1. Detection of Oligonucleotide – Gold
Nanoparticle conjugates Using
Cantilever Arrays Operated in
Dynamic Mode
Larry O’Connell
08390860
School of Physics
Trinity College Dublin
Supervisors:
Prof. Martin Hegner
Ph.D. Student Jason Jensen
Nanobio-Nanomechanics Group
Centre for Research on Adaptive Nanostructures and Nanodevices
Trinity College, Dublin
December 2011
2. i
Abstract
This project endeavoured to demonstrate the ability of a micromechanical cantilever array-
based device, operating in dynamic mode, to detect binding to the cantilever surface of 12-
mer oligonucleotides in solution. The oligonucleotides were attached with a thiol bond to 50
nm diameter gold nanoparticles, while the complimentary oligonucleotide sequence was
similarly attached to the cantilever surface. Only non-specific binding was detected. The
sensitivity of the device was found to be approximately 3.4 pg/Hz.
3. ii
Preface
This final year project was conducted over a 2 month period from September 26th
until
November 25th
, with the Nanobio-Nanomechanics research group in the Centre for Research
on Adaptive Nanostructures and Nanodevices (CRANN), at Trinity College, Dublin (TCD),
under the supervision of Prof. Martin Hegner.
4. iii
Acknowledgments
I would like to thank my supervisor Prof. Martin Hegner for the opportunity to work on this
project. I would also like to thank all staff in the Nanobio-Nanomechanics group for creating
a welcoming atmosphere. Finally, I would like to thank Jason Jensen for his patience and
guidance, without which this project would not have been possible.
All image and photo credits are to the author unless otherwise stated.
7. 1
Chapter 1 Introduction
Micro fabrication techniques previously developed for the semiconductor industry
have found novel applications in producing research and diagnostic tools for the biological
sciences. Micromechanical systems provide a radical new way to carry out qualitative and
quantitative bioassays. In many such scenarios, it is desirable to analyse surface binding in
small fluid volumes to analyze precious samples or to make a large number of measurements
from a single sample. A typical and particularly promising example of such “lab-on-a-chip”
nanosensing technology is the cantilever array.
Cantilever arrays present a particularly versatile sensing technique, where analyte
solutions as dilute as picomolar concentrations have been detected6
. Cantilever arrays offer
the possibility of detection of analytes in environments as diverse as blood1
, urine2
, tap-
water3
, air4
etc. This can be achieved in a shorter timeframe, with greater automation, greater
redundancy, and better field portability than with present techniques.3
Also, since the extent
of binding is proportional to concentration in the sample, the sensor response is proportional
to analyte concentration3
and thus can produce quantative information about analyte
concentration. This type of sensing has been shown to have applications as diverse as:
genomics5
, proteomics6
, food engineering4
and chemistry7
. Dynamic-mode-operated
cantilevers have been shown capable of detecting bacterial pathogens in liquid, notably:
Enterrohemorrhagic Escherechia coli8
, Bacillus anthracis9
(aka Anthrax), Salmonella
typhimurium10
, and Cryptosporidium parvum11
. Thus, cantilever arrays also have debatable
salience as a method of detection of bioterrorism agents.12,13
The ability to detect arbitrary
DNA markers with a high degree of specificity for rapid detection of genetic mutations and
8. 2
Figure 1 – Schematic diagram of successful attachment of a nanoparticle-oligonucleotide conjugate to a
functionalized cantilever surface. Note the complimentarity between the cantilever-bound oligonucleotide sequence
and the nanoparticle-bound oligonucleotide sequence. The mass of many likewise attached gold nanoparticles will
alter the vibrational dynamics of the cantilever, lowering its eigenfrequencies.
disease states is an area of active research, and is essential to a number of new methods of
disease management.13,14
Presently, micro-litre scale bioassays are familiar in the context of certain kinds of
testing. For example, diabetics use a simple device that uses a small droplet of blood to
measure blood sugar level. A similar-scale test is available for testing for anaemia by
measuring blood Iron content. This contrasts sharply with methods for testing for disease
which rely on enrichment culture.
Enrichment culture methods necessitate taking large samples from a patient (e.g.
blood, urine) on the scale of tens of millilitres. These samples (or parts thereof) are then
incubated for several hours or days before analysis by a biologist. Although these methods
offer selective and reliable analysis, they are comparatively slow, normally taking between 8
– 24 hours15
; suffer from potential human error, and are not generally field-portable.
A technical report by the International Union of Pure and Applied Chemistry
(IUPAC) defines a biosensor as “a self-contained integrated device, which is capable of
providing specific quantitative or semi-quantitative analytical information using a biological
recognition element”.16
From this definition, a biosensor has three principal components: the
molecular probe, which binds selectively to the target molecule; the transducer, which
9. 3
produces a measurable signal from the binding reaction; and an output system which
amplifies and presents the signal for interpretation.
This experiment investigated the efficacy of using a cantilever array-based device in
detection of gold nanoparticle-oligonucleotide conjugates suspended in buffer. We attach the
molecular probe - a complimentary oligonucleotide strand – with a thiol bond to one or more
cantilever(s) and bathe them in a nanoparticle-oligonucleotide conjugate suspension. The
complimentary oligonucleotides bind (or hybridize, see Section 1.2) and form a “sandwich
assay” between the target modified gold nanoparticles and the cantilever surface. We use an
optical system to monitor the response of the cantilever to vibration by a piezoelectric
actuator. An amplifier and readout system log and display the frequency-domain time-
evolution of the observed eigenfrequency peaks as the array is bathed in a colloid containing
the nanoparticle conjugates.
Thus the cantilever serves the purpose of a chemo-mechanical transducer, as its
eigenfrequencies shift to smaller values as the molecular probe binds with its complimentary
sequence causing a mass increase on the surface of the cantilever. Suitable control and
baseline cantilevers are employed to show the binding-specific eigenfrequency shift.
Tracking this shift allows us to calculate the added mass as the conjugates attached to the
cantilever surface. This technique is a proof of concept for detection of arbitrary DNA
markers.
1.1 Cantilever arrays
Silicon cantilever arrays were fabricated from high-grade single-crystal silicon in the
Micro/Nanomechanics group, IBM Zurich Research Laboratory, Switzerland. The cantilevers
are arranged in a parallel array of eight with a pitch of 250 µm, each of them 500 µm long,
100 µm wide, and 1 µm thick (see Fig. 2). The array is fabricated in one piece using a top-
down lithography process with the eight cantilevers sharing a common support structure.
In dynamic-mode measurements, the cantilevers are vibrated in a sweep across a
frequency interval containing the cantilever’s eigenfrequencies. Typically the higher modes
of cantilever vibration are monitored, as these give a greater signal-to-noise ratio17
. Vibrating
the cantilever at its higher modes also alters the interaction between the cantilever and the
surrounding fluid.17
At low frequencies, there is significant inertial coupling between the
fluid and cantilever resulting in an inertial loading of the beam. This inertial load is called
virtual mass and the cantilever must displace this virtual mass due to the density and
viscosity of the surrounding medium.
10. 4
Figure 2 – Left: An SEM of a typical cantilever array. Right: An optical microscope image of cantilever details with
hair for size comparison. Each cantilever is 500 µm long, 100 µm wide, and 1 µm thick. Similar arrays have been
produced with thicknesses ranging between 500 nm and 7 µm.
