larry o'connell - thesis

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

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.

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.

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.

Contents

Abstract ................................................................................................................................... i

Preface .................................................................................................................................... ii

Acknowledgments ................................................................................................................ iii

Chapter 1 Introduction .......................................................................................................... 1

1.1 Cantilever arrays ......................................................................................................... 3

1.2 DNA basics ................................................................................................................. 4

1.3 Device design .............................................................................................................. 5

1.4 Oligonucleotide-nanoparticle conjugates .................................................................... 8

1.5 Plasma cleaning ......................................................................................................... 10

1.6 The ζ-potential ........................................................................................................... 10

1.7 Dynamic light scattering ........................................................................................... 12

1.8 Spectrophotometry .................................................................................................... 13

1.9 Q Factor ..................................................................................................................... 13

1.10 Mass-frequency shift relation .................................................................................... 14

Chapter 2 Experimental Method........................................................................................ 17

2.1 Cantilever functionalization ...................................................................................... 17

2.1.1 HF treatment ..................................................................................................... 17

2.1.2 Temescal coating ............................................................................................... 18

2.1.3 Plasma cleaning ................................................................................................. 19

2.1.4 Incubation .......................................................................................................... 19


2.3 Oligonucleotide-nanoparticle conjugation ................................................................ 20

2.4 Dip test ...................................................................................................................... 22

2.5 Dynamic mode measurement .................................................................................... 22

2.6 Data analysis ............................................................................................................. 22

Chapter 3 Results & Discussion ........................................................................................ 24

3.1 Dip test verification of compliment-specific binding ............................................... 24

3.2 Dynamic-mode measurements .................................................................................. 25

3.3 Dynamic light scattering: nanoparticle size distribution ........................................... 26

3.4 ζ-potential measurements .......................................................................................... 27


3.6 Discussion ................................................................................................................. 28

Chapter 4 Conclusion ........................................................................................................ 30

References ................................................................................................................................ 31

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, urine

2, tap-

water3, air

4 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, proteomics

6, food engineering

4 and chemistry

7. Dynamic-mode-operated

cantilevers have been shown capable of detecting bacterial pathogens in liquid, notably:

Enterrohemorrhagic Escherechia coli8, Bacillus anthracis

9(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

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

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.

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).

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

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

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

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

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.

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

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.

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

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

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.

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:

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.

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.

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.

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)



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.


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

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

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.

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

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.

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)

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

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.


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

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

Ab

sorb

ance

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)

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.

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.

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).

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).

larry o'connell - thesis

Documents