A MIXED BIOSENSING FILM COMPOSED OF OLIGONUCLEOTIDES … · students: Anthony Tavares, Rhys Crasto,...

A MIXED BIOSENSING FILM COMPOSED OF OLIGONUCLEOTIDES AND POLY(2-HYDROXYETHYL

METHACRYLATE) BRUSHES TO ENHANCE SELECTIVITY FOR DETECTION OF SINGLE

NUCLEOTIDE POLYMORPHISMS

by

April Ka Yee Wong

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy

Graduate Department of Chemistry University of Toronto

© Copyright by April Ka Yee Wong 2010

ii

A MIXED BIOSENSING FILM COMPOSED OF

OLIGONUCLEOTIDES AND POLY(2-HYDROXYETHYL

METHACRYLATE) BRUSHES TO ENHANCE SELECTIVITY

FOR DETECTION OF SINGLE NUCLEOTIDE

POLYMORPHISMS

April Ka Yee Wong

Doctor of Philosophy

Graduate Department of Chemistry University of Toronto

2010

Abstract

This work has explored the capability of a mixed film composed of oligonucleotides and

oligomers to improve the selectivity for the detection of fully complementary oligonucleotide

targets in comparison to partially complementary targets which have one and three base-pair

mismatched sites. The intention was to introduce a “matrix isolation” effect on oligonucleotide

probe molecules by surrounding the probes with oligomers, thereby reducing oligonucleotide-to-

oligonucleotide and/or oligonucleotide-to-surface interactions. This resulted in a more

homogeneous environment for probes, thereby minimizing the dispersity of energetics associated

with formation of double-stranded hybrids. The mixed film was constructed by immobilizing

pre-synthesized oligonucleotides onto a mixed aminosilane layer and then growing the oligomer

portion by surface-initiated atom transfer radical polymerization (ATRP) of 2-hydroxy

methacrylate (PHEMA). The performance of the mixed film was compared to films composed

iii

of only oligonucleotides in a series of hybridization and melt curve experiments. Surface

characterization techniques were used to confirm the growth of the oligomer portion as well as

the presence of both oligonucleotides and oligomer components. Polyatomic bismuth cluster

ions as sources for time-of-flight secondary ion mass spectrometry experiments could detect both

components of the mixed film at a high sensitivity even though the oligomer portion was at least

200-fold in excess.

At the various ionic strengths investigated, the mixed films were found to increase the

selectivity for fully complementary targets over mismatched targets by increasing the sharpness

of melt curves and melting temperature differences (ΔTm) by 2- to 3-fold, and by reducing non-

specific adsorption. This resulted in improved resolution between the melt curves of fully and

partially complementary targets. A fluorescence lifetime investigation of the Cy3 emission

demonstrated that Cy3-labeled oligonucleotide probes experienced a more rigid

microenvironment in the mixed films.

These experiments demonstrated that a mixed film composed of oligonucleotides and

PHEMA can be prepared on silica-based substrates, and that they can improve the selectivity for

SNP discrimination compared to conventional oligonucleotide films.

iv

Acknowledgments I would like to thank Prof. Ulrich J. Krull for being such a positive influence in my life.

He has guided me with enthusiasm and support from the beginning and especially in the end. I

am extremely grateful for all that he has done for me. I wish to also thank Prof. Aaron Wheeler

and Prof. Scott Prosser for being in my advisory committee and giving helpful comments over

the years.

I must give special thanks to Prof. Chris Yip and Dr. Patrick Yang (IBBME, University

of Toronto) for giving me AFM instrument use time. I would also like to express my gratitude to

Dr. Himadri Mandal and Prof. Shirley Teng (University of Waterloo) for collecting the AFM

images. I would also like to thank Dr. Rana Sodhi and Peter Brodersen (Surface Interface

Ontario, University of Toronto) for collecting XPS and TOF-SIMS data and for their expertise.

And finally, I would like to offer many thanks to Dr. Neil Coombs and Ilya Gouverich (Centre

for Nanostructure Imaging, University of Toronto) for giving me SEM training. I would like to

thank Dr. Peter Mitrakos (University of Toronto Mississauga) for assisting me with the operation

of various instruments. I must offer special thanks to Prof. Claudiu Gradinaru and Dr. Denys

Marushchak (University of Toronto Mississauga), for collecting the lifetime and anisotropy data,

which became important pieces of my thesis.

To my colleagues, most notably Taufik Al-Sarraj, Sameer Al-Abdul Wahid, Russ Algar,

Andrew Chan, I-San Chan, Lu Chen, Yevgenia Kratvsova, Melissa Massey, and Kris Wang,

thank you for your friendship and for many years filled with, not only discussions about research,

but laughter, lunches, trips, dinners, and movies. I would like to give special thanks to Russ

Algar for writing the Eslide program, which was a tremendous help for processing the data. I

would like to thank Dr. Neil McKinnon for contributing his expertise on the NMR spectrum of

v

polymers. I am grateful for the extra help and the great ideas provided by former summer

students: Anthony Tavares, Rhys Crasto, Miki Stanikic, and Lori Chong. I must also thank the

administrative staff, Carmen Bryson and Anna Liza Villavelez, for being such great help and for

taking care of most matters of Departmental life.

I would like to thank my parents and my sister for their continuous support over the

years. I am also grateful to my new Adorjan family who has embraced me from the start. Last

but not least, I wish to thank my husband, Mike, for being always there when I needed him. I

have thought that Mike was crazy when he said that he is looking forward to his own Ph.D.

defence. His enthusiasm has eventually brought me more confidence to overcome the last hurdle

of this long journey.

vi

Table of Contents Abstract ........................................................................................................................................... ii

Acknowledgments.......................................................................................................................... iv

Table of Contents........................................................................................................................... vi

Symbols and Abbreviations ............................................................................................................ x

List of Tables ................................................................................................................................ xv

List of Figures ............................................................................................................................. xvii

List of Equations and Chemical Reactions ................................................................................ xxiii

List of Appendices ...................................................................................................................... xxv

Chapter 1......................................................................................................................................... 1

1 Introduction ................................................................................................................................ 1

1.1 Overview............................................................................................................................. 1

1.2 Principles of Nucleic Acid Properties, Structure, and Function ......................................... 5

1.2.1 Deoxyribonucleic Acids (DNA) ............................................................................. 5

1.2.2 Base-pairing ............................................................................................................ 8

1.2.3 Structure of DNA.................................................................................................... 9

1.2.4 Energetics of the Double Helix............................................................................. 10

1.2.5 Denaturation of the Double Helix......................................................................... 12

1.2.6 Effect of pH on DNA Structure ............................................................................ 14

1.2.7 Oligonucleotides as Probes for Recognition of Target Sequences ....................... 15

1.3 DNA Biosensors ............................................................................................................... 16

1.3.1 Optical Biosensors ................................................................................................ 19

1.3.2 Fluorescence ......................................................................................................... 19

1.3.3 Total Internal Reflection and Evanescent Waves ................................................. 23

1.3.4 Microarrays ........................................................................................................... 26

vii

1.3.5 Examples of Optical DNA Biosensors ................................................................. 26

1.4 Strategies for Immobilization of Oligonucleotide Probes ................................................ 28

1.4.1 Adsorption............................................................................................................. 28

1.4.2 Biotin-Avidin Affinity Pair................................................................................... 29

1.4.3 Thiol-Gold Interaction .......................................................................................... 30

1.4.4 Covalent Linkages ................................................................................................ 33

1.5 Atom Transfer Radical Polymerization ............................................................................ 43

1.5.1 Controlled/“Living” Polymerizations ................................................................... 44

1.5.2 Atom Transfer Radical Polymerization ................................................................ 47

1.5.3 Application to Biosensors ..................................................................................... 52

1.6 Contributions of this Thesis .............................................................................................. 54

Chapter 2....................................................................................................................................... 62

2 Experimental ............................................................................................................................ 62

2.1 Materials ........................................................................................................................... 62

2.2 Instrumentation ................................................................................................................. 63

2.3 Procedures......................................................................................................................... 67

2.3.1 Preparation of Silicon, Fused Silica, and Glass Substrates................................... 67

2.3.2 Synthesis of Benzaldehyde (BZ)-Capped APTMS [Benzylidene-(3-trimethoxysilanyl-propyl)-amine]......................................................................... 68

2.3.3 Immobilization of APTMS or BZ-APTMS .......................................................... 68

2.3.4 Immobilization of Heterobifunctional Linker, Sulfo-SMCC................................ 69

2.3.5 Immobilization of SH-SMN Probe ....................................................................... 70

2.3.6 Hydrolysis of Benzylimine ................................................................................... 72

2.3.7 Synthesis of Bromoisobutyryl NHS Ester Initiator .............................................. 73

2.3.8 Immobilization of Bromoisobutyryl NHS Ester onto APTMS-Modified Silicon Wafers....................................................................................................... 74

2.3.9 ATRP of HEMA on Bromoisobutyryl-Immobilized Silicon Wafers ................... 75

viii

2.3.10 Hybridization of Target Oligonucleotides with Probes on Aminosilane Surface or in Mixture with PHEMA on Glass Substrates.................................................. 77

2.3.11 Measurement of Immobilization Efficiency and Hybridization Yield (Chapter 3.2) ........................................................................................................................ 78

2.3.12 Acquisition of Melt Curves from Oligonucleotide Probe Films with and without PHEMA on Glass Surfaces...................................................................... 79

2.3.13 Conjugation of Thiolated-Oligonucleotides to Au Nanoparticles (Chapter 3.3) .. 80

2.3.14 Dissolution of Gold Nanoparticles by KCN and Construction of a Calibration Curve for Calculation of Surface Density of Oligonucleotides on Gold Nanoparticles ........................................................................................................ 81

2.3.15 Hybridization or Adsorption of the Au nps-DNA Conjugates to Fused Silica Surfaces Modified with Oligonucleotide Films or Mixed Films for SEM Analysis (Chapter 3.3) .......................................................................................... 82

Chapter 3....................................................................................................................................... 83

3 Results and Discussion............................................................................................................. 83

3.1 Syntheses of Benzaldehyde-Protected Aminosilane, Initiator, and PHEMA for Assembly of Mixed Film .................................................................................................. 83

3.1.1 Introduction........................................................................................................... 83

3.1.2 Synthesis of Benzaldehyde-Protected APTMS (N-[(1Z)-phenylmethylene]-3-(trimethoxysilyl)propan-1-amine)......................................................................... 84

3.1.3 Synthesis of the Bromoisobutyryl NHS Ester Initiator (1-[(2-bromo-2-methylpropanoyl)oxy]pyrrolidine-2,5-dione)....................................................... 86

3.1.4 NMR Characterization of PHEMA....................................................................... 89

3.1.5 Conclusions........................................................................................................... 91

3.2 Surfaces for Tuning of Oligonucleotide Biosensing Selectivity Based on Surface-Initiated Atom Transfer Radical Polymerization on Glass and Silicon Substrates .......... 92

3.2.1 Abstract ................................................................................................................. 93

3.2.2 Introduction........................................................................................................... 94

3.2.3 Surface Characterization of Each Immobilization Step........................................ 98

3.2.4 Choice of Monomer ............................................................................................ 108

3.2.5 Effect of Benzaldehyde-Capped Aminosilanes on ATRP Rate.......................... 109

ix

3.2.6 Analytical Performance for Biosensor Development ......................................... 110

3.2.7 Conclusions......................................................................................................... 120

3.3 Bin+ Cluster Ion Sources for Investigation of a Covalently-Immobilized Mixed Film

Composed of Oligonucleotides and Poly(2-hydroxyethyl methacrylate) Brushes......... 121

3.3.1 Abstract ............................................................................................................... 122

3.3.2 Introduction......................................................................................................... 122

3.3.3 XPS Characterization of Mixed Film on Glass Surfaces.................................... 124

3.3.4 Estimation of Density of Surface Immobilized Oligonucleotide Probes by Using Au Nanoparticle-Tagged Complementary Oligonucleotide Targets........ 127