Low frequency actuation of the cantilever thus leads to strong damping of the
vibration, reducing the quality factor.18
The virtual mass that must be displaced at
fundamental frequency is about 40 times the cantilever’s mass, and this number drops to
about 10 times the cantilever’s mass at mode 16.17
1.2 DNA basics
Deoxyribonucleic acid (DNA) is an information-carrying molecule naturally
occurring in biological systems. DNA typically refers to the double stranded form of the
molecule (unless otherwise stated) and is composed of two polymer chains of nucleotides,
held together by a backbone of sugars and phosphate groups. These chains form an
antiparallel double helix. Each nucleotide is composed of a five-carbon sugar (2’-
deoxyribose) and one of four types of nucleobase; the pyrimadines Cytosine (C) and
Thymine (T), and the purine derivatives Adenine (A) and Guanine (G). Across the double
helix, bonds are formed only between adenine-thymine pairs (A-T), or guanine-cytosine pairs
(G-C). The convention of naming the carbon atoms in the deoxyribose sugar ring is to
number them 1 to 5. This gives rise to directionality in a nucleotide sequence. The 5’-end (or
five prime) refers to the end of the nucleotide strand which has the fifth carbon in the sugar
ring of the deoxyribose at its terminus. Likewise for the 3’-end (or three prime).
11. 5
An oligonucleotide consists of a single polymer chain of nucleobases without any
cross-bonding to a second chain of bases. This oligonucleotide has a complimentary sequence
with which it can cross-link and form a stable covalent bond, forming a DNA double helix.
This sequence specific bonding is called DNA hybridization and is relied on heavily in this
experiment as the process by which only the target oligonucleotide is bound.
In this experiment a 12 base-pair long oligonucleotide sequence is attached onto the
gold nanoparticle surface while the complimentary sequence is immobilized on the cantilever
surface. Upon placing the oligonucleotide-functionalized cantilever into an analyte solution
containing the complimentary strand, we expect to see a mass change in the cantilever as
DNA hybridization occurs, binding gold nanoparticles to the cantilever via a DNA double
helix strand (Fig. 1).
Figure 3 – The chemical structure of DNA.19
Here we see the two polymer nucleotide chains composed of Adenine
(green), Thymine (purple), Guanine (blue), and Cytosine (red). These nucleotides are held together by a phosphate-
deoxyribose sugar backbone. Also, visible is the 5’end and 3’ end naming convention, arising from the fifth and third
carbon (respectively) in the sugar ring of the deoxyribose terminus. Hybridization refers to the formation of the
dotted bonds between the two chains.
1.3 Device design
The apparatus used can be split up into three sections: the delivery mechanism, the
analysis & monitoring assembly, and the output system. The delivery mechanism consists of
a valve controlled by the computer. This valve allows pressure from the lab’s compressed air
12. 6
Figure 4 – Schematic Diagram of the device
supply to push liquid from the reservoir through a 75 µl internal volume circuit. This circuit
can either bypass, or be brought into confluence with, the analyte storage loop which is
comprised of a 100 µl capacity tube, pre-loaded with a volume of analyte solution that is
experiment dependant. Flow enhances the kinetics of attachment and results in a greater level
of binding.12
For practical applications a small volume would be used in a continuous flow
closed-loop circuit. However, for this experiment a stop-flow sequence was used (section
2.5). This means the reaction rate is diffusion-limited and thus slower. The advantage,
however is that we observe a greater proportion of oligonucleotides binding to the cantilever
surface. The circuit is fed into the thermally isolated chamber and into the sensor flow cell
where it is sealed with an O-ring. Liquid is passed through this open-loop circuit past the
cantilever array before finally dropping into a waste container.
The analysis and monitoring assembly includes the optical system, the piezoelectric
actuator, and the internal thermocouples. All are housed inside a thermally controlled
chamber. The thermocouples feed into the computer and are used to ascertain when the
chamber’s temperature has equilibrated. A third and less vital thermocouple is used outside
the chamber to log the ambient lab temperature. The sensor flow cell is maintained at 23 °C ±
0.1 °C, although this is experiment dependant. The piezoelectric actuator is placed in close
proximity to the sensor flow cell and so is vibrationally coupled with the cantilever array. It is
controlled by the computer and is vibrated in a sweep across a frequency range as specified
13. 7
Figure 5 - A section through the sensor flow cell housing. The piezoelectric actuator is mounted below the array
beneath a 200 µm thick membrane. The volume of the sensor flow cell is 6 µl. Note the 45° angle formed between the
plane of the array and the glass cover faces, allowing the laser light to pass orthogonally through the glass-liquid
interfaces.
by the computer. The optical system passes monochromatic 632.99 nm light from an external
Newfocus AlGaInP laser through optical fiber into the chamber. Inside the chamber the beam
is passed through a collimator and focused using a lens. These components are mounted on a
computer-controlled motorized stage, which is itself mounted on a manual 3-axis stage. The
light is focused near the apex of the cantilever by manually adjusting this 3-axis optical stage.
The diameter of the laser spot is approximately 12 µm. Fine-tuning of the laser spot position
is carried out by vibrating the cantilever and adjusting the position of the beam along the
cantilever using the computer link to the automatic optical stage until the largest amplitude of
vibration is measured. The optimum position for the beam has the spot focused on a node
rather than an anti-node. At a node the deflected beam shows a maximum variation in the
angle of the reflected beam.
The automatic stage, once calibrated, also acts to move the laser spot from one
cantilever to the next. Using a single laser that is translated using an automatic stage differs
from past cantilever array experiments which typically use an array of Vertical-Cavity
Surface-Emitting Lasers (VCSEL).20
From the cantilever, the reflected light impinges on a Position Sensitive Detector
(PSD). The resulting photocurrents are amplified and stored digitally with time and frequency
14. 8
information on the computer. The amplifier signal is relayed to the computer and collected by
a program operated in the LabView environment. This data is then exported for model-fitting
and analysis in NOSEtools and Origin. A readout sequence can be carried out for all eight
cantilevers in approximately 25 seconds.
Changes in refractive index of the liquid could induce differences in angular
deflection. The refractive index will change as buffer and differing oligonucleotide solutions
are flowed through the chamber. This problem is more relevant for static-mode experiments
rather than dynamic mode, where in the former an artificial bending signal will be observed,
in the latter one would at most see a non-frequency specific change in the amplitude of
vibration. This danger is eschewed by having the beam enter and leave the sensor flow cell
via a glass cover positioned such that the beam is orthogonal to both the liquid-glass and
glass-air interfaces.
Figure 6 - (a) Beam translation at antinode, (b) beam deflection at node. When the laser spot is focused on the
antinode, though this part of the cantilever shows the highest amplitude of vibration, we see the weakest deflection in
the reflected beam. The optimum position is at an antinode since the motion of the cantilever results in an angular
gyration of the reflected beam rather than a translational gyration as at the antinode.
1.4 Oligonucleotide-nanoparticle conjugates
The oligonucleotides chosen for the experiment were two different 12 base-pair long
sequences (known as a 12-mer) from the Bio B Biotin synthase gene (EMBL accession
number: J04423) bought from Microsynth. The oligonucleotides are thiolated at the 5’ end to
facilitate binding with gold-coated surfaces. Two surface-bound molecular probes were used
- Bio B2 Compliment (Bio B2c) and Bio B3 Compliment (Bio B3c) - which were attached
onto the upper and lower surfaces of the cantilever. The target sequences (Bio B2 and Bio B3
respectively) were attached in separate solutions onto 50 nm Gold nanoparticles forming a
self-assembled monolayer around the particles. During the experiment, Bio B2 conjugates
will bind to cantilevers coated with the Bio B2c oligonucleotide strands. This results in a
15. 9
measurable mass change and commensurate drop in the resonant frequency of the cantilever.
