
from hydrothermal synthesis.
32-34
The annealing step also introduced a significant
improvement in the on/off ratios (Figure 3.6b). Most of the devices in group 2 exhibit
on/off ratio lower than 10, while in the case of group 1, although a wide distribution is
observed, roughly 70% of the devices show on/off ratio of 10
2
~ 10
5
, which meet the
requirements of most sensing applications. About 30% of the devices show excellent
on/off ratio of 10
4
~10
5
, which makes them competitive with FETs based on 1D ZnO
nanostructures (nanowires and nanobelts) synthesized by vapor-phase approaches. The
enhancement of device performance upon vacuum annealing is mainly attributed to the
change of the doping level in ZnO nanowires.
3.2.4 Device uniformity
Upon annealing the ZnO products prior to device fabrication, we were able to
fabricate FETs with significantly enhanced performance. However, as discussed above,
the devices show a relatively wide distribution in terms of the performance. Attempts
10
-12
10
-11
10
-10
10
-9
10
-8
10
-7
10
-6
0
20
40
60
80
100
Percentage
on-current (A)
Unannealed
Annealed
10
0
10
1
10
2
10
3
10
4
10
5
0
20
40
60
80
100
Percentage
on/off ratio
Unannealed
Annealed
(a) (b)
Figure 3.6. Statistics of 17 annealed devices and 13 unannealed ones. (a) and (b) shows
the distribution of on-current and on/off ratio of these devices, respectively.
49
were made to suppress the device-to-device variation by reducing the number of ZnO
nanobelts in the products, since the presence of large numbers of nanobelts results in
wide distribution of the nanowire diameter, which in turn affects uniformity of device
performance. It was reported that the growth of nanobelts is assisted by the surfactant
used during the synthesis, sodium dodecyl sulfate (SDS). In the absence of SDS, the yield
of nanobelts can be largely reduced.
25
Based on this observation, we synthesized a new
batch of ZnO products without using SDS. These products contain mostly nanowires and
have a narrower distribution in terms of diameter, as illustrated by the SEM images in
Figure 3.7. After synthesis, these ZnO nanowires went through the same annealing step
followed by the device fabrication to give an array of FETs, and the electrical properties
of these devices were measured as before. However, no significant improvement was
observed in terms of device uniformity. The reason behind the non-uniformity of device
performance is still under investigation, and an effective method to increase the device
yield remains to be discovered.
Figure 3.7. ZnO products synthesized without SDS. This batch has fewer belts.
50
3.2.5 Surface functionalization of ZnO products
It has been reported that ZnO nanomaterial can be functionalized by carboxyl
acids, phosphonic acids and silanes.
35-38
In order to compare the binding affinity of these
molecules, we functionalized ZnO products with three molecules terminated with a
hydrophobic tail (A, B and C shown in Figure 3.8) and measured the contact angle of thin
films prepared with these ZnO products. ZnO nanomaterials were dispersed in 10 mM
solutions of molecule A, B and C in ethanol and stirred overnight. The products were
then filtered and washed by fresh ethanol for several times. ZnO was then re-suspended
in ethanol and drop-coated on Si substrates to form thin films. As shown in Table 3.1,
the contact angle of film B has the highest value, indicating that phosphonic acids have
the best binding affinity to ZnO.
Molecule A B C
Contact Angle (degree) ~20 ~130 ~90
Table 3.1. Contact angle of ZnO nanomaterial films functionalized with molecule A, B
and C.
Figure 3.8. Structure of molecule A, B and C.
51
The successful functionalization of ZnO with molecule B was further confirmed
by ATR-IR. The IR spectrum of free molecule B and functionalized ZnO were recorded
and shown in Figure 3.9a. For free phosphonic acid, the absorption peak between ca.
955 cm
-1
and 930 cm
-1
are associated with P-O-(H) stretching vibrations and the peak
centered at 1005 cm
-1
corresponds to the asymmetric ν
a
P-O-(H) vibration. The surface
bound molecule B shows two broad peaks at 962-1000 cm
-1
and 1000-1070 cm
-1
, raising
from ν
s
PO
2
–
and ν
s
PO
3
–
vibration, respectively. The relative intensity of these two group
of peaks suggest mainly bidentate binding of the phosphonic acid.
39
In order to anchor biological molecules onto ZnO nanomaterials, we
functionalized them with 3-phosphonopropionic acid and 3-aminopropyl phosphonic acid,
Figure 3.9. IR spectrum of ZnO nanomaterials functionalized with (a) molecule B, (b)
3-phosphonopropionic acid and (c) 3-aminopropyl phosphonic acid.
52
two phosphonic acid derivatives terminated with -COOH and -NH
2
, respectively. The
functionalization solution was prepared with ethanol instead of water because we
observed notable etching of ZnO after overnight incubation in aqueous solutions. After
functionalization, ZnO products were examined by ATR-IR and we were able to see the
peaks arisen from surface bound phosphonic acids (Figure 3.9b and 3.9c). These -COOH
or -NH
2
terminated ZnO nanomaterials can then be used for biorecognition molecule
attachment in future sensing experiments.
3.3 Chapter Conclusions
In summary, “mass” production of ZnO nanowires and nanobelts were prepared
using a low cost, low temperature hydrothermal method, and used to fabricate back-gated
FETs. By applying a post-synthesis annealing step to the ZnO products, we were able to
adjust the doping level of the nanowires and nanobelts, and thus significantly improve
their electrical properties. Roughly 70% of the FETs made with annealed ZnO products
exhibit promising performance, with on/off ratio of 10
2
~10
5
, on-current higher than
10
-7
A and field-effect mobility of ~ 64 cm
2
V
-1
s
1
. About 30% of the devices show
excellent on/off ratio of 10
4
~10
5
, and they are competitive with FETs based on 1D ZnO
nanostructures synthesized by vapor-phase approaches. These characteristics exceed
those of the previously reported FETs prepared with ZnO nanowires and nanorods
obtained from hydrothermal methods.
53
3.4 Experimental Section
Chemicals. ZnCl
2
and SDS were purchased from Sigma-Aldrich. Na
2
CO
3
was
purchased from Mallinckrodt. Molecule A, B and C were purchased from UCT, Alfa
Aesar and Sigma-Aldrich, repectively. 3-phosphonopropionic acid and 3-aminopropyl
phosphonic acid were purchased from Sigma-Aldrich. All chemicals were used without
further purification.
ZnO nanomaterial synthesis. 0.08 g ZnCl
2
, 0.6 g SDS and 8 g Na
2
CO
3
were
dissolved in 20 ml HPLC grade water. The mixture was stirred for 30 mins and then
transferred to a Teflon-lined stainless steel autoclave bottle. The autoclave was kept in an
oven at 140
C for 12 hrs. The precipitates were filtered and washed by hot water and
ethanol. The white product was dried in an oven at ~80 C for 4 hrs and kept in a
vacuum desiccator for further use.
Characterization of ZnO products. SEM images were obtained on JEOL-7001
SEM operating at 10.0 kV. Powder XRD analyses were performed on a Rigaku Ultima
IV X-ray diffractometer using a Cu K α radiation source ( λ = 1.54 Å). TEM and SAED
analyses were carried out on a JEOL JEM-2100 microscope at an operating voltage of
200 kV equipped with a Gatan CCD camera. To prepare TEM samples, as-grown
nanomaterials were sonicated at mild power in ethanol, and a drop of this solution was
put on the 300 mesh Cu grid.