3.3.5 ToF-SIMS Characterization of Mixed Films on Glass Surfaces ........................ 133

3.3.6 Conclusions......................................................................................................... 146

3.4 A Mixed film Composed of Oligonucleotides and Poly(2-Hydroxyethyl Methacrylate) Brushes to Enhance Selectivity for Detection of Single Nucleotide Polymorphisms ............................................................................................................... 148

3.4.1 Abstract ............................................................................................................... 149

3.4.2 Introduction......................................................................................................... 149

3.4.3 Selectivity of Various Targets on the Mixed films............................................. 153

3.4.4 Fluorescence Lifetime to Identify Different Microenvironments in Oligonucleotide Films vs. Mixed Films.............................................................. 155

3.4.5 Comparison of Selectivity for SNP Detection between Oligonucleotide and Mixed films......................................................................................................... 160

3.4.6 Conclusions......................................................................................................... 169

Chapter 4..................................................................................................................................... 171

4 Summary ................................................................................................................................ 171

Chapter 5..................................................................................................................................... 174

5 Future Directions.................................................................................................................... 174

References................................................................................................................................... 178

Appendices.................................................................................................................................. 189

x

Symbols and Abbreviations

A Adenine

AFM Atomic Force Microscopy

APTES Aminopropyltriethoxysilane

APTMS Aminopropyltrimethoxysilane

ATRA Atom Transfer Radical Addition

ATRP Atom Transfer Radical Polymerization

b Pathlength of a cuvette

BODIPY Meso-substituted boron-dipyrrin dyes

bpm Base-pair mismatch(es)

bpy 2,2'-bipyridyl

BZ Benzaldehyde

BZ-APTMS Benzaldehyde-protected (3-aminopropyl)trimethoxysilane

C Cytosine or concentration

CDCl3 Deuterated chloroform

cDNA Single-stranded complementary deoxyribonucleic acid

C.I. Confidence Interval

CV Coefficient of Variation

Cy3 Cyanine dye with three carbon atoms in polymethine bridge

DCM Dichloromethane

DMF Dimethylformamide

DMSO Dimethylsulfoxide

DMT-HEG Dimethoxytrityl hexaethylene glycol

DNA Deoxyribonucleic Acid

xi

Δ Phase change of the electric field in ellipsometry

ΔTm Difference in melting temperature

dsDNA Double-Stranded DNA

dT20 Thymidylic acid icosanucleotides

dx Change in temperature for greatest change in fssDNA

E Intensity of the evanescent wave field

E0 Intensity of the electric field

EB Ethidium Bromide

EDC 1-Ethyl-3-(3-Dimethylaminopropyl) Carbodiimide)

EGMEM Ethylene Glycol Methyl Ether Methacrylate

EGMP Ethylene Glycol Methacrylate Phosphate

ε Molar Absorptivity

FC Fully Complementary

Fcc Face-centred cubic

Fmoc Fluorenylmethyloxycarbonyl

FRET Fluorescence Resonance Energy Transfer

fssDNA Fraction of single-stranded DNA

FWHM Full Width Half Maximum

G Guanine

GOPS 3-Glycidyloxypropyltrimethoxysilane

HEG Hexaethylene glycol

HEMA 2-Hydroxyethyl Methacrylate

HOMO Highest Occupied Molecular Orbital

K Instrumental response coefficient

kr Rate of radiative decay

xii

knr Rate of non-radiative decay

I Intensity of a photon

L Ligand in ATRP

λ Wavelength of a photon

LMIG Liquid Metal Ion Gun

LUMO Lowest Unoccupied Molecular Orbital

m/z Mass-to-Charge Ratio

MEHQ Monomethyl Ether Hydroquinone

MeOH Methanol

MEMS Microelectromechanical systems

Mn Number average molecular weight

Mw Weight average molecular weight

NC Non-Complementary targets

NHS N'-Hydroxysuccinimide

NMR Nuclear Magnetic Resonance

NMRP Nitroxide-Mediated Radical Polymerization

nps Nanoparticles

PBS Phosphate-Based Saline

PCR Polymerase Chain Reaction

PEG Poly(Ethylene Glycol)

PHEMA Poly(2-Hydroxyethyl Methacrylate)

Pn• Active species in ATRP

Pn-X Dormant species in ATRP

ppm Parts per million on chemical shifts scale

Ψ Amplitude change of the electric field in ellipsometry

xiii

n Refractive index

Φ Quantum yield

QCM Quartz Crystal Microbalance

RAFT Reversible Addition Fragmentation Chain Transfer

RCA Rolling Circle Amplification or

Surface cleaning method developed by the RCA company

RNA Ribonucleic Acid

SAMs Self-Assembled Monolayers

SARS Severe Acute Respiratory Syndrome

SDS Sodium Dodecyl Sulfate

SE Secondary Electron

SEM Scanning Electron Microscopy

SFRP Stable-Free Radical Polymerization

SHOM Sequence by Hybridization to Oligonucleotide Microchip

Si wafer Silicon Wafer

SIMS Secondary Ion Mass Spectrometry

SMA Spinal Muscular Atrophy

SMN Survival Motor Neuron gene

SNP Single Nucleotide Polymorphism

SPR Surface Plasmon Resonance

ssDNA Single-stranded DNA

SSA Styrenesulfonic acid

Sulfo-SMCC Sulfosuccinimidyl 4-[N-maleimidomethyl]cyclohexane-1-carboxylate

STEM Scanning Transmission Electron Microscopy

T Thymine

xiv

τs Fluorescence lifetime decay of the singlet state

TCEP Tris(2-Carboxyethyl)phosphine hydrochloride

TEM Transmission Electron Microscopy

TEMPO 2,2,6,6-Tetramethyl-1-Piperidinyloxy

θc Critical angle

TIR Total Internal Reflection

TIRF Total Internal Reflection Fluorescence

Tm Melting Temperature

TMS Cl Chlorotrimethylsilane

ToF-SIMS Time-of-Fight Secondary Ion Mass Spectrometry

U Uracil

UHV Ultra-High Vacuum

UV Ultraviolet

XPS X-ray Photoelectron Spectroscopy

xv

List of Tables

Table # Title of Table Page #

1.1 Various types of biosensors based on DNA hybridizations. 18

1.2 Types of static and dynamic fluorescence measurements. 22

1.3 Distinct characteristics of living and controlled polymerizations. 46

1.4 Types of controlled/ “living” radical polymerization. 47

2.1 Oligonucleotide sequences used. 71

3.1 Carbon peak positions of the bromoisobutyryl NHS ester. 89

3.2 Proton peak positions of PHEMA. 91

3.3 Elemental identifications and quantifications of the N 1s spectra. 102

3.4 Densities, immobilization yield and hybridization efficiency of

oligonucleotide probes and targets on oligonucleotide film and

mixed film surfaces.

113

3.5 Tm values of fully complementary and 3 bpm SMN targets on

mixed film and oligonucleotide film immobilized on glass slides.

117

3.6 dx values of fully complementary and 3 bpm SMN targets on

mixed film and oligonucleotide film immobilized on glass slides.

119

3.7 Atomic percentages of nitrogen-containing carbon species unique

to oligonucleotides on Surfaces A to C collected at a take-off

angle of 30° measured by XPS.

126

3.8 List of secondary ion fragments and peak intensities found by

using Bi5+ as a cluster ion primary source in TOF-SIMS for each

140

xvi

type of surface.

3.9 Fluorescence lifetime and steady-state anisotropy values obtained

from intensity time trajectories measured on a confocal

microscope.

156

3.10 Comparison of dx values in all ionic strengths studied. 162

3.11 Comparison of ΔTm values in all ionic strengths studied. 163

xvii

List of Figures

Figure # Title of Figure Page #

1.1 Depiction of a mixed film composed of oligonucleotides and

poly(2-hydroxyethyl methacrylate) brushes immobilized on a

silica-based substrate.

4

1.2 Proposed oligonucleotide conformations in (a) oligonucleotide

films; (b) mixed films.

4

1.3 Proposed interfacial melt curves from (a) oligonucleotide films;

(b) mixed films.

5

1.4 5′- to 3′-end structure of a dinucleotide – a string of 2 nucleotides

– that composes of guanine and adenine on one strand and

cytosine and thymine on the complementary strand.

7

1.5 Chemical structures of nitrogenous bases present in

deoxyribonucleotides and ribonucleotides; (a) purines; (b)

pyrimidines.

7

1.6 Watson-Crick model of the DNA double helix. The ribbon-like

strands represent the phosphate backbone, and the parallel,

horizontal rungs represent the nitrogenous base pairs.

9

1.7 Melt curves of SMN hybrids in 0.5× PBS. 14

1.8 Concept of a biosensor. The binding of the target DNA strands to

the immobilized DNA probes (biological reaction) can be detected

via the fluorescence emission of a dye due to intercalation

(analytical signal).

17

1.9 Electronic transition of absorption and fluorescence. 20

xviii

1.10 The generation of evanescent waves during total internal

reflection.

24

1.11 The ligand-copper structures at reduced and oxidized states in

ATRP.

51

2.1 Synthesis of BZ-APTMS. 68

2.2 Immobilization of 1:1 ratio of APTMS and BZ-APTMS on silica-

based substrates. 69

2.3 Immobilization of sulfo-SMCC onto silica substrates. 70

2.4 Confocal image of a glass slide that was spotted with Sequence 1

and the reference spots (green spots) used Sequence 2.

71

2.5 Immobilization of oligonucleotides onto mixed APTMS and BZ-

APTMS modified silica substrates. 72

2.6 Hydrolysis of remaining BZ-APTMS sites. 73

2.7 Synthesis of the bromoisobutyryl NHS ester initiator. 74

2.8 Immobilization of initiator onto remaining APTMS sites. 75

2.9 Purging the monomer/copper catalyst mixture with Ar gas. 76

2.10 ATRP of HEMA on initiator-immobilized sites. 77

2.11 Hybridization of various targets (Sequences 5–7) to immobilized

probe spots. 78

2.12 Confocal images of hybridized targets (Sequences 5–7) on a

mixed film glass surface and their fluorescence intensities as

temperature was increased.

80

3.1 1H NMR spectrum of BZ-APTMS in CDCl3. The inset contains 85

xix

the structure of BZ-APTMS. Different types of protons are

labeled.

3.2 1H NMR spectrum of the activated bromoisobutyryl NHS ester

initiator. The inset shows the structure of the bromoisobutyryl

NHS ester. The spectroscopically different protons are labeled.

87

3.3 (a) 13C NMR (500 MHz) spectrum of the activated

bromoisobutyryl NHS ester initiator. The inset shows the

structure of the bromoisobutyryl NHS ester. The

spectroscopically different carbon atoms are labeled; (b) high

resolution data showing detail of the downfield region.

88

3.4 1H NMR of PHEMA in DMSO-d6. Structure of PHEMA. The

inset shows different types of protons, which are labeled.

90

3.5 High resolution XPS spectra of N1s and the peak locations of the

various nitrogen-containing species with their atomic percentages.

(a) Cleaned Si wafer; (b) APTMS-BZ modified Si wafer ; (c)

hydrolyzed APTMS Si wafer

100-

101

3.6 AFM images of (a) RCA cleaned bare Si wafer; (b) APTMS-

modified Si wafer; (c) BZ-APTMS modified Si wafer; (d)

PHEMA grown on BZ-APTMS-coated Si wafer (~10 nm thick).

104

3.7 ToF-SIMS ion images of EGMEM on Si wafers grown via ATRP.

(a) cleaned Si wafer; (b) initiator-immobilized Si wafer

106

3.8 Change in elemental composition in an angularly-resolved XPS

experiment of C 1s and Si 2p of Si wafers that were and were not

immobilized with the ATRP initiator using EGMEM as monomer.

(•) no initiator, Si 2p; (♦) initiator, C 1s; ( ) no initiator, C 1s;

(▲) initiator, Si 2p

107

3.9 Low resolution XPS spectra of (a) C 1s at 0 h of ATRP; (b) Si 2p 108

xx

at 0 h of ATRP; (c) C 1s at 1 h of ATRP; (d) Si 2p at 1 h of ATRP.