Similarly for Bio B3 conjugates binding to Bio B3c functionalized cantilevers.21
In practical applications, selectivity of sensor to target is an absolute requirement.
Thus, any investigation into detection efficacy must also assess the selectivity of the sensor to
a specific analyte, to the exclusion of similar analytes that may also be present. Thus, for this
experiment, only two cantilevers in each array are functionalized to bind to a target analyte.
As a control, reference cantilevers were functionalized with either Bio B4 or Unspecific 12.
These sequences act as a control since they share similar properties with the oligonucleotides
that are complimentary to the target analytes; they have comparable molecular weight and
identical monolayer-forming behaviour. However, the controls will not bind to these target
analytes. Thus any frequency shift trend observed in the control cantilevers can be subtracted
from the frequency shift trend of the cantilevers sensitive to the target analytes.
The conjugates were produced such that there was a layer of many oligonucleotides
on the surface of each nanoparticle. Producing conjugates in this way allows them to be
easily separated from free, unreacted thiol-oligonucleotides by centrifugation.22
Also, such
layering obviates the need for a protective shell of anionic phosphate ligands, as is necessary
when producing conjugates with a sparse coating of oligonucleotides.22
When bought from Microsynth, the oligonucleotides come with a thiol modification at
the 5’ end. This facilitates binding with Gold as it forms a competing, stronger bond with the
Gold nanoparticles compared to the nanoparticle citrate covering. This thiol group will tend
to form the oligonucleotides into dimers if left unmodified. This dimerization will drastically
reduce the binding efficiency with gold surfaces.23
In order to prevent this, the manufacturer
binds the thiol group with Dithiothreitol (DTT) which serves as a preservative. Diethyl Ether
(DEE) is used to remove the DTT, exposing the thiol group for conjugation.
The protocol followed gave the following equation for the amount of oligonucleotides
needed to conjugate a given volume of nanoparticle colloid.22
[5]
[6]
where An is the surface area of an individual nanoparticle, cn is the concentration of the stock
nanoparticle solution, D0 is the oligonucleotide density on each particle (taken to be 35
pmol/cm2 24
), V is the desired volume of nanoparticle solution to be conjugated, the radius (r)
referred to is the nanoparticle radius determined from DLS analysis.
16. 10
1.5 Plasma cleaning
Plasma cleaning is an effective way to prepare cantilever surfaces before being gold-
coated. The surface to be cleaned is placed in an evacuated chamber (10-3
atm in this
experiment) and rarefied oxygen gas is fed into the chamber. This gas is excited by high
frequency voltages (typically kHz to MHz). Transitions back to lower energy states illicit the
release of a photon, causing the characteristic glow of the plasma.
The plasma’s activated species include O2
+
, O2
-
, O3, O, O+
,O-
, free electrons, and
photons. These photons are in the short-UV range and are good at breaking organic bonds (C-
H,C-C,C=C,C-O,C-N) of surface contaminants, helping break apart high molecular weight
particles. A second cleaning effect is carried out by the activated species, forming H2O, CO,
and CO2 and other low weight hydrocarbons. These molecules have a high vapour pressure
and so are quickly evaporated from the surface. Plasma cleaning exhibits no surface tension
restrictions and can thus clean into corners that a cleaning solution cannot.
1.6 The ζ-potential
If one state of matter (dispersed phase) is finely dispersed in another (the dispersion
medium), then the system is known as a colloid. In this experiment we deal with a colloid of
gold nanoparticles dispersed in water. When nanoparticles are immersed in a liquid, they
develop a net charge at the particle surface, in the case of gold nanoparticles this is a negative
charge25
. This local surface charge attracts oppositely charged ions of the dispersant to the
particle-dispersant interface, inducing a microheterogenous region. This region can be split
up into two layers; an inner layer called the Stern layer, where the ions are strongly bound,
and an outer, more diffuse layer where the ions are less strongly bound. Thus, an electric
double layer is formed around each particle. Between these two layers is a notional plane
known as the slipping plane. On the proximal side of the slipping plane (nearer the particle
surface) the ions move with the particle as it travels in the fluid, whereas the loosely bound
particles on the distal side of the slipping plane do not travel with the particle. The potential
difference between a point in the slipping plane and the bulk fluid is known as the ζ-potential
(or Zeta potential). The ζ-potential is important as it characterizes the stability of the colloid.
Colloids can come out of suspension, a term called flocculation, under certain
circumstances. This is almost entirely dependent on the sum of attractive and repulsive
interparticle forces.25
Colloid theory developed in the 1940s by Derjaguin, Verwey, Landau
and Overbeek; established that colloidal stability is dependent on the sum of these potentials,
where a sufficiently large repulsive force will make the colloid stable.26
If all particles in
17. 11
Figure 7 - Potential difference as a function of distance from particle surface.
suspension have a large negative or positive ζ-potential, they will repel each other due to
Coulomb forces. A typical value of the ζ-potential that characterizes a stable colloid is larger
than +30mV or less than -30mV.The ζ-potential of a particle in a suspension is heavily
dependent on the pH of the dispersion medium. Indeed a ζ-potential value quoted without a
pH value is largely meaningless as any ζ-potential can be brought to 0 depending on the pH
of the medium. For this experiment, a pH of 7 can be assumed unless otherwise stated. When
an electric field is applied across an electrolyte, the charged particles suspended in the
electrolyte are attracted to the electrodes. The particle’s velocity is a function of electric field
strength, the dielectric constant of the dispersion medium, the viscosity of the medium, and
the ζ-potential of the particle. We define the Electrophoretic Mobility UE as:
[1]
where ζ is the ζ-potential, ε is the dielectric constant, η is the viscosity, and (ka) is known as
Henry’s function, k is the Debye-Huckel parameter, and a is the particle radius. Henry’s
function varies between 1 and 1.5, for the case of an aqueous medium with a moderate
electrolyte concentration, we take the Huckel approximation and set (ka)=1.27
We measure
the Electrophoretic Mobility using Laser Doppler Velocimetry and from this we infer the ζ-
potential using Eq. 1.
18. 12
1.7 Dynamic light scattering
Dynamic Light Scattering (DLS) is a technique for determining the size distribution
of particles in a suspension28
, in this case a colloid of gold nanoparticles suspended in water.
When monochromatic laser light passes through the sample, it forms a speckled
pattern on the other side composed of light and dark regions. These regions are not due to
simple occultation of the incoming laser beam by particles. Rather, this pattern is due to
constructive and destructive interference of monochromatic light that has been scattered by
the particles. Due to Brownian motion of the scattering centres, there is a time-dependant
fluctuation of the scattering intensity. A key aspect of the Brownian motion normally
undergone by particles in a suspension, is that larger particles will move more slowly, as
governed by the Stokes-Einstein equation.27
We fit an autocorrelation function to the intensity trace at a particular point in the
speckle refraction pattern. The time in which the autocorrelation of the intensity trace drops
to zero is dependent on the size of the scattering centres. Larger scattering centres will move
more slowly and hence the intensity trace at a given point will show a longer autocorrelation
timescale. We can also infer the variance in sizes of the scatterers. A suspension of particles
of very similar size is referred to as monodisperse while particles with significantly varied
size are referred to as polydisperse. For our experiment, it is advantageous to have a well-
characterized monodisperse nanoparticle colloid. This is important for quantifying the
attached mass on the cantilever and also aids in the calculation of the amount of
oligonucleotide required for conjugation, as this depends on the radius of the nanoparticles
(see Eq. 6).27
A consequence of this method of calculating the size distribution is that it produces a
scattering intensity distribution rather than a number or volume distribution. To illustrate the
difference, consider a suspension of equal numbers of two sizes of particles: 5 nm and 50 nm.