FET fabrication and electrical testing. The ZnO products were first suspended
in IPA by ultra-sonication and then dispersed onto a 3-inch Si/SiO
2
wafer with a SiO
2
layer of 50 nm. This step was repeated multiple times until a desired surface density
(2~3 nanowires/100 μm
2
) was achieved, which was verified by optical microscopy and
54
SEM. Electrodes were patterned by photolithography followed by e-beam deposition of
5 nm Ti and 45 nm Au. Current-voltage characteristics of FETs were measured using an
Agilent semiconductor parameter analyzer.
Surface modification of ZnO nanomaterials. ZnO powders were suspended in
0.1 mM solution of phosphonic acids in ethanol and stirred overnight. ZnO were filtered,
washed by fresh ethanol for several times and dried in an oven at 80 C.
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W.; Marks, T. J.; Janes, D. B. Nature Nanotechnology 2007, 2, 378-384.
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F.; Saykally, R. J.; Yang, P. D. Angewandte Chemie-International Edition 2003, 42,
3031-3034.
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56
CHAPTER FOUR: Construction of nanobiosensors using cyclotides as capturing
probes
4.1 Introduction
In less than a decade, biosensors based on nanowire/carbon nanotube transistors
have successfully made the transition from proof of concept
2
to highly selective,
ultrasensitive devices capable of detecting specific proteins and DNA sequences.
3-11
These devices utilize a capture agent on the sensor surface to selectively bind the target
biomolecules. The captured biomolecules affect the electronic properties of the
nanowires/nanotubes, resulting in an electronically readable signal. Capturing agents
commonly used in nanobiosensors include antibodies, oligonucleotides, and small
affinity binding agents (e.g., biotin).
2,4,7-11
In contrast to typical antibodies and oligos, small affinity binding agents possess a
few advantages that are important in nanobiosensing applications. First of all, these
molecules are stable to a wide range of pH and electrolyte concentrations, and are
relatively small. Secondly, they are normally synthesized in labs so it is expected that
these binding agents can be produced in large quantity, at relatively low cost. Thirdly,
lots of these binding agents can be evolved/engineered to improve recognition properties
such as selectivity and binding affinity, with the potential to surpass conventional
antibodies and nucleotides. Moreover, it is relatively easy to add various functional
groups at designated locations on these molecules so that they can be immobilized on
sensor surface with certain orientations via desired bioconjugation methods.
Cyclotides are a new emerging family of large plant-derived backbone-cyclized
polypeptides ( ≈ 28-37 amino-acids long) that share a disulfide-stabilized core (three
57
disulfide bonds) characterized by an unusual knotted structure (Figure 4.1).
12,13
In this
motif, an embedded ring formed by two disulfide bonds and their connecting backbone
segments is penetrated by the third disulfide bond. Cyclotides contrast with other circular
poylpeptidesin that they have a well-defined three-dimensional structure, and despite
their small size, can be considered as miniproteins. Their unique circular backbone
topology and knotted arrangement of three disulfide bonds makes them exceptionally
stable to thermal and enzymatic degradation.
14
In this chapter, we use cyclotides as a new class of capture agents for
nanowire/nanotube biosensors. Cyclotide-modified nanobiosensors are constructed for
the detection of trypsin, a protease related to pancreas diseases, using devices based on
In2O3 nanowires. The cyclotide used in this work is MCoTI-II.
Figure 4.1 Primary and tertiary structure of trypsin inhibitor (MCoTI-II). Conserved
cysteine residues are marked in yellow and disulfide connectivities in red. The circular
backbone topology is shown with a blue line.
1
58
4.2 Results and discussion
4.2.1 Construction of nanobiosensors using MCoTI-II
When we use conventional antibodies or oligonuleotides as capturing probes in
nanobiosensing, our choices of surface immobilization methods are limited to amine or
thiol coupling reactions because these are the most common functional groups present in
biomolecules. Each capturing probes normally contains more than one amine or thiol
groups so that when the probes are immobilized to the sensor surface, their orientation is
often random. This may cause the binding region of some capturing probes to be blocked
by sensor surface or nearby molecules, which may decrease the binding affinity towards
analytes. The use of engineered small binding agents as capturing probes can solve these
problems. It is possible to add a variety of functional groups at desired positions of these
molecules so it helps expand our choice of surface chemistry and improve the control
over molecular orientation on sensor surface. In this work, we chose to add an alkyne
group at the N-terminal of MCoTI-II and use the well developed “click chemistry” to
immobilize it on In
2
O
3
NW-based sensors. The alkyne group is located on the opposite
side of the binding region of MCoTI-II toward trypsin so that it allows maximum chances
to capture analytes when MCoTI-II is immobilized on sensor surface. The synthesis and
structure modification of MCoTI-II is done by Professor Julio Camarero’s group. In
order to link the alkyne-terminated MCoTI-II to In
2
O
3
NWs, we designed a bifunctioal
linker (3-azidopropyl phosphonic acid, molecule B) which has a phosphonic acid group
on one end and an azide group on the other. The synthesis route of this molecule is
shown Figure 4.2a and proposed surface chemistry strategy is shown in Figure 4.2b.
59
4.2.2 Click chemistry condition
In order to determine the reaction condition for the azide-alkyne click chemistry,
we first performed a series of reactions in solution using model molecules (molecule A
and ethynylferrocene). Due to limited quantity of MCoTI-II, we tested this reaction at
low reactant concentrations (from 10 μM to 1mM). CuSO
4
and sodium ascorbate were
used as catalyst reagents and a ligand, tris-(triazolyl)amine (TBTA) was added to the
reaction mixture according to an establised bioconjugation protocol.
15-17
The reaction
was conducted in a 9:1 mixture of water and DMF in air. The reaction scheme is shown
in Figure 4.3 and the reaction yields (determined by LC-MS analysis) are summarized in
Table 4.1 for different reaction conditions. When we use 10 mol% of CuSO
4
and 20
mol% of sodium ascorbate without the ligand, the reaction at three reactant
concentrations didn’t proceed. We attribute this to the low concentration of reactants
P
O
O
O
N
3
P
O
O
O
N
N
N
Cyclotide
OH OH
P Br
O
O
O
P N
3
O
O
O
NaN
3
,MeCN
Reflux, 48hours
P N
3
O
HO
OH
i. BrMe
3
Si, MeCN, 24hr
ii. MeOH/H2O,2hr
AB
(a)
(b)
Figure 4.2 (a) Synthesis of bifunctional linker B. (b) Surface chemistry strategy for
immobilizing alkyne-terminated MCoTI-II onto In
2
O
3
NWs.
60
because we were able to obtain nearly 100% yield when both reactants are at 10 mM.
We then increased the concentration of CuSO
4
and sodium ascorbate to 1 mM and 2 mM,
respectively, and we were able to detect the product in LC-MS analysis. The yield
dropped dramaticaly when the reactant concentration decreased from 1 mM to 10 μM.