EGMP was used as monomer.

3.10 Reusability of mixed film of oligonucleotides and PHEMA-

modified glass slide for hybridizations. Note that 3 bpm targets

were used for hybridizations 4 and 5 whereas fully complementary

SMN targets were used for hybridizations 1-3.

114

3.11 Melt curves of fully complementary targets collected from: (■)

oligonucleotide film only; (▲) solution (0.5 × PBS); (●) mixed

film of oligonucleotides and PHEMA. “fssDNA” represents

fraction of single-stranded DNA.

115

3.12 Melt curves of 3 bpm targets collected from: (■) oligonucleotide

film only; (▲) solution (0.5 × PBS); (●) mixed film of

oligonucleotides and PHEMA. “fssDNA” represents fraction of

single-stranded DNA.

116

3.13 High resolution C 1s spectra of all surfaces (A-D) investigated; (a)

Surface A: aminosilanes only; (b) Surface B: oligonucleotides

only; (c) Surface C: mixed film; (d) Surface D: PHEMA only

125

3.14 TEM of 5 nm gold nanoparticles after conjugation with thiolated

SMN probes.

128

3.15 Fluorescence intensity of Cy3-labeled thiolated SMN probe

immobilized onto 5 nm gold nanoparticles (⎯) before addition of

40 mM KCN and (⎯) after addition of KCN. λex = 520 nm.

129

3.16 (a) SEM image of Au np-oligonucleotide targets hybridized to

surface-bound oligonucleotide probes on PHEMA-modified

surfaces; (b) SEM image of Au np-oligonucleotide targets

adsorbed onto PHEMA-modified surfaces (no oligonucleotide

probes were immobilized).

131-

132

xxi

3.17 Possible origins of (a) [C3H5NO2+H]+; (b) [C2HNO2+H]+

fragments.

134

3.18 Proposed fragmentation of PHEMA for selected positive and

negative fragments.

134

3.19 Overlaid ToF-SIMS spectra when Bi3+ was used for the following

ion fragments: (a) PO3- at 78.96 m/z; (b) C4H5O2

- at 85.03; (c)

C2H2NO2+ at 71.96; (d) C2H5O+ at 45.02 m/z.

135-

136

3.20 Peak intensities of (a) PO3- from nucleotides; (b) C4H5O2

- from

PHEMA under various types of Bin+ (n = 1, 3, 5) primary ion

source collected from various surfaces (A-D).

137

3.21 Peak intensities of (a) C2H2NO2+ from nucleosides; (b) C2H5O+

from PHEMA under various types of Bin+ (n = 1, 3, 5) primary ion

source collected from various surfaces (A-D).

139

3.22 Comparison of peak intensities of selected positive fragments from

PHEMA and oligonucleotides between the Bi3+ and Bi5

+ primary

ion sources at various concentrations; (a) C2H5O+ and C4H5O+

fragments from PHEMA; (b) C4H5N4+ and C6H7N2O+ from

oligonucleotides. The control was a mixed BZ-APTMS and

APTMS; conc1 was 0.10 μM, conc2 was 0.48 μM, and conc3 was

1.02 μM (loading concentrations).

144

3.23 Integrated fluorescence intensities of various targets introduced to

oligonucleotide probes (Sequence 1). Standard deviations are

shown.

154

3.24 Integrated fluorescence intensities of various targets introduced to

areas with and without immobilized oligonucleotide probes.

Standard deviations are shown.

154

xxii

3.25 Fluorescence intensity decay of Cy3-ssDNA in different

environments: upper decay corresponds to the mixed film and the

lower one to the oligonucleotide film. Raw data in black, bi-

exponential fit curves in red and green. See table 3.8 for the

numeric results of the fitting analysis.

156

3.26 The structure of 5′-Cy3-labeled oligonucleotides. 157

3.27 Proposed stacked and unstacked interactions of Cy3 with

immobilized oligonucleotide probes (a) in an oligonucleotide film;

(b) in a mixed film.

158

3.28 Fluorescence images of (a) oligonucleotide-only film, and (b)

PHEMA film on glass cover slips. The intensity variations in the

PHEMA image are mostly due to imperfect flattening of the

excitation field in a wide-field microscope.

159

3.29 Individual melt curves of (•) SNP targets and (■) FC targets that

represent average Tm and dx values collected from oligonucleotide

(DNA) and mixed films in increasing PBS strengths.

165-

166

3.30 Interpretation of melting transition width and Tm differences

between different targets hybridized to the mixed film.

168

xxiii

List of Equations and Chemical Reactions

Equation

#

Equation Page #

1.1 Dependence of melting temperature on the percentage of C and G 12

1.2 Factors that determine the fluorescence intensity 20

1.3 Quantum yield described by radiative and non-radiative processes 21

1.4 Quantum yield described by radiative decay and fluorescence

lifetime

21

1.5 FRET 22

1.6 Fluorescence lifetime 23

1.7 Snell’s law 24

1.8 Evanescent wave intensity described by penetration depth and the

distance normal to the surface

25

1.9 Penetration depth 25

1.10 Intensity of electric field 25

1.11 Adsorption of thiols onto gold surfaces 31

1.12 Oxidation of sulfur to thiolate 32

1.13 Hydrolysis of alkoxysilanes 36

1.14 Condensation of silanols 36

1.15 Condensation of silanols with alkoxysilane 36

1.16 Weighted average molecular weight 44

xxiv

1.17 Number average molecular weight 44

1.18 Coupling reaction observed in radical polymerization 44

1.19 Disproporportionation reaction observed in radical polymerization 45

1.20 Termination reaction observed in radical polymerization 45

1.21 Abstraction of halogen atom by copper catalyst to yield a radical 48, 96

1.22 Polymerization of monomer onto radical end of polymer 48, 97

2.1 Calculation of the fraction of single-stranded DNA 79

3.1 Linear regression of the ATRP reaction rate on initiator-

immobilized Si wafers

105

3.2 Linear regression of the ATRP reaction rate on Si wafers without

initiators

105

3.3 Calibration curve of Cy3-modified oligonucleotides to determine

the number of oligonucleotides immobilized on Au nps

129

3.4 Sigmoidal equation 160

xxv

List of Appendices

Appendix

#

Title Page #

A.1 Kinetic study of the growth of PHEMA via ATRP on surfaces

modified with and without initiator.

189

A.2 Comparison of ATRP rates on mixed BZ-APTMS and APTMS

and free APTMS surfaces.

190

A.3 Overlaid ToF-SIMS (a) positive and (b) negative spectra obtained

by using Bi+ on various surfaces (top to bottom: mixed APTMS

and BZ-APTMS, oligonucleotides only, mixed oligonucleotides

and PHEMA, PHEMA only).

191


by using Bi3+ on various surfaces (top to bottom: mixed APTMS



193


by using Bi5+ on various surfaces (top to bottom: mixed APTMS



195

A.6 Overlaid ToF-SIMS positive spectra obtained by using Bi3+ on

surfaces containing various ratios of oligonucleotides to PHEMA

(top to bottom: control, conc1, conc2, and conc3. Conc1 was 0.10

μM, conc2 was 0.48 μM, and conc3 was 1.02 μM (loading

oligonucleotide probe (sequence 2) concentrations).

197

A.7 Overlaid ToF-SIMS positive spectra obtained by using Bi5+ on

surfaces containing various ratios of oligonucleotides to PHEMA

201

xxvi

(top to bottom: control, conc1, conc2, and conc3. Conc1 was 0.10

μM, conc2 was 0.48 μM, and conc3 was 1.02 μM (loading

oligonucleotide probe (sequence 2) concentrations).

1

Chapter 1

1 Introduction

1.1 Overview Water and food contamination cases in Ontario, Canada, such as the E. coli

contamination of water in Walkerton, ON (May 2000) [1] and the Listeriosis contamination of

deli meat in a Maple Leaf Foods plant (August 2008) [2], as well as the outbreaks of Severe

Acute Respiratory Syndrome (SARS) (March 2003) [3] and swine flu (May 2009) [4] have

dominated headlines over the last decade. These incidents have reminded us that early detection

and fast response time are critically needed to be able to keep any contaminated water and food,

and disease, from spreading further. Pathogens such as bacteria and viruses are usually

identified by using markers associated with their genetic code or deoxyribonucleic acid (DNA).

DNA is known for its complementarity, in a process by which one strand can “recognize” the

other complementary strand through specific non-covalent interactions that lead to DNA

hybridization [5]. Using the knowledge that each organism possesses its own unique genome,

DNA can be used as a specific molecular recognition element to identify the presence of

pathogens in a sample.

Gel electrophoresis combined with Southern blotting are methods typically used to detect

specific DNA sequences [6]. Genetic tests, such as those for single nucleotide polymorphisms

(SNP), also use these methods to screen for certain genes associated with diseases. Molecular

biology techniques usually involve time-consuming and laborious steps such as amplification,

enzymatic, and separation steps [6, 7]. Therefore, a more rapid, sensitive, and selective tool is

desired.

2

Biosensors are devices that can satisfy the above requirements. They can convert a

biological or biochemical reaction into a measurable analytical signal [8]. Representative

reactions involve the specific binding of antibodies to antigens, substrates to enzymes, and DNA

strands to complementary DNA strands [8]. Several types of transducers are available, with the

most common being based on piezoelectric, electrochemical, and optical platforms [8]. The

optimization of the properties listed above would make biosensor technology attractive, with the

ideal biosensor also offering field portability and reusability or disposability, and without need

for reagent labels [6].

The interest in this thesis is to develop an optically-based nucleic acid biosensor

chemistry that improves selective discrimination of fully complementary target sequences from

targets that contain SNPs. SNPs contain one base-pair which is not complementary to the probe

sequence [9]. As markers, they are often linked to cardiovascular and autoimmune diseases,

cancer, and psychiatric disorders [9] such as Parkinson’s, Alzheimer’s, and diabetes [10-15].

The DNA sequence that will be used as a model for this work is a short sequence or

oligonucleotide from the Survival Motor Neuron 1 (SMN1) gene, which causes spinal muscular

atrophy in children (SMA) [7]. Not having adequate copies of the gene in a cell can lead to

reduced SMN1 protein levels, which is indicative of SMA [7]. Moreover, a single mutation in

the SMN1 gene can result in the SMN2 gene, which yields a less stable, truncated SMN protein

[7]. It has been discovered that a low copy number for SMN1 is indicative of the disease and the

number of SMN2 copies per cell can predict how severe the disease would be [7]. Thus, this

selection of target offers a real example of how crucial it is to selectively detect and quantitate

each gene sequence with or without SNPs. This task is not trivial since the presence of a SNP

can alter the thermal stability of the DNA duplex by more than 4 °C in solution [16]. However,

3

the melting transition range in interfacial melt curves is usually broadened, reflecting the

diversity of energetics at an interface, hence providing even lower selectivity for SNP detection

[17]. Therefore, selectivity must be maximized to optimize the discrimination of the two

sequences.

The key to increase selectivity between two nearly identical target sequences is to control

the local environment on the surface of the biosensor at the nanoscale level. A “matrix isolation”

concept was proposed in the work by Piunno et al. [18], which entailed the inclusion of nucleic

acid probes with non-nucleic acid oligomeric brushes; this concept is depicted in Fig. 1.1. The

co-immobilization of oligomers would cause the oligonucleotides to exhibit a more upright

conformation with respect to the surface (Fig. 1.2). Additionally, this may reduce any cross

hybridizations (hybridization of one target strand with more than one probe), non-specific

adsorption, and interactions of oligonucleotide probes with neighbouring probes and with the

surface [18]. It was anticipated that the addition of non-nucleic acid material would serve such a

purpose, rendering a highly selective DNA biosensor capable of SNP detection. By collecting

melt curve data, such differences in surface free energies can be distinguished because the

surface would yield a lower number of non-specifically adsorbed DNA and intermediate dsDNA

conformations, leading to a much narrower transition from double-stranded to single-stranded

DNA (Fig. 1.3). This thesis proposes a simple but effective strategy for creating a mixed

oligonucleotide and oligomer film which involves growing the oligomer portion of poly(2-

hydroxyethyl methacrylate) via surface-initiated atom transfer radical polymerization on silica

based substrates which have already been immobilized with oligonucleotide sequences of SMN1.