A number distribution would produce a curve as shown in Fig. 8(a), the area under each
curve is equal as there are equal numbers of each size of particle. A volume distribution
would produce a curve like that shown in Fig. 8(b), with the area under the peak at 50 nm
being 1000 times larger than under the peak at 5 nm. This is due to the fact that the volume
of a 50 nm particle is 1000 times larger than for a 5 nm particle. Finally, an intensity
distribution will produce a curve like that shown in Fig. 8(c). Here, the area under the peak at
50 nm is 1,000,000 times larger than the area under the peak at 5 nm. This is because the
19. 13
Figure 8 – Number, volume, and intensity distributions for the same suspension containing equal numbers of 5 nm
and 50 nm particles. Since scattering intensity is proportional to the 6th
power of the particle diameter29
, even slightly
larger particle species can dwarf smaller particles in a scattering intensity trace. (Diagram adapted from Zetasizer
User Manual27
)
scattering intensity of a particle is proportional to the 6th
power of its diameter, according to
Rayleigh’s approximation.27,29
1.8 Spectrophotometry
In our experiment it is important to measure the concentration of oligonucleotides in
various solutions. These measurements will later be used to calculate the amounts of
oligonucleotides required to form conjugates with a given volume of nanoparticle solution,
and also to subsequently confirm binding of the two. In our spectrophotometer, light of 260
nm wavelength from a pulsed Xenon flash lamp is passed through a liquid sample.30
The
resultant transmitted light is collected along a linear CCD. Beer’s law states that the
Absorbance (A) of a sample is given by30,31
:
[2]
where A is equal to the product of the extinction coefficient (ελ) at wavelength λ, the molar
concentration (c) and the path length (l). Thus if we know the path length of the sample and
its extinction coefficient, we can determine the concentration of absorbers. The extinction
coefficient for a short single-stranded oligonucleotide is dependent on the length and base
composition and can easily be calculated (see Section 2.2)
1.9 Q Factor
In sensing applications of micro-scale cantilevers, a fundamental limit of the
sensitivity is imposed by thermomechanical noise, representing the mechanical analogue of
20. 14
Johnson noise.32
The quality factor (or Q-factor) is one of several ways of characterizing the
bandwidth of a resonator relative to its centre frequency. The narrower the bandwidth, the
lower the noise levels when tracking the frequency response of the cantilever, and so the
more accurate the mass uptake measurements. Thus, much of the design of this experiment
seeks to maximize the Q-factor. The Q-factor parameter is given by18
:
[3]
where is the resonance frequency of the mode and is the full-width half-maximum
(FWHM) of the peak in the frequency domain. The total Q-factor (Q) has several
contributing Q-factors due to internal thermoelastic dissipative loss (QT), loss to the chip
substrate through the cantilever support known as clamping loss (QC), surface effects (QS),
and viscous/acoustic loss to the surrounding medium (QV).18
[4]
Models of the frequency response of the cantilever beam - which assume an
incompressible viscous fluid - indicate that the Q-factor of the resonance peaks increases
commensurately without limit as the mode number increases.33
Realistically, the increase in
Q-factor is not unbounded and a paper by Eysden and Sader – which takes into account fluid
compressibility - predicts a “coincidence point” beyond which the generation of acoustic
waves dramatically reduces the Q-factor.34
This coincidence point marks the point where
acoustic waves are generated, inducing significant loss in the form of sound waves. The
Coincidence point is dependent on the geometry of the cantilever, specifically the length to
thickness ratio.35
However, the same paper concludes that when operated in liquids, this
coincidence point will occur at modes too high to have a significant effect.34
1.10 Mass-frequency shift relation
Models of cantilever vibrational dynamics in liquid previously developed for
understanding their behaviour in AFM are well documented. In a vacuum, the equation of
motion for a cantilever is given by:36
[5]
where E is Young’s Modulus, I is the moment of inertia (together EI is the flexural rigidity),
u(x,t) is the deflection of the cantilever surface as a function of time (t) and position along the
cantilever (x), C0 is the coefficient of intrinsic damping per unit length which describes the
internal dissipative loss, L is the length of the cantilever, mc is the mass of the cantilever.
21. 15
Since in this experiment the cantilever is vibrating in liquid, two effects must be taken
into account. Firstly, the effect of the virtual mass mv of the inertially coupled liquid must be
added to the above equation. Secondly, an additional dissipative force per unit length that is
proportional to the velocity must also be included. The commoving mass produces an
additional inertial force (gi) given by:
[6]
where mv is proportional to the displaced mass of the liquid (md). This displaced liquid mass
is itself proportional to the cantilever volume by:
[7]
where p is a coefficient equal to 1 for an ideal fluid, ρ is the density of the fluid, and Vc is the
volume of the cantilever. Note that the added virtual mass becomes asymptotically smaller
for higher modes.17
Since we are not using an ideal fluid, the additional dissipative force (gv)
per unit length is given by:
[8]
where Cv is the dissipation coefficient. Combining these additional inertial (gi) and
dissipative (gv) forces, as well as a periodic driving force F(x,t) provided by the piezoelectric
actuator, we can expand Eq. 5 to get:
[9]
In order to solve this equation, we need to ascertain the virtual mass mv and damping force gv.
Thus, we need to determine the coefficients p and Cv For the deflection of the cantilever u(t)
we use:
[10]
The resonance frequencies for the nth
mode are the complex solutions of Eq. 5, which are
given by:
[11]
The term is the fundamental eigenfrequency in vacuum for a cantilever with its mass
concentrated at one point, and in the absence of damping. The αn terms are related to the
different eigenvalues of the modes and are the nth
positive root of the equation
, [α1=1, α2=1, α3=1,..., αn=π(n-0.5)] The damping factor γ in Eq. 11 is
given by:
22. 16
[12]
A rectangular cantilever with distributed mass has, without taking into account damping,
eigenfrequencies given by:
[13]
The frequency for which the amplitude of the response of the nth
harmonic is maximized, as a
function of driving force frequency, is given by:
[14]
We can see heuristically from the above equation that the actual resonance frequency is
shifted to lower values from by the damping effect of the liquid. Thus it is important to
make a distinction between the eigenfrequency which does not take account of damping,
and the predicted resonance frequency which is what is predicted as the observed
frequency as it does take account of damping.
The total mass change due to analyte binding (∆m) in this experiment is assumed to
be uniformly distributed on the cantilever surface. Thus the total mass that has to be
accelerated is given by:
[15]
Thus we simply modify Eq. 13 with the mass change to get:
[16]
Since the mass change is likely to be in the nanogram range, (i.e. ) we can
make the approximation:
[17]
We can calculate the attached mass ∆m from the frequency shift ∆f:
[18]
Where . We can also define the sensitivity S of the cantilever in terms of
frequency shift per unit attached mass:
[19]
From this we can see that the mass sensitivity increases with the order of the harmonic of the
cantilever vibration.