After adding 2 mM ligand into the reaction system, the yield was greatly improved for all
three reactant concentrations.
Although above experiments revealed the effect of TBTA in improving the
reaction yield, we found that it may cause complications for the reaction between
molecule A and MCoTI-II. 100 μM molecule A and MCoTI-II reacted for 4 hours with
the presense of 1 mM CuSO
4
, 2 mM sodium ascorbate and 2 mM TBTA, and the reaction
mixture was analyzed by HPLC followed by electrospray mass spectrometry (ES-MS)
(Figure 4.4a). The HPLC trace showed that the reaction produced multiple side products.
Reactant
concentration
20% sodium ascorbate
10% CuSO
4
?5H
2
O
2 mM sodium ascorbate
1 mM CuSO
4
?5H
2
O
2 mM sodium ascorbate
1 mM CuSO
4
?5H
2
O
2 mM TBTA
1 mM 0 73% 98%
100 µM 0 42% 94%
10 µM 0 12% 71%
Table 4.1 Reaction yield of molecule A and alkyne-terminated ferrocene under different
reaction conditions.
P
O
EtO
OEt
N
3 + Fc P
O
EtO
OEt
N
N
N
Fc
TBTA
N
N
N
N
Ph
3
CuSO
4
Sodium ascorbate
Figure 4.3 Reaction between molecule A and alkyne-terminated ferrocene.
61
We then did the same reaction without TBTA, the result (Figure 4.4b) indicated that we
obtained a much cleaner product mixture. The original cyclotide at 17.1 min was fully
consumed and the product showed up at 18.6 min with the correct molecular mass. It
appears to us that TBTA may cause degradation of MCoTI-II and therefore can’t not be
used in tthis reaction. We decided to use the reaction condition of 1 mM CuSO
4
and 2
mM sodium ascorbate for further experiments.
min 0 5 10 15 20 25
No r m .
0
20
40
60
80
100
M W D1 A , S ig = 220,1 6 R e f= 8 0 0,100 ( L U IS \0 1 2110 0 0 0 0 0 2.D )
Product
18.6 min
(MS~ 3657)
21.4 min
17.1min
Original cyclotide
min 0 5 10 15 20 25
No r m .
0
20
40
60
80
100
M W D1 A , S ig = 220,1 6 R e f= 8 0 0,100 ( L U IS \0 1 2110 0 0 0 0 0 1.D )
Product
18.6 min
(MS= 3657)
21.4
23
24.8
25.9
16.8
(a)
(b)
Figure 4.4 HPLC traces of reaction mixtures with (a) and without (b) TBTA. Reactants:
100 μM molecule A and 100 μM MCoTI-II. Catalyst: 1 mM CuSO
4
and 2 mM sodium
ascorbate.
62
4.2.3 Surface click chemistry
We then applied the click reaction on planar and nanostructured surfaces and the
results were confirmed by cyclic voltammetry (CV) and fluorescent imaging. A
monolayer of molecule B was first assembled on indium tin oxide (ITO)-coated glass
(Figure 4.5a) and it showed no redox peaks when scanned by CV (Figure 4.5b, black
trace). The azide-derivatized ITO slide, together with a bare ITO slide, were exposed to a
solution of ethynylferrocene under above mentioned reaction conditions and then
examined by CV. After reacting with ethynylferrocene, the azide-derivatized ITO
showed redox peaks correspoding to ferrocene (Fc) moiety (red trace) while the bare ITO
showed no peaks (blue trace). By this experiment we were able to confirm the covalent
binding of ethynylferrocene to the azide-modified ITO.
The monolayer of molecule B was then assembled on In
2
O
3
NW mats and alkyne-
terminated MCoTI-II was immobilized on the surface (Figure 4.5c). FITC labeled
trypsin was used to visualize the presence of MCoTI-II under fluorescent microscope
(Figure 4.5d). A control experiment was carried out where FITC-trypin was directly
added to the azide-derivatized In
2
O
3
NW mats. The fluorescent image of this sample
(Figure 4.5e) apears dark.
63
4.2.4 Binding affinity of surface bound MCoTI-II towards trypsin
It has been reported that free MCoTI-II binds to trypsin at a binding affinity of 25
pM.
18
However, when it is immolized on surfaces, the binding affinity might change due
to steric hinderence or conformational changes. In order to determine the binding affinity
of surface bound MCoTI-II towards trypsin, we performed kinetic analysis using surface
plasmon resonance (SPR) with a custom-derivatized sensor chip. As shown in Figure
4.6a, we started with a standard Biacore CM5 chip which has carboxyl groups on top of a
-0.2 0.0 0.2 0.4 0.6 0.8
-1.5x10
-5
-1.0x10
-5
-5.0x10
-6
0.0
5.0x10
-6
1.0x10
-5
1.5x10
-5
Current (A)
potential (V)
N3 modified ITO
N3 modified ITO +Fc
Blank ITO+Fc
ITO-coated
glass
P
N
O
Fc
N
N
O O
P
N
O
O O
N
N
Fc
Covalently attached Fc
(a)
(b)
P
O
O
O
N
3
P
O
O
O
N
N
N
Cyclotide
P
O
O
O
N
N
N
Cyclotide
Trypsin-FITC
P
O
O
O
N
3
P
O
O
O
N
3 Trypsin-FITC
+
100 祄
100 祄
(c)
(d)
(e)
Figure 4.5 (a) Surface functionalization of ITO-coated glass with molecule B and
ethynylferrocene. (b) CV traces for azide-modified ITO (black), after coupled with
ethynylferrocene (red) and a bare ITO exposed to ethynylferrocene (blue). (c) Surface
functionalization of In
2
O
3
NW mats with molecule B and MCoTI-II. (d) and (e) are
fluorescent images of the top and bottom sample illustrated in (c), respectively.
64
layer of dextran matrix. Streptavidin (SA) was immobilized onto the surface via the
amine coupling reaction using a Biacore T100 system and biotinylated MCoTI-II was
then attached to the SA layer via the specific binding between SA and biotin. A low
immobilization level (20 resonance unit (RU)) was mantained for biotinylated MCoTI-II
due to the requirement of SPR kinetic analysis. Once we derivatized the sensor chip
surface with MCoTI-II, we flow different concentrations of trypsin solutions over the
sensor chip and sensorgrams for each concentration are shown in Figure 4.6b. From the
sensorgrams we were able to calculate the binding affinity of surface bound MCoTI-II
(0.66 nM) using the Biacore T100 evaluation software. Compared to the free MCoTI-II,
surface bound ones exhibit lower binding affinity as expected.
65
4.2.5 Real-time sensing of trypsin using MCoTI-II derivatized In
2
O
3
NW sensors
We then perfomed real-time sensing of trypsin using MCoTI-II derivatized In
2
O
3
NW sensors. In
2
O
3
sensors were fabricated according to our establised protocols and
functionalized with monolayer coatings of molecule B. Alkyne-terminated MCoTI-II
was then attached to the surface via click chemistry and the final configuration of the
sensor is shown in Figure 4.7a. MCoTI-II derivatized sensors were placed under a Teflon
mixing cell filled with 0.01X PBS buffer (Figure 4.7b). During the real-time sensing, a
source-drain voltage of 200 mV and a liguid-gate voltage of 200 mV were applied to the
Carboxyl group
Amine
coupling
Ligand
capture
Streptavidin level :
~3000 RU
Biotinylated-MCoTI-II
level: ~20 RU
Biacore CM5 chip
Trypsin:
600 nM
2.5 nM
Binding Dissociation
(a)
(b)
Figure 4.6 (a) Surface derivatization strategy for Biacore CM5 sensor chip. (b) Kinetic
analysis sensorgrams for surface bound MCoTI-II and trypsin.