This thesis discusses the characterization of each immobilization step towards the making and

characterization of the mixed film, and the comparison of oligonucleotide and mixed films to

4

discriminate SNPs from fully complementary targets. Before providing details about this novel

sensing surface, some background about DNA and biosensors will first be introduced.

Fig. 1.1 : Depiction of a mixed film composed of oligonucleotides and poly(2-hydroxyethyl methacrylate) brushes

immobilized on a silica-based substrate.

(a)

(b)

Fig. 1.2: Proposed oligonucleotide conformations in (a) oligonucleotide films; (b) mixed films.

5

Fig. 1.3: Proposed interfacial melt curves from (a) oligonucleotide films; (b) mixed films.

1.2 Principles of Nucleic Acid Properties, Structure, and Function

1.2.1 Deoxyribonucleic Acids (DNA)

Genetic information is stored in the nucleus of all eukaryotic or multicellular organisms,

or the nucleoid region in prokaryotic or unicellular organisms [19]. The biomolecule which

stores gene information is called deoxyribonucleic acid (DNA); it is composed of a chain of

molecular units composed of a deoxyribose sugar, a nitrogenous base, and a phosphate group.

These three components form a nucleotide, and nucleotides are strung together to form a

polyelectrolyte macromolecule known as DNA. The structure and function of DNA was first

described by Watson and Crick in 1953 [20] and was reinforced by the X-ray diffraction data of

DNA contributed by Wilkins, Stokes, and Wilson. The latter paper was published immediately

after Watson and Crick’s seminal publication in Nature [21].

Genes are the basic physical and functional units of heredity which contain the DNA

information needed to transcribe into messenger ribonucleic acid (mRNA), which can be

translated into specific proteins [22]. Other RNA molecules which are used in the synthesis of

proteins are transfer RNA (tRNA), each of which brings in specific amino acids, and ribosomal

RNA (rRNA), which is the assembly plant that connects all the amino acids into a polypeptide

6

chain [23]. Other types of RNA include small interfering RNA (siRNA) and micro RNA

(miRNA), which have gene regulatory functions [24].

Each gene is based on a serial information sequence created from four possible

nucleotide units, each containing a different nitrogenous base. Sequence information is decoded

from the DNA by copying the DNA code into mRNA using DNA as a template, and then

synthesizing the protein from the mRNA. Each gene is unique due to different sequences of

nucleotides. With only four different DNA bases, it is possible to produce an incredible richness

of different sequences. For example, it is possible to find 425 ≅ 1015 unique sequences for a short

25 base-pair DNA sequence or oligonucleotide [25]. As for protein-coding genes, about 100,000

proteins were expected in the human proteome [22]. Surprisingly, according to The Human

Genome Project, which has completed sequencing the entire human genome, has corrected the

previously estimated number of protein-coding genes to 20,000–25,000 genes [22].

As briefly mentioned earlier, DNA contains three components. The sugar and phosphate

groups serve as the backbone of the DNA, linking the deoxyribose sugars together from the C5

position of the lower sugar to the O3 position of the upper sugar through phosphodiester bonds

(Fig. 1.4). Nitrogenous bases are attached to the C1 atom of the deoxyribose sugar and they are

divided into purines and pyrimidines. Figure 1.5 shows that purines are nine-membered double-

ringed nitrogenous compounds, which in the case of DNA are guanine (G) and adenine (A) [19].

Pyrimidines, which are six-membered single-ringed nitrogenous compounds, include cytosine

(C), thymine (T), and uracil (U) [19]. Uracil substitutes for thymine in RNA chemistry. Instead

of having deoxyribose sugar, RNA uses ribose sugar (contains an extra hydroxyl group at the 2′

position) [19].

7

Fig. 1.4: 5′- to 3′-end structure of a dinucleotide – a string of 2 nucleotides – that composes of guanine and adenine

on one strand and cytosine and thymine on the complementary strand. Dotted lines indicate hydrogen bonds

between A:T and G:C base pairs.

N

N

NH

N

NH2

Adenine

N

NH

NH

N

NH2

O

Guanine

N

NH

NH2

O

Cytosine

NH

NH

O

O

CH3

Thymine

NH

NH

O

O

Uracil

(a)

(b)

Fig. 1.5: Chemical structures of nitrogenous bases present in deoxyribonucleotides and ribonucleotides; (a) purines;

(b) pyrimidines.

8

1.2.2 Base-pairing

Base pairing involves the association of two different DNA strands via hydrogen bonds.

From Chargaff’s discovery that the molar ratio of A:T and C:G were both unity [26], Watson

and Crick deduced that each base pair binds together through hydrogen bonds and that A

associates with T and C has an affinity for G [5]. This is known as Watson-Crick pairing.

Figure 1.4 shows that the two base-pairs are held together by hydrogen bonds. −NH groups are

usually good hydrogen bond donors while sp2-hybridized electron pairs on oxygens of C=O and

on ring nitrogens are better hydrogen bond acceptors than oxygen in phosphate or pentose [27].

Watson-Crick hydrogen bonds are 2.8–2.95 Å apart with 68 ± 2° between two glycosylic bonds

for both A-T and C-G base pairs, a distance of 10.60 ± 0.15 Å from the C1 (deoxyribose sugar

carbon attached to nitrogenous base of one DNA strand) to the C1 (deoxyribose sugar carbon

attached to complementary nitrogenous base) [27]. There are three hydrogen bonds for the G-C

pair and only two hydrogen bonds in the A-T pair. The G-C content of a particular DNA is one

of the predictors of helix stability under stringent conditions such as high temperature, pH and

low salt concentrations [19].

9

Fig. 1.6: Watson-Crick model of the DNA double helix (Adapted from [19]). The ribbon-like strands represent the

phosphate backbone, and the parallel, horizontal rungs represent the nitrogenous base pairs.

1.2.3 Structure of DNA

Figure 1.6 shows the conformation and structural features of a conventional DNA found

in cell nuclei with Watson-Crick pairing. Figure 1.4 shows that each DNA strand is directional.

Each DNA strand starts with a free phosphate group on the 5′-carbon of a deoxyribose unit and

terminates with the 3′-carbon of deoxyribose with its –OH group [19]. The notation is important

because when two DNA strands hybridize (the process of two complementary strands coming

together by base pairing) the two DNA strands run antiparallel of each other, that is, one strand

runs in the 5′ to 3′ direction while the other strand runs from 3′ to 5′ [19]. The antiparallel

directions and the asymmetry of the base give rise to structural features known as the major and

10

minor grooves of the DNA helix, which is shown in Figure 1.6 [5]. The directionality of DNA

strands is important not only for biomolecular processes but for biosensor design. For example,

it is essential to decide in advance which terminal end will be immobilized to the biosensor

surface. This will affect the sequence of the target strand, location of mismatched bases (if any),

and location of intrinsic labels such as fluorophores.

The nitrogenous bases are stacked on top of one another and are separated by a distance

of 3.4 Å. One complete turn of the helix contains 10 nitrogenous bases or 34 Å long [19]. The

diameter of the double helix is 20 Å [19]. This DNA structure, called B-DNA, is only one of the

three types of DNA structures [19]. A- and Z-DNA also exist [19]. They differ in the separation

distances between bases and the energetics by which the two strands interact [19].

1.2.4 Energetics of the Double Helix

Hybridization or the association of the complementary strands of short oligonucleotides

is a bimolecular event which is concentration dependent, with a rate constant of 106 M-1 s-1 [27].

Nucleation of hybridization of DNA strands is rate-limiting at low concentration but each duplex

hybridizes to completion almost instantly (> 1000 bp s-1) [27]. Short oligonucleotides undergo

nucleation, zippering up, and loop-migration during hybridization [27]. For longer DNA,

hybridization involves numerous steps including the formation of intrastrand loops, a slow

nucleation step, loop formation, and zippering up [27].

The stability of DNA hybrids is affected by sequence composition, secondary structure,

strand length, ionic strength, pH, and temperature [28]. Furthermore, there are two major forces

that stabilize double-stranded DNA, which are hydrogen bonding and base-stacking [29]. Base-

11

stacking is the dipole-induced dipole-dipole interaction between two contiguous bases [30].

Šponer and co-workers have discussed the results obtained by advanced ab initio quantum

chemical calculations concerning the electronic properties, hydrogen bonding, and stacking of

DNA bases [31]. This review supported the findings that base stacking is stabilized by

dispersion forces, and it summarized data concerning hydrogen bonding of DNA bases [31].

Studies have also shown that base stacking at a nick introduced by two contiguous DNA tandem

sequences hybridized to a longer strand adds stability and efficiency to the hybridized DNA [30].

Yuan and co-workers used an SPR-based DNA biosensor to compare the association and

dissociation rate constants of DNA hybridization between samples that contain base stacking and

those without, and determined that base stacking assists in DNA hybridization [30]. Guckian et

al. compared base stacking capability between natural bases and non-nucleic acid aromatic

compounds such as pyrene [32]. The study led to the conclusion that the order of base stacking

capability from highest to lowest is: A > G > T = C, noting that purines are more effective in

base stacking than pyrimidines [32]. Their paper showed that a large surface area has a positive

effect on base stacking, whereas dispersion forces and polarizability of the bases are weak

contributors [32]. The other major factor in promoting base stacking is the solvent-driven

hydrophobic effect, which occurs when the solvation of hydrophobic aromatic bases is not

energetically favourable and results in the expulsion of water molecules [32].

There are a few ways that the DNA duplex can become temporarily separated under the

non-denaturing conditions. A process known as DNA breathing occurs when the terminal ends

are not hydrogen bonded to each other [27]. The strands in those regions are temporarily

separated by a short distance. Another event known as “soliton excitation” occurs when the

stretching vibration of DNA chain travels like a wave longitudinally along the helix axis [27]. If

12

there is sufficient energy, it would cause a local unstacking of adjacent bases involving the

deformation of sugars and other bond conformations [27].

1.2.5 Denaturation of the Double Helix

The separation or denaturation of a double helix is a cooperative process and can be

accomplished by increasing the temperature, raising the pH, lowering the salt concentration, or

adding chaotropic agents [19]. These factors cause increased phosphate group repulsion [33]

and/or decreased hydrogen bonds, triggering the helix to unwind, and the strands to separate

[19]. The ease of denaturation is also a function of sequence, presence of mismatches, and chain

length [27]. Homopolymers have a sharper melting transition than random-sequence polymers

since the presence of a single type of base-pairing would mean all the base pairs should melt at

the same temperature, versus a range of temperatures for a mixed polynucleotide. It would take

more heat to denature homopolymers containing G-C pairs than A-T pairs due to a higher

number of hydrogen bonds. Each mismatch on a sequence can lower the stability by 2 °C (for an

A-T pair) or 4 °C (for a G-C pair) [16].

The process of denaturation is often tracked by observing the melt curve of a particular

DNA sequence. The melting temperature, Tm, is designated as the temperature at which 50% of

the double-stranded DNA is denatured [19]. This value can be predicted by the following

equation for oligonucleotide duplexes:

Tm = X + 0.41 [%(C+G)] (1.1)

where X is a constant dependent on salt concentration and pH and it has a value of 69.3°C for 0.3

M sodium ions at pH 7 [19]. Tm is also dependent on DNA concentration [33]. Owczarzy and

13

co-workers have proposed a new general empirical equation to predict the melting temperature

based on salt concentration and G-C content but is independent of DNA concentration and

oligonucleotide length [34].