23. 17
Chapter 2 Experimental Method
2.1 Cantilever functionalization
2.1.1 HF treatment
It was noticed under optical microscopy that many of the cantilevers showed not-
insignificant levels of heavy metal contamination on arrival from the manufacturer (IBM).
Additionally, due to mishandling an entire batch of cantilever arrays was damaged when the
wafer came loose in transit. Sheer stress on the arrays broke many of them and resulted in the
rest being covered in Silicon Dioxide (SiO2) dust. These contaminants can interfere with
functionalization and operation of the arrays. The damaged arrays represent a significant
investment (approximately €8000) and so it is desirable to find a protocol for salvaging
them.
It is possible that if the SiO2 layer underneath these contaminants could be removed,
the contaminants too would be removed. Hydrofluoric Acid (HF) can be used to etch away
the top layer of SiO2. Thus, immersing the arrays in HF could make them useable again. HF
is extremely corrosive however, with moderate reactivity towards metals and high reactivity
towards glass. Polytetrafluotoethylene (PTFE) is resistant to corrosion by HF, although it is
semi-permeable to it.31
Thus a PTFE holder was designed to facilitate immersion of several
arrays simultaneously. It featured a stand designed to accept six arrays at a time and a
fastening clamp which is held against the arrays and tightened with a separate rod. This
fastening arrangement prevents sheer forces on the arrays. The design also called for a long
PTFE rod for lowering the holder into the HF chamber, the rod is threaded to fix the fastener
to the holder.
24. 18
Figure 9 – Top left: array holder, the holder features grooves shaped to accept the arrays, six aligning slots mate with
protrusions on the fastener upon tightening. Top right: the fastener (inverted) has protrusions which hold arrays
upon fastening. Bottom left: 3D rendering of the holder before fabrication. Bottom right: finished PTFE holder for
hydrofluoric acid treatment of cantilever arrays.
2.1.2 Temescal coating
A Temescal FC2000 Bell Jar electron-beam evaporative deposition System is used to
coat the arrays first with a 2 nm layer of titanium, then with a 20 nm layer of gold. The
titanium serves as an adhesion layer for the gold18
since it has an intermediate crystal
structure between silicon and gold. The purpose of the gold is two-fold. Firstly, it is necessary
to enable thiol-modified oligonucleotides to bind and form a monolayer on the cantilever
surface. Secondly, the gold layer also provides a reflective surface which improves the signal
to noise ratio in the reflected beam signal at the PSD. Both sides of the cantilever were thus
coated. Both metals were evaporated using electron beam deposition at a rate 0.02 nm/s for
Titanium and 0.05 nm/s for Gold.
25. 19
2.1.3 Plasma cleaning
Cantilever arrays were rinsed in nanopure water, dried with nitrogen, rinsed in ethanol
and then dried again. The arrays were then mounted on a metal stage and placed in a plasma
cleaner under clean room conditions. After 5 minutes of exposure to 0.3mbar oxygen plasma,
the arrays were removed and placed in ethanol to passivate their now-reactive outer layer.
2.1.4 Incubation
Figure 10 – Left: A cantilever positioned for insertion into incubation capillaries. Right: Cantilevers inserted into
capillary tubes. Note that capillary tube colour is due to a dye added to illustrate the cantilever functionalization
process, this dye is not present in the experiment.
The cantilever arrays were placed in a UV cleanser and then put in ethanol to
passivate them. Next, the arrays are placed on the functionalization stage, adjacent to eight
glass microcapillaries of internal diameter 180 µm and external diameter 250 µm (King
Precision Glass Inc.). With one cantilever inserted into the end of each capillary. Using a
micropipette, the solution containing the molecular probe - to be attached onto the cantilever
surface – is introduced into a larger reservoir capillary at the distal end of the capillary.
Capillary action draws the solution along the capillary to the proximal end, thus bathing the
cantilever arm in the solution. During this incubation period, the thiol-modified
oligonucleotides form a self-assembled monolayer on the cantilever surfaces. The larger
reservoir capillary serves as a physically more manageable target for manual loading of
solution and also counteracts evaporative loss of the solution at the proximal end.
Table 1 – Cantilever array functionalization pattern
Cantilever number Functionalization
1,2 Bare gold
3,4 Bio B4 compliment (Bio B4c)
5,6 Bio B3 compliment (Bio B3c)
7,8 Bio B2 compliment (Bio B2c)
26. 20
A typical functionalization pattern is shown in Table 1. The cantilever is allowed to
incubate for 20 minutes, allowing the probe molecules to self-assemble into a monolayer on
the cantilever’s upper and lower surfaces. The arrays are then placed in 10 mM sodium
phosphate buffer in a refrigerator.
2.2 Spectrophotometry
All oligonucleotide solutions bought from Microsynth were analysed using a
NanoDrop 1000 Spectrophotometer before being used for conjugation. The procedure
involved first calibrating the spectrophotometer before each measurement using nanopure
water. Samples on the scale of 4µl of each solution were used. Using a nearest-neighbour
model from Tataurov, You, and Owczarzy [2008] we can calculate ελ=260nm for arbitrary
sequences, and we arrive at the values in Table 2.38
2.3 Oligonucleotide-nanoparticle conjugation
Table 2 - Oligonucleotide sequences used in this experiment. The ‘SH’
denotes the thiol group bound to the 5’ end of each nucleotide species.
Name Sequence Calculated extinction coefficient at 260 nm
(L mol−1
cm−1
)
Bio B2 SH-5’-TGC TGT TTG AAG-3’ 113500
Bio B2 Compliment SH-5’-CTT CAA ACA GCA-3’ 117700
Bio B3 SH-5’-CCG GAA GAT TGC-3’ 116200
Bio B3 Compliment SH-5’-GCA ATC TTC CGG-3’ 109600
Unspecific 12 SH-5’-ACA CAC ACA CAC-3’ 119200
Bio B4 SH-5’-GGA AGC CGA GCG-3’ 120800
The oligonucleotides are mixed and agitated with DEE to remove the DTT from the
thiol group. DEE and the water are immiscible and so the DEE and oligonucleotide
suspension separate within seconds with a visible meniscus. A micropipette is used to remove
the DEE and the process is repeated 6 times. To completely remove the DEE, the
oligonucleotide solution is placed in a SpeedVac concentrator for 5 minutes. The
oligonucleotides are then ready for conjugation.
The conjugation protocol that was followed necessitated Dynamic Light Scattering
analysis of the nanoparticles to ascertain their mean diameter and the extent of polydispersity,
prior to conjugation. A Malvern Zetasizer Nano ZS dynamic light scattering apparatus was
used. After carrying out the DLS measurement of the mean diameter, we carry out
27. 21
spectrophotometric analysis of the oligonucleotide solution to determine an exact figure for
the molar concentration of the oligonucleotides (Table 3.5). We use these values to calculate
the amount of oligonucleotide solution needed to conjugate the particles according to Eq. 5.22
An example calculation of oligonucleotide necessary for conjugation is as follows:
The specification sheet for our gold nanoparticles indicates a value of 7.473 108
or
equivalently 4.49 1010
nanoparticles per ml. Using Eq. 5 we calculate the required amount of
oligonucleotides to conjugate 1 ml of nanoparticles:
[20]
Adding the 50% molar excess as recommended by the protocol, we obtain a value of 0.1855
nmol needed to ensure good coverage. Using the oligonucleotide concentration values from
spectroscopic analysis, we can calculate the volume of oligonucleotide solution needed. We
produced separate solutions of Bio B2-functionalized nanoparticles, and Bio B3-
functionalized nanoparticles. Both our solutions were at 100 µM concentration, thus we
needed 1.855 µl of the solution to add to the 1 ml of gold nanoparticles.