66
sensor and the source-drain current was monitored constantly. We first established a
stable baseline and then trypsin solutions of different concentrations were added into the
cell progressively. At the end of the sensing, we also added a non-target protein,
chymotrypsin, to demonstrate the selectivity of the sensor. The real-time sensing result is
shown in Figure 4.7c with each addition marked and the numbers above the arrows
represent the final concentration of trypsin or chymotrypsin after each addition. At
10 ng/ml of trypsin, the source-drain current started to drift downwards. However, the
change of signal in the next few steps is not proportional to the tryspin concentration.
Also, the sensor responded to 10 ug/ml of chymotrypsin in the same way as to 10 ug/ml
of trypsin. Therefore, in this sensing experiment, we were not able to obtain sensing
signals specific to trypsin.
67
4.3 Chapter Conclusions
In this chaper, we introduced a new class of small affinity binding agents,
cyclotides, as capturing probes in nanobiosensing. MCoTI-II was integrated with In
2
O
3
NW sensors for trypsin sensing. We investigated the bioconjugation condition for
alkyne-terminated MCoTI-II and achieved fluorescent detection of trypsin using
1000 1500 2000 2500 3000
285
290
295
300
305
310
315
320
325
330
Cu rren t (nA )
Tim e (s)
0.01X
PBS
10ng/ml
100ng/ml
1ug/ml
5ug/ml
10ug/ml
10ug/ml
Chymotrypsin
D
S
Gate (Si)
P
O
O
O
N
N
N
Cyclotide
P
O
O
O
N
N
N
Cyclotide
10ul trypsin solution
(a)
(b)
(c)
Figure 4.7 (a) Schematic view of a MCoTI-II derivatized In
2
O
3
NW sensor. (b)
Schematic view of the sensing setup. (c) Real-time sensing result for trypsin using a
MCoTI-II derivatized In
2
O
3
NW sensor.
68
MCoTI-II derivatized In
2
O
3
NW mats. We also determined the binding affinity of
surface bound MCoTI-II towards trypin by SPR kinetic analysis. Finally, we performed
preliminary experiments on real-time, electrical detection of trypsin using MCoTI-II
derivatized In
2
O
3
NW biosensors.
4.4 Experimental Section
Materials All chemicals were purchased from Sigma-aldrich except otherwise
stated. Alkyne-terminated MCoTI-II and biotinylated MCoTI-II were provided by
Professor Julio Camarero. TBTA was synthesized according to literature procedures.
19
FITC-trypsin was prepared using FITC labeling kit from Sigma-aldrich. SPR sensor
chips were purchased from GE healthcare. Buffers used in SPR sensor chip
derivatization were 1X PBS buffer containing 0.005% (v/v) tween-20. Buffers used in
SPR kinetic study were 1X PBS buffer containing 0.005% (v/v) tween-20 and 0.2 mg/ml
BSA. All SPR buffers were filtered by 0.22 μm membrane.
Azide-alkyne click chemistry reaction All azide-alkyne reactions were
performed with CuSO
4
and sodium ascorbate in a 9:1 mixture of water and DMF and the
reaction time was 4 hours. Ligand TBTA was added when necessary. The reactions in
solutions were conducted in closed vials with no further efforts to exclude oxygen while
the reactions on surfaces were conducted in a nitrogen-purged glovebag.
HPLC-MS HPLC-MS analysis for small molecules was performed on a
Shimadzu LCMS-2020 system with 270 nm detection using a C8 column. All runs used
linear gradients of 0-70% over 15 min of water (solvent A) vs. acetonitrile (solvent B).
HPLC analysis for reaction mixtures of MCoTI-II was performed on a HP1100 series
69
instrument with 220 and 280 nm detection using a Vydac C18 column (5 micron, 4.6 x
150 mm) at a flow rate of 1 mL/min. All runs used linear gradients of 0-70% over 30min
of 0.1% aqueous trifluoroacetic acid (TFA, solvent A) vs. 0.1% TFA, 90% acetonitrile in
H
2
O (solvent B). UV/Vis spectroscopy was carried out on an Agilent 8453 diode array
spectrophotometer. ES-MS was performed on a Sciex API-150EX single quadrupole
electrospray mass spectrometer.
Surface functionlization of ITO and In
2
O
3
NWs The monolayer of molecule B
was assembled on ITO and In
2
O
3
NWs according to procedures described in chapter 2.
Ethylnylferrocene and alkyne-termianted MCoTI-II was attached to surface as described
above and any unbound molecules were removed by multiple rinsing steps.
Electrochemistry CV for ITO samples was performed with
Potentiostat/Galvanostat Model 263A (Princeton Applied Research) in an
electrochemical cell filled with PBS buffer (pH 7.4), with Pt wire as the counter electrode
and Ag/AgCl as the reference electrode.
Fluorescent imaging The fluorescence images were captured using a Nikon
Eclipes ME600 fluorescence microscope equipped with a Microfire digital color camera
at 20X magnification.
SPR Kinetic analysis for surface bound MCoTI-II and trypsin were carried out
using Biacore T100 system. Carboxymethylated dextran on Biocore CM5 chip surface
was activated with the mixture 0.2 M EDC/0.05 M NHS injected over the sensor chip for
7 min at a flow rate 10 μl/min and washed with PBS buffer for 3 min at the same flow
rate. Subsequent immobilization of SA was carried out by injecting SA solution
(100 μg/ml in 10 mM acetate buffer (pH 5.0) during 7 min at a flow rate 10 μl/min) over
70
the activated sensor surface. The residual active groups of dextran were blocked by 1 M
ethanolamine (pH 8.5) for 7 min at a flow rate 1 μl/min. A solution of biotinylated
MCoTI-II in PBS was injected over the SA surface until a immobilization level of 20 RU
was achieved. Interaction of trypsin with the immobilized MCoTI-II was studied using
the range of concentrations 2.5-600 nM. After injection of each trypsin sample the chip
was regenerated by injection of glycine solution (pH 2.5) for 20 sec at a flow rate
10 μl/min. A channel without the immobilized MCoTI-II was used as the reference.
Kinetic parameters for the reaction of affinity binding were calculated using the program
Biacore T100 evaluation software and the mathematical model 1 : 1 Langmuir binding.
Real-time sensing MCoTI-II functionalized sensors were initially immersed in
0.01X PBS and the real-time source-drain current was monitered by Agilent B1500
analyzer. Trypsin solutions (prepared in 0.01X PBS) were added into the mixing cell
from low to high concentrations. The non-target chymotrypsin solution (final
concentration 10 μg/ml in 0.01X PBS) was added at the end of the sensing experiment to
test the sensors’ selectivity.