The proportion of DNA that is denatured can be determined spectroscopically by

illuminating the DNA sample with a UV source at 260 nm [19]. Due to a phenomenon known as

hypochromicity, in which base stacking in the double-stranded form decreases molar extinction

coefficient, the transition from double-stranded DNA (dsDNA) to single-stranded DNA (ssDNA)

can be tracked by an increase in absorption at 260 nm [27]. This is because when two ssDNA

hybridize to form dsDNA, the nitrogenous bases that are located inside the DNA helix are

shielded from the external environment. Therefore, ssDNA absorbs UV stronger than dsDNA

[19]. The molar absorptivity constant of oligonucleotides is around 104 dm3 mol-1 cm-1 but it is

affected by base composition, state of base-pairing interactions, salt concentration and pH.

Ribose-phosphate, on the other hand, is transparent to UV. Therefore, UV absorption

spectroscopy is very specific for quantifying the amount of DNA via the presence of the

nitrogenous bases. Figure 1.7 shows the typical shape of a melt curve collected from solution by

plotting the absorbance of DNA at 260 nm against temperature. Absorbance or any other signals

such as fluorescence can be normalized to quantify the fraction of either ssDNA or dsDNA as the

temperature is increased. The plot reveals the sigmoidal nature, signifying a two-state transition,

of the melting profiles of dsDNA. A comparison of Tm values between two DNA samples, one

of which contains a SNP, shows that the sample with the fully complementary strand has a

higher Tm due to more hydrogen bonds being present to stabilize the DNA helix [19].

14

20 25 30 35 40 45 50 55 60 65 70 75 80 85 90

-0.2

0.0

0.2

0.4

0.6

0.8

1.0

1.2

FC SNP

F ssD

NA

Temperature ( oC)

Fig. 1.7: Experimental melt curves of SMN hybrids in 0.5× PBS. FC: fully complementary; SNP: single nucleotide

polymorphism. FssDNA represents the fraction of single-stranded DNA.

1.2.6 Effect of pH on DNA Structure

The various ionizable states determine the DNA tautomeric structure, overall charge, and

most importantly, the ability to participate in hybridization. All bases, as well as pentose sugars,

are uncharged from pH 5–9 [27]. Under acidic conditions, adenine, guanine, and cytosine

become protonated at the ring nitrogens but not at the exocyclic primary amines [27]. Each

phosphate group is singly charged, which adds repulsive effect when two strands are hybridized.

The first proton in the phosphate groups deprotonate at pH 1 and the second ionization occurs at

pH 7 [27], which causes DNA to be highly sensitive to pH at physiological conditions.

15

1.2.7 Oligonucleotides as Probes for Recognition of Target Sequences

Molecular biorecognition is the highly specific association of biomolecules. Examples of

molecular recognition elements are mainly proteins and nucleic acids. Subclasses of proteins

used are enzymes, antibodies, and cell membrane receptors [35] [36]. They are used due to their

specificity towards substrates, antigens, and cellular solutes, respectively. For the detection of

nucleic acids, the base complementarity makes DNA detection very selective and reversible

because no covalent bonds are formed [37, 38]. During a hybridization event, the selectivity can

be tuned by varying the sequence, length, and stringency conditions as already discussed [36].

For practical purposes, DNA is a better candidate probe than RNA to develop nucleic

acid biosensors. First of all, the phosphate backbone of DNA does not succumb to hydrolysis as

easily as RNA, due to the absence of an extra –OH group on the 2′-carbon [19]. Moreover, the

advent of polymerase chain reaction (PCR) and solid phase nucleic acid synthesis have made the

preparation of DNA probe sequences very straightforward [37].

In biosensing applications, immobilized probes of 10–30 nucleotides are typically

employed [36]. Assays that involve longer probes tend to take a longer period of time because

there are more bases to be hybridized [36]. For shorter probes of 10–20 nucleotides,

hybridization is complete in minutes compared to hours for 100 nucleotide-long probes under

identical conditions such as ionic strength, temperature, and pH [36]. Furthermore, shorter

probes are less likely to hybridize in the presence of mismatches compared to longer probes for

the same number of mismatches because of the lower probability of finding complementary

sequences. For SNP studies, it has been shown through a mathematical model that permutations

of oligonucleotides of at least 17 bases are required to detect every possible unique sequence

from a randomly-organized DNA sample with the length of the human genome [39]. PCR

16

products of about 30 bp long are used in forensics cases to study different sets of SNPs (usually

50-60 SNP loci) [40]. One can use a probability function to determine if two sets of SNPs are

identical within a statistical range [40]. Therefore, oligonucleotide probes are generally 20–30

bp long.

Oligonucleotides can also be easily modified with chemical functional groups or

molecules such as biotin [33] (See Chapter 1.4) for immobilization purposes. To detect DNA

hybridization, numerous types of labels such as radioisotopes, enzymes, colourimetric reporter

groups, luminescent labels, and electroactive species have been attached to DNA [33, 35]. Non-

labeled oligonucleotides are also used in biosensing systems such as surface plasmon resonance

(SPR) and quartz crystal microbalance (QCM) [6]. Electrochemical approaches have relied on

the redox reaction of guanine as an intrinsic label for DNA [41]. Therefore, DNA probes are

versatile and can be used in a variety of sensing platforms.

1.3 DNA Biosensors A biosensor is defined as an analytical device that contains biological recognition

elements that are closely associated with a transducer and that are able to interact with target

molecules [36]. Transducers are components of a biosensor that convert a biological or chemical

binding event into an analytical signal as illustrated in Figure 1.8 [36].

17

Fig. 1.8: Concept of a biosensor. The binding of the target DNA strands to the immobilized DNA probes

(biological reaction) can be detected via the fluorescence emission of a dye due to intercalation (analytical signal).

The choice of the transducer in part depends on the type of biochemical reaction, the

targets, whether labels are used, and the analytical performance requirements. For example, the

very first biosensor was invented in 1953 for the determination of glucose. By using the

knowledge that glucose is oxidized in the human body and is converted to gluconic acid by the

enzyme glucose oxidase, this redox reaction is coupled to a reduction reaction of oxygen into

hydrogen peroxide by using a Clark electrode [35]. This technological advance ultimately

18

impacted millions of people who are afflicted with diabetes, and initiated the biosensor research

area. Beyond electrochemical transduction, biosensors based on piezoelectric and optical

measurements were developed. Table 1.1 lists the earliest demonstrations of each type of DNA

biosensor.

Table 1.1. Various types of biosensors based on DNA hybridizations.

Biosensor type Transducer Description Reference

Electrochemical – cyclic

voltammetry

Glassy carbon

electrodes

Used cobalt complexes as

electroactive markers of DNA

hybridization

[38]

Piezoelectric Quartz crystal

microbalance

Demonstrated difference in

resonance frequency change between

complementary and non-

complementary DNA

[42]

Optical - Surface

plasmon resonance

(SPR)

Gold (BIAcore ™ SPR) Detected hybridization within 10

minutes

[43]

Another type of an optical biosensor is based on the fluorescence transduction of DNA

hybridization on optical fibers, which was introduced in 1994 [44]. Since the goal of this work is

to improve the selectivity of the nucleic acid biosensor based on silica-based substrates, the focus

of this chapter is to discuss the background theory, pioneering work, and recent advances in

DNA optical biosensor development using fluorescence as a transduction method for DNA

hybridization.

19

1.3.1 Optical Biosensors

Optical sensors use the properties of light, which include absorbance, reflectance,

fluorescence, (bio- or chemi-) luminescence, and refractive indices to detect specific targets [45].

Optical biosensors are known to be non-destructive as well as having sensitivity and speed [8].

Specifically, fluorophore-labeled oligonucleotides and the use of DNA intercalators are

commonly used to signal the presence/absence of DNA hybridization at an optical fiber interface

[44, 46, 47]. This section will describe the detection of DNA targets by using fluorescence and

the total internal reflection (TIR) excitation of fluorophores. A brief review of the principles of

fluorescence spectroscopy is also presented.

1.3.2 Fluorescence

Fluorescence is an electronic relaxation process that occurs after an electron has been

promoted to an excited electronic energy level in the singlet state by the absorption of photons.

The excited electrons can return to the ground state first via vibrational relaxations, for example,

to the lowest vibrational state of the excited state, and then releasing energy in the form of light.

This is called fluorescent emission, which has a wavelength longer than the excitation

wavelength, due to the loss of energy through non-radiative relaxation processes [48]. Figure 1.9

is a Jablonski diagram which describes this process. The shift to longer wavelengths in

fluorescence emission is known as Stokes’ shift [48]. The Stokes’ shift also indicates the

presence of collisional processes between molecules, solvent effects and reactions during the

residence of the molecule in the excited state [48]. Therefore, it is an important parameter to

consider for these studies.

20

Fig. 1.9. Electronic transition of absorption and fluorescence.

The fluorescence intensity is dependent on molar absorptivity (ε) of the fluorophore,

pathlength of the sample holder (b), and concentration of the fluorescent analytes (C), as

described by the Beer-Lambert law [48]. Additionally, fluorescence is affected by the quantum

yield (Φ), the instrument response coefficient (K), and the initial intensity of the excitation

radiation (I0) as described by the following equation:

Φ= bCKIF ε0303.2 (1.2)

21

which is for a dilute concentration of the fluorescent species (< 0.01 M) [48]. However, in a

population of excited fluorescent molecules, they may not all undergo relaxation via

fluorescence emission, as some will relax back to the ground state via non-radiative decay. The

quantum yield is a parameter used to describe the efficiency of fluorescence emission, and it can

be approximated by examining the rates of radiative decay (kr) versus non-radiative decay (knr)

[48]:

)/( nrrr kkk Σ+=Φ (1.3)

Since the quantum yield is dependent on decay rates, the lifetime of the excited state or

fluorescence decay time of the singlet state (τs) of the fluorescent molecule can also be

described:

srk τ=Φ (1.4)

Fluorescence spectroscopy allows one to collect a variety of information about the

molecule in question as well as its environment. Thus, fluorescence has been dubbed

“multidimensional” [49]; it can be divided into two kinds of measurements, namely static and

dynamic. Table 1.2 provides details on the kind of information which can be obtained by each

fluorescence feature [49].

22

Table 1.2. Types of static and dynamic fluorescence measurements. Adapted from [49].

Static measurements Information

Intensity Concentration, quenching

Spectral Local environment, number of emissive

components, average distance between sites due to

energy transfer

Polarization/Anisotropy Average size of a rotationally reorienting species,

range of mobility or rotational freedom

Dynamic measurements

Excited-state intensity decay Resolve static emission into contribution from each

individual emissive centers, determine excited-state

decay kinetics, find origin of quenching processes

Excited-state decay of anisotropy Determine reorientational dynamics of non-

spherical rotors, shape of rotating body, compare

differences in local and global motions, find how

surfaces affect solute mobility and dynamics

One type of static measurement is a fluorescence resonance energy transfer (FRET)

experiment which can determine the average distance between two sites. It occurs when an

excited donor molecule undergoes relaxation by transferring its energy, via a near-field non-

radiative process, to an acceptor molecule [48].

D* + A D + A* (1.5)

If there is sufficiently short distance between D and A, and if fluorescence emission and

absorption spectra overlap, then it is possible that FRET would occur. The significance of the

23

overlap is such that there are enough vibronic transitions which are equal in energy for both

acceptor and donor [48]. These transitions occur due to short-range intermolecular orbital

overlap or long-range Coulombic interactions [48]. The latter interaction involves dipole-dipole

interactions, which are also known as the Förster quenching [48]. These interactions lead to an

excited electron localized on the LUMO of the donor to pass on its energy to an electron in the

HOMO of the acceptor, resulting in promotion of the electron to the LUMO of the acceptor

molecule [48]. Long-range interactions can occur within 8–10 nm, whereas short-range

interactions occur within few tens of Angstroms [48].

If the quantum yield changes due to environmental changes, then the lifetime would be

affected as well. Therefore, lifetime measurements are often carried out to probe the nature of

the environment. The lifetime is defined as the average time that a molecule spends in its excited

state before returning to the ground state [50]. In general, the lifetime is related to the emissive

rate of the fluorophore and the rate of radiationless decay:

nrr kk +=

1τ (1.6)

In general, the average fluorescence lifetime of organic dyes is approximately 10 nsec [50].