The mixture is placed in a glass vial which is covered in tin foil and agitated on a
linear shaker at ~1 Hz for 16 hours. The tin foil serves to prevent exposure to light which
hinders the reaction.24
The mixture is then brought to a 10 mM Sodium Phosphate concentration which acts
as a pH 7 buffer. The addition of Sodium Phosphate buffer is split into 5 smaller additions
and gradually added over 5 hours as is recommended for nanoparticles larger than 20 nm.24 22
The sodium phosphate serves as a pH7 buffer to facilitate DNA binding.
The solution was then centrifuged at 4000rpm for 15 mins in low-adhesion Eppendorf
tubes wherein they form a crimson oil of nanoparticles beneath a clear supernatant of excess
oligonucleotide in solution. The supernatant was removed and retained for analysis. This is
done to remove the free oligonucleotides from the suspension which would otherwise
hybridize with the cantilever-bound complimentary strands, thus preventing those sites from
binding with the nanoparticle-bound oligonucleotides. The nanoparticle oil was then
resuspended in the same volume of identical molar concentration of 10 mM sodium
phosphate buffer. This solution was then centrifuged again and the process was repeated 6
times, with the supernatant retained each time.
The final solution should be virtually devoid of free oligonucleotides at this point.
Spectrophotometric analysis is then carried out on the final solution and each of the
supernatants. This data is used to quantitate the successful binding of the oligonucleotides to
28. 22
the nanoparticles. A drop in the total number of free oligonucleotides in the supernatant
indicates successful conjugation with the gold nanoparticles.
2.4 Dip test
This experiment relies on the successful conjugation of the nanoparticles with the
oligonucleotides. Running the experiment involves a significant time investment. If the
procedure fails to produce an obvious resonance frequency shift, it is important to know
whether this is due to a measurement error in the device or a failure to conjugate the
nanoparticles and/or bind them to the cantilever surface. To test this, we functionalized
several cantilevers using the standard protocol outlined in Section 2.1.4. Rather than attempt
to use a frequency shift measurement in the full apparatus to detect binding, we bathed the
functionalized arrays in low-adhesion Eppendorf tubes, each containing one of several
solutions (either Bio B2, Bio B3, or bare gold nanoparticles) and then imaged them under a
Scanning Electron Microscope (SEM). This is known as a preliminary “dip test”.
2.5 Dynamic mode measurement
The sensor flow cell and supplying circuit were flushed with ethanol at a rate of 225
µl/minute for ~90 minutes. The sensor flow cell was then flushed with nanopure water at the
same rate for ~90 minutes. For all subsequent solutions, a flow rate of 18.2 µl/min was used,
corresponding to a bulk velocity in the circuit of 1.2 m/s . As a cleaning process, the circuit is
filled with 10 mM Sodium Phosphate pH7 buffer for 2.5 mins and then left static for 42.5
minutes. The Bio B2 nanoparticle solution was then injected into the analyte storage loop and
flowed into the sensor flow cell at 18.2 µl/min for 2.5 mins after which the flow was stopped
and left static for 42.5 mins. Following this 10 mM Sodium Phosphate was flowed through
the device at 18.2 µl/min for 10 mins and the flow was left static for 30 mins. The Bio B3
nanoparticle solution was then injected into the storage loop and flowed through the sensor
flow cell at 18.2 µl/min for 2.5 mins after which the flow was left static for 42.5 mins. Finally
10 mM Sodium phosphate buffer was again flowed through the device at 18.2 µl/min for 10
mins, followed by a final 30mins of static flow.
2.6 Data analysis
All data analysis was carried out using data analysis software NOSEtools. This
software runs in the IGOR Pro environment. The model used in this software is described in
Braun et al. [2005]36
and is outlined in Section 1.10.
29. 23
The 7th
and 8th
resonant modes were monitored. 1000 data points in the frequency
interval 120 kHz – 270 kHz were taken giving a frequency resolution of ~150 Hz. Each
frequency is excited for 1ms and the response rate was sampled at a rate of 107
samples per
second. The peaks were fitted with a amplitude spectrum of a simple harmonic oscillator, and
the time evolution of the centre frequency of the peaks was used to calculate mass uptake.
The peak centre frequency ( ) and width ( ) were taken from the fit and used to calculate
the quality factor for each peak ( ). The standard error was also calculated using
the statistics function in OriginPro 8.
The data were baseline corrected by calculating the overall linear drift of the
cantilever resonance frequencies over the baseline period (the initial 45 mins of the
experiment, see Fig. 14a). This was done for each cantilever individually. The resonant
frequency trace for each cantilever was saved in a time-stamped file and analysed in
post-processing using NOSEtools. The data was then normalized such that only relative
frequency shifts are apparent (Fig. 14b). A median filter (box size 7) was applied to the data
to reduce noise. Finally the mass uptake was calculated from this frequency trace by fitting
each frequency spectra with the model outlined below. Plots were made of bound mass vs.
time to determine the binding behaviour during the experiment.
Figure 11 – The amplitude spectrum for a simple harmonic oscillator, fitted to the 7th
and 8th
resonant modes
30. 24
Chapter 3 Results & Discussion
3.1 Dip test verification of compliment-specific binding
Figure 12 – A typical SEM indicating a positive result in a dip-test. The cantilever shows bound nanoparticles of
mean diameter 50 nm. The visible halo surrounding this cluster is due to local charge concentration. Note that this
image is from an earlier run of the experiment which used the same equipment and fabrication techniques. This
particular image shows non-specific binding of bare gold nanoparticles to an unfunctionalized cantilever.
A typical SEM image showing successful binding will exhibit randomly distributed
particles with a mean diameter of approximately 50 nm (Fig. 13), while the reference
cantilever will not. The SEM images of our cantilever showed low levels of non-specific
binding of nanoparticles to the upper surfaces of all cantilevers. There was no obvious target
specific binding of the nanoparticles, and no apparent correlation between specificity of a
cantilever’s functionalization and the observed binding to that cantilever.
31. 25
3.2 Dynamic-mode measurements
Our data shows no discernable specificity to the mass uptake during the course of the
experiment. This suggests non-specific binding of Bio B2 nanoparticle conjugates to all
cantilevers, followed by a universal drop in mass-uptake during the period that buffer was
flowed through the cell (90- 95 mins) following the Bio B2 stop-flow period. This is possibly
due to a rinsing effect of the buffer, removing loosely bound nanoparticles. Following this we
see a common mass uptake across all cantilevers during the end of the buffer flow-through
period (95 – 100 mins). The trace from this period (90 – 100 mins) could be an artefact of the
change in flow conditions in the device. Following this, during the stop-flow buffer period
(100 – 130 mins), we see common drift across all cantilevers which seems to continue at the
same rate during the introduction of the Bio B3 functionalized nanoparticles (130 – 175 mins)
and the period of buffer flow-through (175 - 215 mins).