4.5 References
(1) Gould, A.; Ji, Y.; Aboye, T. L.; Camarero, J. A. Current Pharmaceutical
Design, 17, 4294-4307.
(2) Cui, Y.; Wei, Q. Q.; Park, H. K.; Lieber, C. M. Science 2001, 293, 1289-
1292.
(3) Allen, B. L.; Kichambare, P. D.; Star, A. Advanced Materials 2007, 19,
1439-1451.
(4) Bunimovich, Y. L.; Shin, Y. S.; Yeo, W.-S.; Amori, M.; Kwong, G.;
Heath, J. R. Journal of the American Chemical Society 2006, 128, 16323-16331.
(5) Curreli, M.; Zhang, R.; Ishikawa, F. N.; Chang, H.-K.; Cote, R. J.; Zhou,
C.; Thompson, M. E. Ieee Transactions on Nanotechnology 2008, 7, 651-667.
(6) Gruner, G. Analytical and Bioanalytical Chemistry 2006, 384, 322-335.
71
(7) 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. Acs
Nano 2009, 3, 1219-1224.
(8) Ishikawa, F. N.; Curreli, M.; Chang, H.-K.; Chen, P.-C.; Zhang, R.; Cote,
R. J.; Thompson, M. E.; Zhou, C. Acs Nano 2009, 3, 3969-3976.
(9) Li, C.; Curreli, M.; Lin, H.; Lei, B.; Ishikawa, F. N.; Datar, R.; Cote, R. J.;
Thompson, M. E.; Zhou, C. W. Journal of the American Chemical Society 2005, 127,
12484-12485.
(10) Patolsky, F.; Zheng, G.; Lieber, C. M. Nanomedicine 2006, 1, 51-65.
(11) 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. Nature 2007,
445, 519-522.
(12) Craik, D. J.; Daly, N. L.; Bond, T.; Waine, C. Journal of Molecular
Biology 1999, 294, 1327-1336.
(13) Rosengren, K. J.; Daly, N. L.; Plan, M. R.; Waine, C.; Craik, D. J. Journal
of Biological Chemistry 2003, 278, 8606-8616.
(14) Colgrave, M. L.; Craik, D. J. Biochemistry 2004, 43, 5965-5975.
(15) Chan, T. R.; Hilgraf, R.; Sharpless, K. B.; Fokin, V. V. Organic Letters
2004, 6, 2853-2855.
(16) Gurcel, C.; Vercoutter-Edouart, A.-S.; Fonbonne, C.; Mortuaire, M.;
Salvador, A.; Michalski, J.-C.; Lemoine, J. Analytical and Bioanalytical Chemistry 2008,
390, 2089-2097.
(17) Wang, Q.; Chan, T. R.; Hilgraf, R.; Fokin, V. V.; Sharpless, K. B.; Finn,
M. G. Journal of the American Chemical Society 2003, 125, 3192-3193.
(18) Camarero, J. A.; Kimura, R. H.; Woo, Y.-H.; Shekhtman, A.; Cantor, J.
Chembiochem 2007, 8, 1363-1366.
(19) Lee, B.-Y.; Park, S. R.; Jeon, H. B.; Kim, K. S. Tetrahedron Letters 2006,
47, 5105-5109.
72
CHAPTER FIVE: PolySilicon microribbon biosensors: surface functioanlization
and sensing applications
5.1 Introduction
In recent years, label-free, electrical nanobiosensors have drawn lots of research
interest due to the potential of achieving superior time and cost efficiency to current state-
of-the-art biosensing platform such as ELISA. Among nanobiosensors studied by various
research teams, most of them are fabricated by “bottom-up” technique, that is,
nanostructures are assembled to make devices.
1-7
One of the major challenges for
nanosensors fabricated by bottom-up technique is assembly, which can significantly limit
the yield and uniformity of such nanosensors.
8
The yield is highly related to cost and
throughput, and uniformity is essential to the reliability of nanobiosensors. Although
intensive research efforts have been made toward assembly of nanostructures, most of the
techniques still lack controllability, reproducibility and scalability. The other school of
process in nanotechnology is “top-down” fabrication, which seeks to create nanoscale
devices by using larger, externally-controlled ones to direct their assembly. Top-down
fabrication is much more controllable than its bottom-up counterpart, thus leads to more
uniform device performance. As a result, nanobiosensors fabricated using top-down
approaches can yield more reliable sensing results. One of the major challenges for top-
down nanobiosensors, however, is to achieve large surface-to-volume ratio, as surface-to-
volume ratio is directly linked to sensitivity. Research efforts have been made toward
reducing the critical dimension of nanostructures (hence increase surface-to-volume ratio)
by applying techniques such as electron beam lithography and directional etching.
8-11
However, such techniques are extremely time-consuming, cost-inefficient, and have poor
73
scalability. Those drawbacks significantly limit the commercial impact the
aforementioned nanobiosensors can potentially generate. In this regard, a recent study
demonstrates that highly sensitive nanobiosensors can be achieved without pushing
critical dimensions to a limit.
12
By carefully limiting the active layer thickness, silicon
nanoribbon based biosensors can be fabricated by conventional photography (~µm
critical dimension) with clinically relevant sensitivity. Despite the promising result,
single-crystalline silicon on insulator (SOI) wafers with an extremely thin (~50nm) active
layer are required to produce the nanobiosensors. SOI wafers with such thin active layers
are not easily available, and thus can be very expensive. Also, precise oxidation and wet
etching are needed to achieve the desired thickness, further limiting time and cost
efficiency of such platform.
In this chapter, we describe a nanobiosensor platform based on polysilicon
nanoribbons. The devices are fabricated with ~100% yield and great uniformity due to
top-down process. The polysilicon is deposited using a highly scalable, precise and cost-
efficient low-pressure chemical vapor deposition (LPCVD). Unlike SOI wafers, the
thickness of polysilicon active layer and dielectric layer can be well controlled and easily
customized for different applications. The whole process is compatible with
conventional photolithography with only two masks required, thus are incredibly time
and cost efficient. Moreover, the fabrication can be performed on full wafers, which
results in great scalability. By performing pH sensing experiment with a wide dynamic
range and high sensitivity, we demonstrate that the polysilicon nanoribbon sensors are
highly sensitive to ions. Finally biomarker detection is performed with clinically relevant
sensitivity, and thus confirms the practical value of polysilicon nanoribbon biosensors.
74
Multiple devices are monitored simultaneously during all sensing experiments, and the
response shows great uniformity. We demonstrate that polysilicon nanoribbon can act as
a highly efficient, reliable and scalability platform for nanobiosensors. Such a platform
exhibits great potential toward label-free, electrical biosensor with enormous practical
impact.
5.2 Results and discussion
This project was a collaborative effort of Thompson group and Dr. Chongwu
Zhou’s group in electrical engineering department, USC. The device fabrication and
performance data was obtained by Hsiaokang Chang, Shelley Wang and Noppadol
Aroonyadet in Zhou’s group. Surface chemistry was done by myself with the help of
Yan Song in Thompson group. pH sensing and biomarker sensing was performed by
Hsiaokang Chang and myself.