1.3.3 Total Internal Reflection and Evanescent Waves

Although fluorescence has the ability to provide abundant physical information and a low

detection limit, it is not very surface selective [49]. This last feature is especially significant for

sensing applications. A sophisticated way to interrogate surface-bound or near-surface localized

fluorescent molecules is total internal reflection fluorescence (TIRF) [49]. A method first

24

developed by Hirschfeld [51] in 1965, TIRF has since emerged as a powerful optical technique

that allows the study of fluorescent molecules near an interface. Surface-localized excitation

occurs when an incident beam of electromagnetic radiation passes from a medium of higher

refractive index (n1) to one which has a lower refractive index (n2) at an angle greater than the

critical angle (θc). Such a medium is often found in optical fibers, for which the optical fiber

core has a higher refractive index than the cladding [52]. The critical angle is defined by the

Snells’ law as [48]:

1

2sinnn

c =θ (1.7)

When this occurs, a small fraction of the incident light will penetrate into the lower refractive

index medium, and the process is associated with the evanescent wave [49]. The generation of

an evanescent wave is shown in Figure 1.10.

Figure 1.10: The generation of evanescent waves during total internal reflection. Adapted from [48].

25

The intensity of the resultant evanescent wave is given by equation 1.8:

Λ−

=x

eEE 0 (1.8)

where E is the intensity of the evanescent wave, E0 is the intensity of electric field at the

interface, x is the distance normal from the interface, and Λ is the penetration depth [53]. As

equation 1.8 states, the intensity of the evanescent wave field will decay exponentially as a

function of the distance from the surface [54]. The penetration depth is defined as the distance

into the second medium orthogonal to the surface at which the electric field has decreased to 1/e

of its interfacial value [53]. It is related to the wavelength of the excitation source, the incident

angle, n1, and, θc, as given by Equation 1.9:

2/1221 )sin(sin4 cin θθπ

λ−

=Λ (1.9)

where λ is the wavelength of the excitation source and the rest of the parameters are defined as

previously. The penetration depth usually spans from 100 nm to 200 nm from the surface [49,

54]. Thus, fluorophores can then be excited within this region, without perturbing those in the

bulk solution.

The intensity of the electric field, which has parallel and perpendicular components, is

also related to the angle of incidence and the ratio of the refractive indices [53]:

212

2

0 )/(1cos4

nnE i

−=

θ (1.10)

26

It can be seen from Equation 1.8 that by varying the wavelength, angle and refractive

indices, one can tune the penetration depth [53]. The polarization of the electric field is crucial

as it will determine the amount of light that will leak out as evanescent wave field.

1.3.4 Microarrays

Microarrays are platforms which enable the large-scale and parallel analysis of a large

number of genes (~50,000) simultaneously [55, 56]. Oligonucleotide probes are immobilized in

micron-size elements and are introduced to fluorescently-labeled oligonucleotide targets [56].

The presence of hybridization can then be observed with a confocal fluorescence microscope.

The presence of a fluorescent signal may indicate that the target sequence is complementary to

the probe sequence. A specific application is sequencing by hybridization to oligonucleotide

microchip (SHOM), whereby the sequence of an oligonucleotide target can be determined by

which specific oligonucleotide probe it has bound to [55, 56]. Gene expression and SNP

analysis are also analyzed on a microarray [57]. In this thesis, “macroarrays”, which contain

oligonucleotide spots on the order of millimeters in diameter, are used to conduct parallel

analysis of melt curve data from various targets collected from the same type of surface.

1.3.5 Examples of Optical DNA Biosensors

One of the first evanescent wave optical DNA biosensors was reported by Piunno et al.

[58]. Single-stranded (ssDNA) thymidylic acid icosanucleotides (dT20) were immobilized onto

optical fibers [58]. The fibers were first modified by a silane coupling agent known as 3-

aminopropyltriethoxysilane (APTES) onto which a linker or spacer arm of 1,10-decandiol bis-

succinate was covalently attached [58]. The ssDNA was grown onto the linker base-by-base

27

through solid phase phosphoramidite chemistry [58]. The hybridization event between a

complementary DNA sequence (cDNA) and the immobilized oligonucleotide was detected by

introducing a fluorescent intercalating dye (ethidium bromide) and capturing the emitted

fluorescence within the fiber [58]. The fluorescence intensity was proportional to the amount of

DNA [58, 59]. A detection limit of 86 ng mL-1 of cDNA was reported by using standard

hybridization assay conditions and the analysis time took 46 min [58, 59]. The sensor was able

to remain active and reusable for 3 months using washing conditions based on sonication [58,

59].

A recent development in optical fiber-based DNA biosensor technology is the

incorporation of quantum dots as energy donors in a FRET system that uses fluorophore-labeled

nucleic acid targets [60]. Nucleic acid probes were immobilized onto quantum dots, and the

quantum dots were immobilized onto optical fibers. Using FRET, transduction of hybridization

was achieved as FRET-based emission increased with the concentration of the complementary

target [60]. Selectivity was also demonstrated by introducing the probes on the QD-optical fiber

to non-complementary, three base-pair mismatch, and fully complementary targets [60]. The

limit of detection was reported to be 5 nM [60].

It is clear that the use fluorescence labels can provide various types of information for the

labeled molecule, whether they are static or dynamic. The coupling of TIR with fluorescence

measurements makes optical fibers very sensitive biosensing platforms to detect surface-

localized biomolecules. Moreover, fluorophores can be used on a microarray chip platform,

providing parallel analysis of DNA hybridization and denaturation events on a single surface by

using confocal microarray chip scanners. However, regardless of which type of biosensor is

28

used, good control of oligonucleotide immobilization onto surfaces must be achieved. The next

chapter explores the various methods of oligonucleotide immobilization.

1.4 Strategies for Immobilization of Oligonucleotide Probes An important step in developing a biosensor is selection of a suitable immobilization

strategy for oligonucleotide probes. A variety of different immobilization chemistries can be

chosen, each having advantages and disadvantages, and degree of complexity in terms of

processes. The following will discuss how each immobilization strategy (adsorption, biotin-

avidin affinity pair, thiol-gold interactions, and covalent bonding) can be used to immobilize

oligonucleotides onto various types of substrates. The immobilization of oligonucleotides onto

silica-based substrates by using silane coupling agents will be emphasized as it is the method

used in this thesis.

1.4.1 Adsorption

Adsorption relies on interactions such as van der Waals forces, electrostatic interactions,

and hydrophobic interactions to immobilize oligonucleotides [61]. Electrostatic interactions

involve the negatively-charged phosphate backbone while hydrophobic interactions involve the

nitrogenous bases. The more non-covalent interactions there are, the more stable the adsorption

process becomes [61]. Adsorption can be described by the Langmuir adsorption isotherm, which

can represent relations of the fraction of a surface that is covered by an adsorbent with various

kinetic parameters [62].

E. M. Southern was the first to demonstrate the immobilization of oligonucleotides onto a

surface by adsorption for practical applications [63]. The DNA fragments were separated by gel

29

electrophoresis and were physically transferred to a nitrocellulose membrane filter by letting the

gel come into contact with the filter [63]. The immobilized DNA fragments demonstrated the

ability to hybridize to complementary sequences on the surface [63]. Substrates have included

nitrocellulose, nylon, polystyrene, metal oxide surfaces, and carbon electrodes [62].

Although adsorption is a fast and simple technique for the immobilization of

oligonucleotide probes, it tends to result in low packing densities, restricted probe mobility, and

lack of specific orientation of probe molecules [64, 65]. Furthermore, oligonucleotides can be

easily desorbed from surfaces since the forces involved in adsorption are very weak; application

of heat, washing steps, further chemical steps, changes in pH, and ionic strength can result in

desorption [36]. Therefore, this technique for immobilization is only useful for assembly of

biosensing systems intended for short-term applications in carefully controlled environments

[62].

1.4.2 Biotin-Avidin Affinity Pair

Another non-covalent strategy to immobilize oligonucleotide probes is the biotin-avidin

pair, which represents the strongest non-covalent interaction that is known, having an association

constant of 1015 M-1 [66]. Biotin is a small molecule and a natural vitamin found in living cells

[66]. It serves as a cofactor for carboxylating enzymes [66]. Avidin is an egg white protein [67].

A non-glycosylated form of avidin is known as streptavidin, which is found in Streptomyces

avidinii, a soil bacterium [67]. Neutravidin is a third variant of avidin. It is a non-glycosylated

preparation of native avidin [66]. All three forms of avidin form a tetramer from four identical

subunits; the resultant quarternary structure is stable [67]. Each subunit can bind to one biotin,

30

i.e. each avidin/streptavidin molecule can bind to four biotin molecules [67]. The binding energy

is 330 kJ mol-1 for the tetramer [66].

The extremely strong binding is known to be caused by hydrophobic interactions,

hydrogen bonding, and structural stabilization from a loop in the avidin subunit [66]. The loop

causes the binding pocket to close, thus preventing solvent from penetrating and weakening the

avidin-biotin binding interaction [66]. Due to the combination of these strong interactions, the

biotin-avidin pair can withstand extreme pH (pH 2-13) and temperature ranges, chaotropic salts,

organic solvents, denaturing agents such as 8 M guanidine-HCl, and detergents [66].

1.4.3 Thiol-Gold Interaction

The binding of thiol to gold is a chemisorption reaction. Since Nuzzo and Allara reported

that a monolayer of disulfide molecules could adsorb onto gold in 1983 [68], the thiol-gold

interaction has been extensively explored for many applications including the detection of DNA

by SPR [43] and colorimetric assays based on oligonucleotide-modified gold nanoparticles [69].

The macroassembly of these conjugates was first demonstrated by Mirkin et al. [70]. As a result,

the conjugation of oligonucleotides to gold [66], and to gold nanoparticles as well as quantum

dots has become popular for the development of biosensors.

Self-assembled monolayers (SAMs) are the result of a spontaneous process leading to

organized surface structures or patterns from randomly mobile molecules [66]. The assembly of

thiol-terminated molecules such as alkyl thiols on gold is a result of their binding affinity

towards the metal. Each alkanethiol chain is composed of three components: a head group, a

spacer, and tail group. The head group is the thiol group, which is responsible for binding to

31

gold. The spacer is typically an alkyl chain. The tail group can be any chemical functional

groups such as -CH3, -CF3 (hydrophobic), -COOH, -NH2, OH (hydrophilic), -SH [71], which can

be used for subsequent surface chemical reactions if further surface derivatization is desired.

Alkanethiols are able to self-assemble onto a gold surface via specific chemisorptive and

non-covalent interactions (van der Waals, hydrophobic, electrostatic, and hydrogen/coordination

bonds) [66]. The binding energy of a sulfur-gold bond is ~160 kJ mol-1 compared to tens of

kilojoules per mole for alkyl chain-chain interactions [70]. The balance of the attractive

chemisorption interactions and repulsive forces from intermolecular weak van der Waals forces

and the repulsion of water molecules help stabilize SAMs [66].

A generally accepted picture of ordered SAMs is that the alkanethiol chains are not

standing straight up, but rather possess a tilt angle of 30° [71]. The packing density can also be

influenced by the tail group [72]. The assembly of alkanethiols to a gold surface is accomplished

in two steps: 1) diffusion-controlled adsorption; 2) slow (re-) crystallization process [72]. The

binding of thiol to gold is an exothermic process and is described by the following reaction:

RS-H + Au0n RS-Au+⋅ Au0

n + ½H2 (1.11)

where R is an alkyl group. An electron is transferred from Au to sulfur, making the thiol bond

weak, leading to the dissociation of a hydrogen atom. This thiol bond cleavage occurs at the on-

top site of a gold atom and then the alkanethiol moves into a hollow site, which is a space

defined by three gold atoms in a face-centred cubic (fcc) lattice system, as its final equilibrium

position [71, 73]. The rate of migration of thiolate depends on the dielectric constant of the

solvent [74]. However, there is growing evidence that the sulfur can bind to a gold adatom (a

lone atom adsorbed on the surface) which is adjacent to a hollow site [75]. The hydrogen atoms

32

combine at the metal surface to yield H2, an exothermic step, which drives the adsorption process

of alkanethiols onto the gold substrate [73].