Figure 13 – Left: Non baseline-corrected data. Right: Baseline-corrected data. The hatched regions indicate the
period of liquid flow through the sensor flow cell. The unhatched regions indicate the stop-flow period during which
no liquid flowed through the sensor flow cell. The red region indicates the period during which Bio B2 conjugates
were present in the bulk liquid in the cell, and similarly the blue region indicates the period during which Bio B3
conjugates were present in the bulk liquid. The white regions indicate the period during which 10 mM sodium
phosphate buffer was flowed through the cell. The traces of each of the compliment-functionalized cantilevers are
shown, with their respective functionalizations indicated in the legend. The trace of mass uptake has been
normalized, to show relative mass uptake; and baseline-corrected, to disregard drift due to extraneous factors
(temperature drift, loosening of the clamp against the array due to vibration, etc.).
(a) (b)
32. 26
3.3 Dynamic light scattering: nanoparticle size distribution
Functionalization Mean Diameter
(nm)
FWHM (nm)
None (Bare gold) 50.52 22.02
Bio B2 57.83 30.53
Bio B3 65.19 45.31
Figure 15 – Dynamic Light Scattering measurements for unfunctionalized, Bio B2-functionalized, and Bio B3-
functionalized nanoparticles.
The DLS results are shown in Fig. 15. The nanoparticles bought from BBI Life
Sciences were found to be sufficiently monodisperse, having a mean diameter of 50.52 nm
with a full-width half-maximum (FWHM) of the intensity trace of 22.02 nm. The Bio B2
functionalized nanoparticles were found to have a mean diameter of 57.83 nm, which
conforms to our expectation of the size increase. Intuitively, we would expect the diameter of
the nanoparticles to increase by double the length of the attached oligonucleotides. The
oligonucleotides have a length of approximately ~4 nm, and so the observed diameter
increases (after conjugation) of 7.8 nm and 15.2 nm are reasonable.
However, the intensity trace shows a leg on the left hand side for both conjugated
nanoparticles, indicating significant scattering around the 6-8 nm mark.
0
5
10
15
20
1 10 100
Intensity (%)
Radius (nm)
Bare gold nanoparticles
0
5
10
15
1 10 100
Intensity (%)
Radius (nm)
Bio B2 Conjugates
0
5
10
15
1 10 100
Intensity (%)
Radius(nm)
Bio B3 Conjugates
33. 27
3.4 ζ-potential measurements
Functionalization ζ-potential (mV)
None (Bare gold) -0.535
Bio B2 -0.393
Bio B3 0.488
Figure 16 – ζ-Potential measurements for unfunctionalized, Bio B2-functionalized, and Bio B3-functionalized
nanoparticles.
The ζ-potential measurements are shown in Fig. 16. The results gave nonsensical
values for the ζ-potential of all particle solutions. We would expect to see a value of -30mV
or less, since gold exhibits a negative surface charge25
and the colloid is empirically observed
to be stable. The ζ-potential appears to vary around 0mV. This value is not possible given the
observed stability of the colloids.
3.5 Spectrophotometry
Spectrophotometric analysis of the oligonucleotide solutions allowed us to calculate
the 10mm absorbance of the oligonucleotide solutions (Fig. 17). The calculated
oligonucleotide concentration was 258.6 ng/µl for the Bio B2 solution, and 156.2 ng/µl for
the Bio B3 solution. Dividing these values by the molecular weight of each oligonucleotide
species, we can calculate the molar concentration of their respective solutions.
0
50000
100000
-50 0 50
Total
Counts
ζ-potential (mV)
Bare gold nanoparticles
0
50000
100000
-50 0 50
Total
Counts
ζ-potential (mV)
Bio B2 conjugates
0
40000
80000
120000
160000
200000
-50 0 50
Total
Counts
ζ-potential (mV)
Bio B3 conjugates
34. 28
Figure 147 – Spectrophotometry results for Bio B2 and Bio B3 solutions
Analysis of the conjugates solutions yielded oligonucleotide concentrations which were too
low to be measured. Successful conjugation is suggested, however, by the observed diametric
size increase of the nanoparticles after conjugation (Section 3.3).
3.6 Discussion
Selective metallization of the cantilever surface has been shown to significantly
improve the Q-factor of single-crystal silicon cantilevers.40
A 2005 study18
looking at gold-
coated silicon cantilevers, observed a severe degradation in Q factor compared to bare
cantilevers. Their work suggested confining the metalized layer to the tip of the cantilever as
a method of reducing dissipation. To illustrate the potential improvement from selective
metallization, the damping caused by metallization of the hinge accounted for ~60% of the
total damping caused by a full coat.40
The signal to noise ratio in the cantilever response
could possibly be improved by refraining from metalizing the cantilever hinge. This
improvement may be negligible, however, as the viscous damping contribution to the Q-
factor typically dominates at atmospheric pressure.18
We know from Rayleigh’s approximation that the intensity of scattering of a particle
is proportional to the 6th
power of its diameter29
(see Section 1.7). The observed leg on the
intensity traces for the conjugated nanoparticles (Fig. 16) could be an obscured “peak”
indicating a significant presence of 6-8 nm scale scatterers. Since this is size measurement is
consistent with expected oligonucleotide length, this could indicate the presence of free
oligonucleotides that either came loose after conjugation with the nanoparticles, or perhaps
were never removed during the conjugation protocol. Alternatively, the observed small scale
0
1
2
3
4
5
6
7
8
9
220 240 260 280 300 320 340
Absorbance
Wavelength (nm)
10mm Absorbance vs Wavelength
Bio B2 solution
(66.2 µM
258.585 ng/µl)
Bio B3 solution
(40.4 µM
156.19 ng/µl)
35. 29
scattering could be explained as simply arising from contamination. The conjugated
nanoparticles are put through several processes that the bare gold nanoparticles are not,
exposing them to air and passing them to and from different containers, and this could result
in an unknown contaminant. However, the small scale of the apparent contaminant scatterers
(6-8 nm) makes it unlikely that it is due to dust particles (on the order of 500 µm) or even
bacteria (on the order of 500 nm). Thus it is reasonable to assume the observed small-scale
scattering is due to scattering from free oligonucleotides. This could explain the absence of a
clear mass-uptake trend in the dynamic mode cantilever experiment, since free
oligonucleotides would hybridize with their cantilever-bound compliments, thus passivating
potential binding sites for the mass-tagged conjugates.
36. 30
Chapter 4 Conclusion
In the Dynamic mode experiment, we would have expected to see a larger mass-
uptake in the Bio B2c functionalized cantilevers during the period that the cantilever is
bathed in Bio B2 conjugates, followed by a plateau in mass uptake while buffer and then Bio
B3 conjugates were flowed through the sensor flow cell. Concurrently, the Bio B3
functionalized cantilevers should have shown no uptake during the same Bio B2 bathing
period. We then should have seen a larger mass-uptake in the Bio B3c functionalized
cantilevers during the period that the cantilever is bathed in Bio B3 conjugates. We would
expect to see some binding of nanoparticles to reference cantilevers, and slightly less to the
control cantilevers (Bio B4 functionalization) due to a passivation effect of having a mono-
layer of oligonucleotides that would not hybridize with nanoparticle-bound oligonucleotides.
The results did not conform to expectations. We can conclude after baseline correction and
normalization that all cantilevers showed mass binding.