5.2.1 Device fabrication and performance
The FET fabrication process is shown in Figure 5.1. The fabrication starts with a
simple silicon oxide on silicon wafer as in Figure 5.1a. The oxide thickness is
determined by the desired dielectric thickness (500 nm in our work). A thin layer
(typically 50 nm) of polysilicon is deposited via LPCVD and doped with boron at desired
doping concentrations (shown in Figure 5.1b). Photolithography and a CF
4
dry etch are
then performed to define the contact lead and nanoribbon area (Figure 5.1c). A second
photolithography followed by thermal evaporation is performed to define the metal
contact (5 nm Ti and 45 nm Au) (Figure 5.1d). Finally, a nitride passivation layer can be
75
deposited via plasma-enhanced chemical vapor deposition (PECVD) to cover metal
electrodes if necessary. The overall fabrication process is highly efficient and scalable
with only two masks required and all the steps involved can be performed on wafer scale.
Shown in Figures 5.1e and 5.1f is a SEM image of a group of 6 polysilicon nanoribbons
and a photographic image of hundreds of polysilicon nanoribbon sensors fabricated on a
3” wafer, respectively. The devices are fabricated at a yield of nearly 100% and exhibit
very little device-to-device variation due to the controllable top-down process.
In our work, we tested three different doping concentrations, 1×10
17
, 5×10
17
and
1×10
18
, and FETs were fabricated using each concentration. The electrical properties of
Figure 5.1 (a)-(d) show the fabrication process of polysilicon nanoribbon FETs. (e) and
(f) is a SEM image of a group of 6 polysilicon nanoribbons and a photographic image of
hundreds of polysilicon nanoribbon sensors fabricated on a 3” wafer, respectively.
76
these devices were characterized using a back-gate and the results are shown in Figure
5.2. Plotted in Figure 5.2 a and b is the source-drain current (I
DS
) v.s. source-drain
voltage (V
DS
) characteristics under various back-gate voltage (V
GS
) and I
DS
v.s. V
GS
under various V
DS
, respectively, for a device with 1×10
17
doping. The device exhibits
ohmic contact behavior with strong gate dependence. The on/off ratio reaches 250 at V
DS
=1 V and is high enough for nanobiosensor application. This value can be further
improved by using dielectrics with higher dielectric constant or by applying the gate
voltage via liquid-gate. The I
DS
v.s. V
DS
and I
DS
v.s. V
GS
characteristics for a device with
5×10
17
doping concentration are shown in Figure 5.2 c and d, respectively. The device is
about 10 times more conductive compared to the 1×10
17
device. However, the on/off
ratio is reduced to 62 at the same V
DS
. Data for a device with 1×10
18
doping
concentration is shown in Figure 5.2 e and f. The device exhibits even higher
conductance and lower on/off ratio compared to the 5×10
17
doping concentration. Base
on this study, it is clear that doping level is positively related to the FET conductance but
is negatively related to the on/off ratio. Our previous experiences with nanosensors
indicate that devices with high on/off ratio usually yield better sensitivity during sensing.
Therefore the 1×10
17
doped devices are selected for further sensing experiments in this
chapter unless otherwise stated.
77
5.2.2 pH sensing
We preformed pH sensing to demonstrate that the polysilicon FETs are ion-
sensitive. During the sensing experiment, the device was exposed to buffer solutiolns of
different pH and the I
DS
was monitored constantly. The device was first soaked in pH 4
Figure 5.2 The I
DS
v.s. V
DS
and I
DS
v.s. V
GS
characteristics for devices with 1×10
17
,
5×10
17
and 1×10
18
doping concentration.
78
buffer using a mixing cell and then the liquid was replaced by equal volume of pH 6
buffer. The device showed significant increase in conductance after the buffer exchange,
as shown in Figure 5.3a, and such increase in conductance can be explained by the p-type
transistor behavior of the polysilicon FETs. Polysilicon is known to be covered by a
layer of native silicon oxide. When the pH value of surrounding buffer increases, the
surface hydroxyl groups are deprotonated and the negative charges on the surface induce
an accumulation of charge carriers in polysilicon. Therefore an increasement in
conductance was observed. Further buffer exchanges to higher pHs caused constantly
increase in device conductance and devices showed a wide dynamic range to pH
variation (pH 4 to pH 10). Buffers were then exchanged back to lower pHs to show the
reversibility of device responses to ions. Further pH sensing experiments were carried
out with a pH step of 0.2 around neutral pH, and the result is shown in Figure 5.3b. The
device shows a response of ~10% with high signal-to-noise ratio even to a 0.2 variation
in pH, suggesting the polysilicon nanosensor is highly sensitive to ions.
79
5.2.3 Surface functionalization of polysilicon with native oxide layer
In order to use polysilicon nanoribbon FETs as nanobiosensors, we need to
functionalize the surface of active area (the ribbon area) with certain biorecognition
pH 4 pH 6 pH 8 pH 10 pH 8 pH 6 pH 4
pH 7.2 pH 7.4 pH 7.6 pH 7.8 pH 8
(a)
(b)
Figure 5.3 Real-time pH sensing using a polysilicon FET at pH range of (a) pH 4 ~10
and (b) pH 7.2 ~ 8.
80
molecules, such as antibodies and oligonuleotides, and these molecules can searve as
capturing agents for the detection of desired biomarkers. There have been numerous
reports of surface functioanlization of silicon nanowire-based FETs, and the most
common method is to covalently attach capturing agents to the native oxide layer using
alkoxysilane linkers.
13-16
The strategy used to functionalized polysilicon is shown in
Figure 5.4a. The pre-cleaned polysilicon wafers were functionalized with a biotin
derivative and the successful attachment of this molecule was confirmed by both x-ray
photoelectron spectroscopy (XPS) and fluorescent imaging. In our work, we chose a
monoalkoxysilane derivative, 3-aminopropyldimethylethoxy silane (APDMS), instead of
the commonly used trialkoxysilanes to prevent the formation of a thick polymer film due
to self-polymerization. The terminal amine groups were first converted to carboxyl
groups by reacting with succinic anhydride in the presense of triethylamine, and then an
amine-terminated biotin molecule (amine-PEG-biotin) was attached to the surface by the
NHS/EDC coupling reaction.
XPS high-resolution spectra of the N 1s region is plotted in Figure 5.4b for
samples after each functionalization step. The increase in peak intensity from the black
trace (oxide surface) to the blue trace (APDMS surface) indicates the formation of silane
monolayer. After converting the surface to carboxyl groups, the peak intensity (red trace)
remains almost the same since the number of N atoms per molecule didn’t change. The
intensity increased by a factor of ~2.5 from the red trace to the green trace (biotin surface)
after calibrating the N 1s signal against each sample’s Si 2p signal. Considering that each
amine-PEG-biotin molecule contains 4 N atoms, we can calculate that the yield of the
biotin immobilization reaction is roughly 50%. The surface functionalization with biotin
81
derivative is further confirmed by fluorescent imaging with the aid of a fluorescent
labeled streptavidin molecule (Figure 5.4c). The polysilicon wafer used as control was
directly exposed to amine-PEG-biotin and then fluorescent streptavidin right after pre-
cleaning step, and showed no fluorescence in Figure 5.4d.