1.4.3.1 Preparations of SAMs

SAMs can be made via a solution or vapor deposition method. In solution deposition,

one only requires a gold substrate and the alkanethiol of interest dissolved in an organic solvent.

Typical solvents used include ethanol, hexane, methanol, benzene, and toluene [66, 71]. The

gold substrate is exposed to the alkanethiol for a period of time. The longer the substrate is

immersed in the solution, the higher the surface coverage will be until saturation is achieved.

Random adsorption is usually triggered by nucleation sites, which are initiated by dust and

particles. However, this usually leads to irreproducible SAMs in terms of surface coverage. For

a highly ordered structure, it usually takes about 12–15 hours for an alkanethiol concentration of

micro- to millimolar range [66]. Multilayers do not form for alkanethiols with more than 5

carbons, but they can be formed if the immersion time is as long as 6 days [72]. To improve

reproducibility, vapour deposition can be used. The alkanethiol solution is usually transported

into a closed ultra-high vacuum (UHV) chamber by nitrogen gas as a carrier [66]. This method

allows precise control of environmental conditions, but it is more difficult to achieve higher

packing densities [76]. There are ways to reduce the amount of contaminants. Ion sputtering,

oxygen-rich flame annealing, and chemical etching with Piranha solution, which is composed of

hydrogen peroxide and sulfuric acid, are common methods to clean the substrates [66].

Oxidation of sulfur leads to surface binding of the thiolate ion to gold as the following reaction

proposed by Weisshaar et al. demonstrates [77]:

Au + X (CH2)nS- → AuS(CH2)nX + 1e- (1.12)

33

This method provides similar structures and interfacial properties of SAMs compared to those

formed from wet deposition [66]. The improvement is in the repulsion of contaminants,

resulting in denser films with fewer defect sites [66].

There has been a growing interest in making mixed SAMs, i.e. SAMs comprised of two

or more types of alkanethiols. It was found that the mole ratio of two components in the loading

solution was the same as the ratio found within the immobilized mixed SAMs [72]. Mixed films

can be used to better control surface wetting behavior, chemical reactivity, and biological

response [78], therefore offering potential for new electronic and chemical properties, nanoscale

structure, and new applications [79] such as microelectromechanical systems (MEMS),

molecular switches, and biosensors, which depend heavily on interfacial characteristics [80].

Recently, mixed SAMs composed of thiol-modified oligonucleotides and short alkanethiol

chains have been found to enhance DNA hybridization efficiency because shorter alkanethiols

cause the oligonucleotides to adopt an upright position, instead of interacting with the surface at

multiple non-specific attachment sites [81, 82]. It is clear that mixed films offer opportunity to

better control surface chemistry through the engineering of wettability, dynamics and

morphology, and this represents a major theme in this thesis.

1.4.4 Covalent Linkages

Covalent linkages provide for coupling of reactive groups that result in the most stable

bonding to surfaces. Common methods to covalently immobilize oligonucleotides onto silica-

based substrates are often based on the fact that hydroxylated surfaces can be derivatized to yield

electrophilic or nucleophilic groups to react with chemically modified oligonucleotides [83].

Unlike immobilization by adsorption, packing density and strand orientation can be readily

34

controlled for covalently-immobilized oligonucleotides, which can greatly improve kinetics of

hybridization [36, 66]. Covalent linkages are also more stable in stringent washing conditions,

and extreme pH and temperature conditions [66]. However, procedures to form covalent

linkages can become complicated and often multiple procedural steps are required [36].

The earliest demonstrations of covalent immobilization of nucleic acids to solid supports

was demonstrated by Gilham and co-workers in 1964 [84], who used a phosphodiester linkage

between a hydroxyl group from cellulose and the phosphate group from the 5′-end of DNA for

the immobilization. Many other reactions have been demonstrated. For example, Kremsky et al.

immobilized thymidine onto cellulose by using dicyclohexylcarbodiimide to form stable

carbodiimide linkages [85].

The immobilization of oligonucleotides onto silica-based optical biosensors is the focus

of this work, and it usually involves the activation of silanol surfaces with alkoxysilanes.

Furthermore, important aspects of linker and oligonucleotide immobilization will be discussed.

1.4.4.1 Cleaning Protocols for Silica-Based Substrates

Metal and organic contaminants are prone to adsorb onto surfaces. Before any surface

derivatization, the surface must be free of these surface-adsorbed contaminants. There are

numerous cleaning protocols for cleaning silicon and silica surfaces. Two of the more common

ones are cleaning with Piranha solution and the “RCA method” [86]. The first method, as the

name implies, is an extremely harsh and dangerous mixture of 30% hydrogen peroxide and 98%

sulfuric acid in a 1:3 volume ratio [87]. The Piranha solution simultaneously eliminates organic

and metal contaminants. The RCA method is considered much safer than the Piranha solution,

35

and involves two cleaning steps: 1) an alkaline cleaning solution composed of 1:1:5 30%

hydrogen peroxide:30% ammonium hydroxide:water, which is followed by 2) an acidic cleaning

solution composed of 1:1:5 30% hydrogen peroxide:concentrated hydrochloric acid:water. The

alkaline wash eliminates any organic matter by the solvation power of ammonium hydroxide and

the oxidizing ability of peroxide [88]. Ammonium hydroxide can also complex with Groups I

and II metals [88]. Hydrochloric acid further rids any heavy metal contaminants from the

surface by forming soluble complexes [88]. The result of these cleaning protocols is the

generation of a layer of silanols on the surface, which is typically less than 50 Å thick [86, 88].

1.4.4.2 Surface Activation Using Silane Coupling Agents

To immobilize oligonucleotides, surface silanol groups are reacted with different

alkoxysilane coupling agents such as (3-glycidyloxypropyl)trimethoxysilane (GOPS) and (3-

aminopropyl)triethoxysilane (APTES) to yield terminal reactive epoxy and amine groups,

respectively. GOPS, along with APTES, have been used since the 1970’s for the immobilization

of enzymes and antibodies on a wide range of solid supports including fused silica [64]. Surface

epoxide groups can react with amino- or thiol-modified oligonucleotides [12]. Alkylsilanes can

also be used to activate silica surfaces. Chemically modified oligonucleotides can be

immobilized on amino-functionalized surfaces via the use of homo or heterobifunctional linkers,

which possess identical or different terminal functional groups.

The alkoxysilane groups immobilize onto surface silanol groups through siloxane bonds

[83]. Whether the silane coupling agent carries one, two, or three alkoxysilane group(s), the

mechanism of attachment to the surface is identical. However, reactivity decreases with

36

decreasing number of leaving groups, which are typically methoxy or ethoxy groups [66].

Trialkoxysilanes are the most reactive species.

The first step in the functionalization of the surface is the condensation of the

alkoxysilane groups to the silanol groups using a base such as Hunig’s base (N,N-

diisopropylethylamine) as a catalyzing reagent in a non-polar solvent [89]. Methoxy groups

attached to Si can be released easily in the presence of water to form methanol and silanol, which

proceeds by the SN2 mechanism [90]:

≡ SiOMe + H2O ≡ SiOH + MeOH (1.13)

where ≡ represents the three alkyl and/or alkoxy groups that Si is singly-bonded to. Silanols can

condense easily with surface silanols, with each other or with another methoxy group that is

attached to another Si atom to form siloxane bonds [90]:

≡ SiOH + ≡ SiOH ≡ Si-O-Si ≡ (1.14)

≡ SiOH + ≡ SiOMe ≡ Si-O-Si ≡ (1.15)

When more than one alkoxygroup exists, it is possible for multiple surface attachments to take

place [83]. However, it has been determined that the dominant binding arrangement of APTES

is having two ethoxy groups condensing with surface silanol, while the third ethoxy group

remains free [83].

An oligomerized network of silanes can occur before and after surface attachment, and

can subsequently form a multilayer structure via the polymerization of siloxane bonds [90],

especially in the presence of water. For example, a 0.25% aqueous solution of GOPS may

induce a multilayer formation involving 8 molecular layers [91]. Therefore, attempts have been

37

made to minimize polymerization of silanes so that the result is a homogeneous monolayer

versus a heterogeneous polymerized surface [83]. A homogeneous surface is desired for the

development of a biosensor, affording potential for better reproducibility, selectivity, sensitivity

and kinetics of hybridization. AFM studies of GOPS-derivatized silica-based substrates have

shown a coverage of GOPS nodules of approximately 50–100 nm in diameter [92]. Immobilized

oligonucleotides on nodules of ~50 nm may experience improved hybridization rates as

compared to being immobilized on a flat film surface [92]. This may be due to the influence of

geometry on surface density of probes (a fanning effect as encountered in polymer brushes) [92].

Silane polymerization can be ameliorated by: 1) minimizing the amount of alkoxysilanes

introduced to a surface by using vapor deposition, anhydrous solvents, oven-dried glassware,

monoalkoxysilanes, and postsilanization curing [83].

Monoalkoxysilanes may seem like an ideal coupling agent but their ease of hydrolysis

which leads to surface detachment is not practical [93]. Dialkoxysilanes are known to

polymerize like trialkoxysilanes, and be hydrolyzed like monoalkoxysilanes [83]. Therefore, the

silane coupling agents of choice are typically the trialkoxysilanes. Postsilanization curing has

been shown to be effective in reducing hydrolysis by cross-linking free silanols [94, 95],

however, it still allows the growth of heterogeneous silane layers [83].

The complexity of interactions increases when using agents such as aminosilanes, for

example, (3-aminopropyl)trimethoxysilane (APTMS). Such agents can chemically react with

free surface silanols. According to Kowalzyck et al., Horr et al., and Chiang et al. [96-98],

besides reacting with the surface via siloxane linkages, the amino-terminated end of

aminosilanes can react with the surface due to protonation either at a silanol site or by other free

silanols [99]. Further possibilities include the formation of siloxane bonds, as well as hydrogen

38

bonding between the primary amine and silanol on a surface. Postsilanization capping with an

alkylsilane has indicated an increase in the immobilization efficiency of aminosilanes due to the

reduction of side reactions involving the amine groups [100]. Hicks and Jones have also used

alkylsilanes to cap unreacted silanols to prevent undesired side reactions [101].

Despite problems of irreproducibility of structures, alkoxysilanes are routinely used due

to the straightforward immobilization procedure, low cost, and availability of numerous silanes

with different end functional groups (thiol, amine, epoxide, and halogenated) that are suitable for

subsequent surface modifications. Du et al. described a method in which thiolated

oligonucleotides were conjugated to mercaptosilanes via a disulfide bond, producing silanized

oligonucleotides, which can be immobilized directly on unmodified glass surfaces [102]. The

silanized oligonucleotides were found to be immobilized at a density of 2 ×105 molecules μm-2

and were capable of surviving 3-4 rounds of hybridization and denaturation of target sequences

[102].

Another immobilization method based on alkoxysilanes is found in the work by Dendane

et al., in which they describe a method to pattern oligonucleotides inside a glass capillary by first

immobilizing an aminoxy-derivatized triethoxysilane, which is protected by a photolabile

protecting group [103]. Aldehyde-modified oligonucleotides were immobilized via the

formation of an oxime bond with the deprotected aminoxy groups upon UV irradiation [103].

Steinberg and co-workers explored several different immobilization chemistries which

can attach amino- or thiol-modified oligonucleotides to aminosilanized surfaces [104]. It was

determined that cyanuric chloride, diisothiocyanate, nitrophenyl chloroformate, and hydrazone

chemistries provided the highest hybridization efficiencies amongst the chemistries explored

[104]. A probe density of 2.1 × 104 probes μm-2 or a mean distance of 69 Å between two

39

adjacent probes was indirectly determined from a deprotection reaction of

fluorenylmethyloxycarbonyl (Fmoc), a protecting group for amines [104].