This experiment was unsuccessful in establishing the efficacy of dynamic mode
cantilever array detection of gold nanoparticle-bound oligonucleotides. Though we observed
a negative result for the dynamic mode experiment, we cannot ascertain why this was
unsuccessful. The problem may lie in a failure to conjugate the nanoparticles with the
oligonucleotides, or alternatively the problem may be a failure of the nanoparticles to retain
their oligonucleotide covering. Indeed, due to the wealth of existing research done using
nanosensing cantilevers, we conclude that the observed results are a consequence of the
conjugation protocol followed, rather than the device design itself. We arrive at this
conclusion since our device successfully detected mass-uptake during the course of the
dynamic-mode experiment, (albeit it due to non-specific binding) while exhibiting sensitivity
on the scale of ~4 pg/Hz with very low noise of approximately ±0.5 ng,
It stands as a testament to the precision of the device’s design, that we can detect such
small mass uptake since very few methods allow such a degree of sensitivity. Furthermore,
even fewer technologies allow such sensitivity in a liquid that resembles physiological
environments, and it is this prerequisite which places cantilever arrays in a crucial position in
the field of probing biological processes. Further work is needed to prove that this method is
as promising as many similar methods being investigated in this exciting sub-field of
biophysics.
37. 31
References
1
Joseph Wafula Ndieyira, Moyu Watari, Alejandra Donoso Barrera, Dejian Zhou, Manuel
Vogtli, Matthew Batchelor, Matthew A. Cooper, Torsten Strunz, Mike A. Horton, Chris Abell,
Trevor Rayment, Gabriel Aeppli, and Rachel A. McKendry, Nat Nano 3 (11), 691 (2008).
2
David Maraldo, Fernando U. Garcia, and Raj Mutharasan, Analytical Chemistry 79 (20), 7683
(2007).
3
Sen Xu and Raj Mutharasan, in Environmental Microbiology - Current Technology and Water
Application, edited by Nicholas J. Ashbolt Keya Sen (Caister Academic Press, 2011), pp. 65.
4
Natalia Nugaeva, Karin Y. Gfeller, Natalia Backmann, Marcel Düggelin, Hans Peter Lang,
Hans-Joachim Güntherodt, and Martin Hegner, Microscopy and Microanalysis 13 (01), 13
(2007).
5
J. Zhang, Hans-Peter Lang, F. Huber, A. Bietsch, W. Grange, U. Certa, R. McKendry, H.J.
Guntherodt, Martin Hegner, and Ch. Gerber, Nat Nano 1 (3), 214 (2006).
6
Kathrin Gruber, Tim Horlacher, Riccardo Castelli, Andreas Mader, Peter H. Seeberger, and
Bianca A. Hermann, ACS nano 5 (5), 3670 (2011).
7
Rüdiger Berger, Emmanuel Delamarche, Hans Peter Lang, Christoph Gerber, James K.
Gimzewski, Ernst Meyer, and Hans-Joachim Güntherodt, Science 276 (5321), 2021 (1997).
8
Gossett A. Campbell and Raj Mutharasan, Analytical Sciences 21 (4), 355 (2005).
9
Gossett A.Campbell and Raj Mutharasan, Biosensors and Bioelectronics 21 (9), 1684 (2006).
10
Qing Zhu, Wan Y. Shih, and Wei-Heng Shih, Biosensors and Bioelectronics 22 (12), 3132
(2007).
11
Gossett A.Campbell and Raj Mutharasan, Biosensors and Bioelectronics 23 (7), 1039 (2008).
12
Gossett Campbell, David Maraldo, and Raj Mutharasan, in Nanoscience and Nanotechnology
for Chemical and Biological Defense (American Chemical Society, 2009), Vol. 1016, pp. 25.
13
Nathaniel Rosi and Chad Mirkin, Chemical Reviews 105, 1547 (2005).
14
J. A. Dougan, C. Karlsson, W. E. Smith, and D. Graham, Nucleic acids research 35 (11), 3668
(2007).
15
L. M. Prescott, J. P. Harley, and D.A. Klein, Microbiology, 6 ed. (McGraw-Hill Education,
Boston, 2005).
16
Daniel Thevenot, Klara Toth, Richard Durst, and George Wilson, Biosensors &
Bioelectronics 16, 121 (2001).
17
Murali Krishna Ghatkesar, Thomas Braun, Viola Barwich, Jean-Pierre Ramseyer, Christoph
Gerber, Martin Hegner, and Hans Peter Lang, Applied Physics Letters 92 (4), 043106 (2008).
18
R. Sandberg, K. Mølhave, A. Boisen, and W. Svendsen, J. Micromech. Microeng. 15 (12),
2249 (2005).
19
Madeleine Price Ball, (Wikimedia Commons, 27 March 2007).
20
Bharat Bhushan, Springer Handbook of Nanotechnology, 2 ed. (Springer, 2006).
21
Hans Peter Lang, Martin Hegner, and Christoph Gerber, Materials Today 8 (4), 30 (2005).
22
T. Andrew Taton, in Current Protocols in Nucleic Acid Chemistry (John Wiley & Sons, Inc.,
2001).
23
Sigma-Aldrich, (Sigma-Aldrich, 2011).
24
Linette M. Demers, Chad A. Mirkin, Robert C. Mucic, Robert A. Reynolds, Robert L. Letsinger,
Robert Elghanian, and Garimella Viswanadham, Anal. Chemistry 72 (22), 5535 (2000).
25
Douglas H. Everett, Basic Principles of Colloid Science. (The Royal Society of Chemistry,
1988).
26
B.V. Derjaguin, L.D. Landau, E. J. W. Verwey, and J. T. G. Overbeek, Acta Physiochim. 14
(633) (1941).
38. 32
27
Zetasizer Nano User Manual. (Malvern, Malvern Instruments Ltd., Worcestershire, United
Kingdom, 2009), p.302.
28
Bruce J. Berne, Dynamic light scattering : with applications to chemistry, biology, and physics
/ Bruce J. Berne and Robert Pecora. (Wiley, New York :, 1976).
29
Masuo Hosokawa, Kiyoshi Nogi, Makio Naito, and Toyokazu Yokoyama, (Elsevier Science,
2007), pp. 372.
30
(Thermo Fischer Scientific, Wilmington, U.S.A, 2010).
31
(Ocean optics, 2012).
32
K. Y. Yasumura, T. D. Stowe, E. M. Chow, T. Pfafman, T. W. Kenny, B. C. Stipe, and D. Rugar,
Microelectromechanical Systems, Journal of 9 (1), 117 (2000).
33
John Elie Sader, Journal of Applied Physics 84 (1), 64 (1998).
34
Cornelis A. Van Eysden and John E. Sader, Journal of Applied Physics 106 (9), 094904 (2009).
35
Jason Jensen and Martin Hegner, Journal of Sensors 2012 (2011).
36
Thomas Braun, Viola Barwich, Murali Krishna Ghatkesar, Adriaan H. Bredekamp, Christoph
Gerber, Martin Hegner, and Hans Peter Lang, Physical Review E 72 (3), 031907 (2005).
37
Jean Aigueperse, Paul Mollard, Didier Devilliers, Marius Chemla, Robert Faron, René
Romano, and Jean Pierre Cuer, in Ullmann's Encyclopedia of Industrial Chemistry (Wiley-VCH
Verlag GmbH & Co. KGaA, 2000).
38
Andrey V. Tataurov, Yong You, and Richard Owczarzy, Biophysical Chemistry 133 (1-3), 66
(2008).
39
Thomas Braun, Murali Krishna Ghatkesar, Natalija Backmann, Wilfried Grange, Pascale
Boulanger, Lucienne Letellier, Hans-Peter Lang, Alex Bietsch, Christoph Gerber, and Martin
Hegner, Nat Nano 4 (3), 179 (2009).
40
G. Sosale, K. Das, L. Frechette, and S. Vengallatore, J. Micromech. Microeng. 21 (10) (2011).