82
Si
NH
2
O
Si
NH
O
HO
O
O
Si
NH
O
HN
O
O
PEG-Bioti n
OHOH
Si
NH
O
HN
O
O
PEG-Bi otin
APDMS
Succinimide
anhydride
NH
2
-PEG-biotin SA-dye
(a)
(b)
405 400 395
Oxide
APDMS
COOH
Biotin
Intensity (a.u.)
Binding Energy (eV)
(d) (c)
Figure 5.4 (a) Scheme of surface functionalization of polysilicon using a biotin
derivative. (b) XPS high-resolution spectra of the N 1s region after each step of
functionalization. (c) Fluorescent image of the biotin-functionalized polysilicon after
exposed to fluorescent labeled streptavidin. (d) Fluorescent image of a polysilicon
wafer as control. The wafer was directly exposed to amine-PEG-biotin and then
fluorescent streptavidin right after pre-cleaning step.
83
5.2.4 Biomarker sensing
Biomarker detection was also performed using polysilicon nanoribbon sensors.
We chose cancer antigen 125 (CA-125), an ovarian cancer biomarker, as the target of
study. The polysilicon devices were first conjugated with CA-125 antibody so that they
could recognize CA-125 specificly. During the sensing experiment, a 200 mV V
DS
and
a -200 mV V
GS
were applied to the devices and the source-drain currents of three devices
were monitored simultaneously. CA-125 solutions of different concentrations were
progressively added to the sensing environment.
The normalized current versus time for three devices is plotted in Figure 5.5.
Note that three devices showed uniform responses to the presence of CA-125 at each
concentration. All three devices started to show response at CA-125 concentration of
10 U/ml (50 pM). This limit of detection is one order of magnitude lower than the
clinically relevant level for diagnosis (100 ~ 275 U/ml). The devices showed larger
responses when CA-125 of higher concentrations were added. In comparison, none of
the devices showed response to the addition of bovine serum albumin (BSA) at 150 nM
(data shown in inset Figure 5.4), a concentration much higher than that of the target
analyte. The sensing data showed that the polysilicon nanosensor could detect
biomarkers with clinically relevant sensitivity and great selectivity, suggesting that
polysilicon nanoribbon biosensors have great potential as a sensing platform for clinical
diagnosis of diseases such as cancers.
84
5.2.5 Surface functionalization of polysilicon without native oxide layer
The insulating layer of native SiO
2
on the surface of polysilicon nanoribbons
screens out the electrical signals generated from binding of target molecules that we use
for our real-time sensing.
17
In order to achieve higher sensitivity, we also explored the
functionalization methods that are directly applied to silicon hydride surface. The
processing starts by removing the native oxide using HF etching and leaving a
hydrogen-terminated silicon surface. The method involves using UV light to rapidly
photo-dissociate the Si-H bond to generate radical species on the polysilicon surface.
These radicals can subsequently react with terminal olefin groups on the linker molecule,
5-hexenoic acid, thus forming stable Si-C bonds at the Si surface.
17-19
The linker
molecule usually carries a carboxyl group, which can be used to attach biological probes
Figure 5.5 Real-time sensing of CA-125 using polysilicon nanoribbon biosensors.
Signals from three sensors were monitored simultaneously. A non-target protein, BSA,
was added to the sensing media at the end of the sensing experiment to demonstrate the
sensors’ selectivity. This part of sensing signal is shown in inset.
85
for further functionalization. In this case, an amine-PEG-biotin was attached by the
NHS/EDC coupling reaction (Figure 5.6a) and the result was confirmed by both XPS and
fluorescent imaging. Figure 5.6b shows the XPS survey spectra of two hydrogen-
terminated polysilicon wafers after being immersed in 5-hexenoic acid in N
2
for 4 hours.
With UV illumination, the surface (red trace) showed much higher signal in the C 1s
region than the one in dark (black trace), indicating the covalent attachment of
5-hexenoic acid on the surface. The two samples were then treated by amine-PEG-biotin
as mentioned above and exposed to fluorescently labeled steptavidin. Fluorescent images
of the two samples are shown in Figure 5.6c (UV) and Figure 5.6d (Dark), confirming the
successful functionalization of polysilicon with the aid of UV illumination.
86
5.3 Chapter Conclusions
Polysilicon nanoribbon field-effect transistors (FETs) were fabricated uisng a
simple two-mask photolithography method on a wafer scale. Three doping
concentrations, 1×10
17
, 5×10
17
and 1×10
18
were tested and the FETs with 1×10
17
doping
concentration were identified as the best candidate for further sensing experiments. pH
sensing was performed with these FETs in two pH ranges and it demonstrated the
devices’ high sensitivity to ions. Biofunctionalization of polysilicon surface were
H H
OH OH
O O
UV, 4 hours
NH NH
O O
PEG PEG
Biotin Biotin
NH
2
-PEG-biotin
OH
O
1000 800 600 400 200 0
UV
Dark
Intensity (a.u.)
Binding Energy (eV)
O 1s C 1s
Si 2s
Si 2p
(b)
(a)
(c)
(d)
Figure 5.6 (a) Scheme of surface functionalization strategy of hydrogen-terminated
polysilicon. (b) XPS survey spectra of two hydrogen-terminated polysilicon samples
after being immersed in 5-hexenoic acid in N
2
for 4 hours with (red) and without
(black) UV illumination. (c) and (d) are fluorescent images of these two samples after
being treated with amine-PEG-biotin and fluorescently labeled streptavidin. (c) is the
image of the UV illuminated sample and (d) is the image of the sample kept in dark.
87
conducted with and without the native oxide layer and the results were confirmed by both
XPS and fluoresent imaging. Finally, electrical, label-free detection of a cancer
biomarker, CA-125, was performed using three polysilicon nanoribbon sensors. The
sensing results indicated the potential of these FETs to be used as nanobiosensors with
promising sensitivity and selectivity.
5.4 Experimental Section
Materials and instruments Spin-on dopant was purchased from Emulsitone
cooperation. APDMS was purchased from Alfa Aesar. Amine-terminated biotin was
purchased from Pierce. Streptavidin-Alexa fluro 568 was purchased from Invitrogen.
CA-125 antibody and antigen were purchased from Fitzgerald. All other chemicals were
purchased from Sigma-Aldrich. UV-assisted reaction was performed in a Luzchem ICH-
2 photoreactor. XPS was performed on an M-probe surface spectrometer (VG
Instruments). Monochromatic Al KR X-rays (1486.6 eV) incident at 35 from horizontal
were used to excite electrons from the sample, and the emitted electrons were collected
by a hemispherical analyzer at a takeoff angle of 35 from the plane of the sample surface
(horizontal). Fluorescent images were taken by a Nikon Eclipse ME600 fluorescent
microscope equipped with an Microfire digital color camera. The electrical testing of
devices was performed by an Agilent 4156B semiconductor analyzer. The real-time
sensing was done using an Agilent B1500 semiconductor analyzer.
FET fabrication A thin layer (typically 50 nm) of polysilicon was deposited via
LPCVD on silicon substrate with a layer of oxide (500 nm). Spin-on dopant of desired
doping concentration was applied to the polysilicon surface via spin coating and a
88
subsequent drive-in annealing was performed at 1100°C for 15 minutes in nitrogen
environment. A buffered HF solution was then used to remove spin-on dopant and a CF
4
dry etch was then performed to define the contact lead and nanoribbon area. A second
photolithography was then performed to define the metal contact. 5nm Ti and 45nm Au
was deposited as electrode via thermal evaporation. A HF dipping was required before
the metal evaporation to remove native oxide. A nitride passivation layer can be
deposited on top of metal electrodes via PECVD if necessary.
pH sensing pH sensing was performed in 10 mM phosphate buffers of different
pHs. The total ionic strength of these buffers was adjusted to 100 mM by adding NaCl.
A polysilicon nanoribbon sensor was first inmmersed in the buffer of pH 4 using a teflon
mixing cell. A 200 mV source-drain voltage and a 200 mV liquid gate voltage were
applied to the sensor. The source-drain current is constantly monitored by a
semiconductor analyzer (Agilent B1500). During sensing, the sensor was exposed to
buffers of different pHs by changing the buffer in the mixing cell with a micropipette.
Surface functionalization - with oxide layer Polysilicon wafers or sensors were
first cleaned by boiling in aceton and IPA for 5 minutes each, and then treated by UV/O
3
for 10 minutes (2 minutes in the case of sensors). The cleaned wafers or sensors were
immediately transferred to a APDMS solution in dry tolune (2% v/v) and incubated for 2
hours in N
2
. They were then washed by fresh tolune and methanol and annealed under
N
2
at 120 C for 12 hours. The carboxyl surface were generated by immersing annealed
polysilicon in a 5 mg/ml succinic anhydrous solution in dry THF (containing 5% v/v
triethylamine) for 4 hours in N
2
. The functionalized polysilicon was then subject to the
standard NHS/EDC coupling reaction to attach amine-terminated biotin or antibodies.
89
For fluorescent imaging experiments, the biotinylated sample was exposed to a solution
of streptavidin-Alexa fluro 568 (in 1X PBS) for 1 hour and then rinsed by 1X PBS for 3
times.
Surface functionalization - without oxide layer Polysilicon wafers were first
cleaned by solvent boiling and UV/O
3
as discribed above, and then dipped into a
deoxygenated HF solution (2%) for 30 seconds. The wafers were then quickly rinsed
with D.I. water and transferred into a pre-dried custom-made quartz tube. The tube was
immediately pumped down to vacuum to ensure a oxygen-free enviorentment. After
back-filling the tube with N
2
, a deoxygenated solution of 5-hexenoic acid in dry methanol
was added onto the polysilicon wafers. A UV light of 256 nm was shined on the
polysilicon wafers and the reaction was allowed to run for 4 hours. After that, the wafers
were taken out of the tube and sonicated in fresh methanol for 3 times, 5 minutes each
time. Amine-ternimated biotin was attached to the surface via the NHS-EDC coupling
reaction. Fluorescent imaging was done as discribed above.
Biomarker sensing Polysilicon nanoribbon sensors with native oxide were
functionalized with CA-125 antibody and immersed in 0.01X PBS. During the sensing
experiment, a 200 mV V
DS
and a -200 mV V
GS
were applied to the devices and the
source-drain currents of three devices were monitored simultaneously by an Agilent
B1500 analyzer. CA-125 antigen solutions (prepared in 0.01X PBS) were added into the
mixing cell from low to high concentrations. The non-target BSA solution (10 mg/ml in
0.01X PBS) was added at the end of the sensing experiment to confirm the sensors’
selectivity.
90
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Abstract (if available)
Abstract
In the past 20 years, material scientists and engineers have progressively miniaturized the materials that constitute the building blocks of various biomedical devices. This progressive downscaling has led to the creation of materials with at least one critical dimension falling within the 1-100 nm range. These nanomaterials have been considered as ideal candidates for biosensing applications due to their high surface-to-volume ratio and small sizes. A diversity of sensor architectures has been designed and fabricated during the last decade that utilizes different nanomaterials as sensing elements. Among them, sensors based on FETs have drawn increasing attention because of their capability of performing rapid and label-free detections. Since the first demonstration of FET-based biosensors in 2001, people have achieved detections of proteins, oligonucleotides and viruses with high sensitivity and selectivity. However, to facilitate the widespread adoption of nanobiosensing technology, researchers still need to address a few challenges, including multiplexing, cost efficiency and signal reproducibility. This dissertation tries to tackle these challenges by improving nanosensor fabrication techniques and developing novel surface functionalization methods. ❧ Chapter 1 briefly introduces the fundamentals of nanobiosensors, including nanomaterial synthesis, device fabrication and sensing setup, and discusses current challenges in the nanobiosensing field. ❧ Chapter 2 describes a newly designed electroactive surface modifier that can be used in selective biofunctionalization of nanomaterials. This molecule offers promising control over the surface reaction of each sensor in an array and can be considered as a key element in the fabrication of high-density biosensor arrays for multiplexed biosensing in the future. ❧ Chapter 3 focuses on sensing platforms based on solution grown ZnO nanostructures. Mass production of ZnO nanowires and nanobelts are prepared using a low cost, low temperature hydrothermal method, and used to fabricate back-gated FETs. By applying a post-synthesis annealing step to the ZnO products, we adjust the doping level of the nanowires and nanobelts, and thus significantly improve their electrical properties. These FETs shows comparable performance with those based on ZnO nanostructures synthesized via vapor-phase approaches. ❧ Chapter 4 introduces a new class of small affinity binding agents, cyclotides, as capturing probes in nanobiosensing. These backbone-cyclized polypeptides with a disulfide-stabilized core are chemically more stable than conventional antibodies and can be produced in relatively large quantities at low cost. A cyclotide, MCoTI-II, was integrated with In₂O₃ NW mats and sensors for trypsin sensing in this chapter. ❧ In the end, chapter 5 studies top-down polysilicon nanoribbon sensors and their sensing applications. Top-down sensors are believed to have better device uniformity and thus can yield more reliable sensing signals. In this chapter, polysilicon nanoribbon FETs are fabricated using a simple two-mask photolithography method on a wafer scale and functionalized with and without the native oxide coating. The pH sensing and biosensing performed with these sensors demonstrates their promising sensitivity and selectivity.
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Asset Metadata
Creator
Zhang, Rui
(author)
Core Title
Surface functionalization of nanomaterials and the development of nanobiosensors
School
College of Letters, Arts and Sciences
Degree
Doctor of Philosophy
Degree Program
Chemistry
Publication Date
11/05/2012
Defense Date
08/02/2012
Publisher
University of Southern California
(original),
University of Southern California. Libraries
(digital)
Tag
biosensors,field-effect transistors,nanomaterials,OAI-PMH Harvest,surface functionalization
Language
English
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Advisor
Thompson, Mark E. (
committee chair
), Goo, Edward K. (
committee member
), Zhou, Chongwu (
committee member
)
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ruizhang@usc.edu,ruizhangusc@gmail.com
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UC11289928
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etd-ZhangRui-1266.pdf
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107242
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Dissertation
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Zhang, Rui
Type
texts
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University of Southern California Dissertations and Theses
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The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the a...
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Tags
biosensors
field-effect transistors
nanomaterials
surface functionalization