Walsh et al. have shown that aminosilane-derivatized bead surfaces can be further

reacted with acetic anhydride, polyethylene glycol (PEG)-dicarboxymethyl, and 1,4-phenylene

diisothiocyanate to form carboxyl ends (for the first two reagents), and thiocyanate (for the last

reagent) [105]. Amino-modified oligonucleotides were then linked to the carboxylated surfaces

using common activating reagents such as 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide)

(EDC) and sulfo-N′-hydroxysuccinimide (NHS) to facilitate the linkage between carboxyl and

amine [105]. EDC promotes the formation of an activated ester intermediate, which can be

attached to primary amines to form a peptide bond under neutral conditions [105]. With the

addition of sulfo-NHS, a stable sulfo-NHS ester intermediate is formed to react with primary

amines under slightly acidic conditions [105]. Isothiocyanates, on the other hand, can react

readily with primary amines under basic conditions, forming a new C-N bond. The one-step

EDC immobilization procedure resulted in the highest immobilization efficiency of 82-89%

[105].

The work by Watterson and co-workers [37] demonstrated the compatibility of

automated oligonucleotide synthesis with oligonucleotide immobilization. First, optical fibers

were activated with GOPS and were further reacted with dimethoxytrityl hexaethylene glycol

(DMT-HEG) [37]. DMT was used to ensure that the reaction only occurred at one end of HEG

and avoided formation of closed loop structures with unreacted epoxide sites on the surface [37].

Chlorotrimethylsilane (TMS⋅Cl) was used to cap any unreacted silanols and hydroxyl groups

[37]. Stepwise oligonucleotide synthesis was performed directly on the fibers by using

phosphoramidite chemistry carried out by an automated DNA synthesizer [37]. A mean

40

separation distance of 263 Å for a low density of oligonucleotide probes and 100 Å for a high

density were achieved [37].

1.4.4.3 Control of Packing Density

The coupling efficiency of the chosen immobilization strategy must be considered as it

can affect the immobilization yield and packing density, which ultimately influences the

hybridization efficiency. Beattie and co-workers have found that when oligonucleotide probes

were closer than ~5-10 nm apart, hybridization efficiency was poor [106]. This high packing

density triggered steric hindrance and charge repulsion amongst oligonucleotide targets [66].

However, if the packing density is too low, then not enough hybridization would occur to

develop a useful analytical signal [66]. An abundance of unmodified sites may lead to increased

levels of non-specific adsorption as well [66]. The packing density itself is affected by the

availability of functional groups and the coupling efficiency of end functionalized probes [66].

Therefore, judicial choice of immobilization chemistry is paramount.

One way to maintain a high density but reduce steric hindrance is to use dendrimers as

shown by Hong and co-workers [107]. Dendrimers are highly branched polymers with uniform

size and molecular weight with a three-dimensional structure (elliptical- or cone-shaped) with

terminal functional groups [107]. It was shown that the immobilization of oligonucleotides can

be spaced out by using bulky dendrimers as anchor points, with the resulting surface density of

amine sites being 0.1-0.2 amine per 100 Å2 [107]. Others have used linear and star-shaped

molecules as well [66]. Another strategy is to use photochemistry. Piunno et al. used a

photolabile protecting group to protect HEG linkers and to limit the amount of oligonucleotides

being immobilized [18]. Controlled deprotection by UV light was used to selectively cleave

41

some portion of the photoprotecting groups. The remaining HEG linkers were deprotected and

were further reacted with non-nucleic acid oligomers [18]. Hicks and Jones used benzaldehyde

to protect APTMS [101]. This method also caused APTMS to be immobilized at reduced

density and avoided undesired polymerization reactions [101]. Fluorescence spectroscopy

experiments which employed pyrene as a marker showed that the distance between adjacent,

bulkier trityl-capped amine sites were, on average, longer than the less bulky benzaldehyde-

capped APTMS [108].

1.4.4.4 Linkers and Oligonucleotide Immobilization

No matter which method is chosen to immobilize oligonucleotides, one must ensure that

the probes are freely available and mobile to undergo hybridization with target strands. One

obvious strategy is to immobilize the probes through the 3′ or 5′-end of the DNA strand to free

up the nitrogenous bases for hybridization [64]. However, instead of attaching the terminal end

of the oligonucleotide directly to the surface, a linker is usually attached between the

oligonucleotide and the surface to provide further mobility and a fluid-like environment to mimic

a real hybridization environment for the oligonucleotide [64]. It has been reported that

interfacial hybridization is 10-fold slower than solution-phase hybridization [64]. Therefore, one

must choose a covalent immobilization method which can optimize hybridization reaction

kinetics and efficiency [64].

The properties of a linker that must be considered are length, charge, hydrophobicity,

flexibility, degree of solvation, and absence of hydrogen bond donors [64]. These characteristics

can dramatically influence the efficiency of DNA hybridizations. It was found that a linker

length of at least 28 Å long would be needed to improve the rate of hybridization to about 4

42

times compared to oligonucleotides that were directly attached to a solid substrate [109].

Furthermore, it was determined that the length of the linker had a greater impact than packing

density on the hybridization yields [110]. The hydrophobicity of linkers should also be minimal,

otherwise linkers would form aggregates in an aqueous environment, thus resulting in an

inhomogeneous coverage [64]. Moreover, hydrophobic linkers would offer sites for non-specific

protein and lipid adsorption [64]. Hydrophilic linkers were shown to work quite well for the

attachment of oligonucleotides, but negatively-charged linkers are not desired as they may repel

incoming negatively-charged target DNA strands [64]. Positively-charged linkers should not be

used as they would electrostatically attract target oligonucleotides and impede or block

hybridization [64]. It was found by Ganachaud and co-workers that the optimal properties of a

DNA linker include a low charge density and significant hydrophilicity [111]. Examples of

useful hydrophilic linkers are HEG, pentaethylene glycol, or ethylene glycol [64]. It is known

that they can form hydrogen bonds with water in a highly organized manner which minimizes

the tendency for hydrophobic interactions [64]. The advantage is that a monolayer of these

molecules provides a relatively fluid and highly hydrated state with sufficient flexibility and

mobility to allow hybridization of immobilized oligonucleotides [64]. Furthermore, such surface

chemistry minimizes protein adsorption, which otherwise would have caused drift in calibration

and loss of sensitivity due to surface blockage [64].

Of all these immobilization methods, covalent linkage is the most practical choice for

stable immobilization such that the yield of oligonucleotide immobilization and hybridization

efficiency can be maximized. The use of biotin-avidin, even though this non-covalent linkage is

extremely strong, may still run the danger of detachment of biotinylated-oligonucleotides from

denatured avidin in experiments that require high temperature conditions, such as a melt curve.

43

In this thesis, APTMS was chosen to activate silica-based substrates for its chemical

compatibility with the substrate and its high reactivity. Using the method proposed by Hicks and

Jones, a mixture of benzaldehyde-capped and unprotected APTMS served as the first

immobilization step towards the goal of this project, which was the preparation of the mixed

film. The benzaldehyde-cap served to minimize interactions between neighbouring APTMS

molecules and APTMS molecules with the surface, and also protected the amine group in

APTMS for subsequent immobilization of the oligomer portion of the mixed film.

1.5 Atom Transfer Radical Polymerization Immobilization of entire polymers to surfaces through end-functionalized groups, which

is often called a “grafting to” method, cannot yield a high surface density due to steric hindrance

[112]. A more effective way to graft polymer brushes of high density onto surfaces is called a

“grafting from” method or surface-initiated polymerization, in which immobilized initiators are

used to grow the polymer chains from the substrate one monomer at a time. The method chosen

here to synthesize oligomers using the “grafting from” technique is called atom transfer radical

polymerization (ATRP). ATRP is a specific type of controlled/“living” polymerization which

can carry out such “grafting from” method. A general introduction of controlled/“living”

polymerizations, the theory behind ATRP and how it has been applied to the biosensing field

will be elaborated.

44

1.5.1 Controlled/“Living” Polymerizations

Controlled/“living” polymerization is a term given to polymerization reactions that may

possess both controlled and “living” characteristics, but since the nature of this polymerization

system is still ambiguous and uncertain, the dual definition is usually given [113]. In general,

radical polymerization reactions tend to be uncontrollable in terms of molecular weight and

polydispersity, which is defined by Mw/Mn, where:

Mw is the weighted average molecular weight:

)()( 2

ii

iiw MN

MNM

ΣΣ

= (1.16)

where Ni is the number of polymers of molecular weight Mi. Mn is the number average

molecular weight:

i

iin N

MNMΣ

Σ=

)( (1.17)

where the total weight (ΣNiMi) is divided by the number of polymers (ΣNi). For example, ionic

polymerizations tend to succumb to termination due to the presence of impurities [114].

Conventional radical polymerizations are often uncontrollable due to disproportionation,

coupling, and premature termination reactions [114]. Disproportionation often leaves saturated

and unsaturated ends whereas coupling leads to a single dead end with no further reaction. The

following chemical equations describe the coupling, disproportionation, and termination reaction

often observed in radical polymerization:

Coupling

R• + T• R-T (1.18)

45

Disproportionation

R-T R• + T• (1.19)

Termination

R• + R• RR (1.20)

where R• and T• are two distinct radical species, and R-T is the product of two radical species R•

and T•.

Living systems require “active” and “dormant” species. “Active” species are capable of

undergoing polymerizations while “dormant” species are inactive from doing so [114]. Active

species are present at only part-per-million concentration while most species are dormant [114].

This results in less termination reactions [114]. Terminations might still occur, but it occurs in

less than 10% of all reactions [114]. However, termination reactions cannot be completely

eliminated in radical polymerization because they involve the same active radical species as the

propagation reaction [114]. There are also no irreversible chain-breaking reactions in “living”

systems [113].

Controlled polymerization simply means obtaining polymers of predefined molecular

weight, low polydispersity, and controlled functionality [113]. Transfer and termination can still

occur but are less dominant factors [113]. Table 1.3 compares the definition of “living” vs.

controlled polymerizations.

46

Table 1.3: Distinct characteristics of living and controlled polymerizations.

Living polymerization Controlled polymerization

Chain growth Chain or step growth

Slow initiation possible Limited chain breaking possible

Slow exchange possible Fast initiation

Uncontrolled MW possible Fast exchange

High polydispersity possible *Controlled MW

*No chain breaking (transfer or termination) *Low polydispersities

denotes key feature(s) of each polymerization. Adapted from Matyjaszewski [113].

As Table 1.3 indicates, the advantages of controlled/ “living” polymerization are defined by

molecular weight and low polydispersity. Polydispersity is usually less than 1.5 [115] and this

value is small compared to conventional cationic and radical systems [115].

There are three types of polymerization mechanisms that yield controlled/“living” radical

polymerizations. These are: stable-free radical polymerization (SFRP) or nitroxide-mediated

radical polymerization (NMRP); atom transfer radical polymerization (ATRP); and reversible

addition-fragmentation chain transfer (RAFT). Table 1.4 describes the mechanism and

monomer(s) used for each type of polymerization [114].

47

Table 1.4: Types of controlled/ “living” radical polymerization.

Mechanism Propagating radical/regulator Monomers used

SFRP or

NMRP

Reversible homolytic cleavage of weak covalent bond leads

to propagating radical and stable free radical, which only

reacts with propagating nitroxide radical called 2,2,6,6-

tetramethyl-1-piperidinyloxy (TEMPO)

styrenes

ATRP Reversible redox reaction between alkyl halides and

transition metal complexes

Acrylates, methacrylates

RAFT Degenerative chain transfer with alkyl iodide or thioesters thiocarbonylthio compounds,

such as didthioesters,

dithiocarbamates,

trithiocarbonates, and

xanthates

The key characteristic of these reactions is the extension of radical lifetime from 1 s to a few

hours by making use of a dynamic equilibrium which involves a rapid exchange between

dormant and active states [114]. A persistent radical (deactivator) species or a highly active

transfer agent is often used to react with propagating radicals, which then become dormant, with

a rate kd [114]. Active species are regenerated by activating the dormant species by the

corresponding mechanism with a rate ka [114].

1.5.2 Atom Transfer Radical Polymerization

The ATRP reaction spun off from the atom transfer radical addition (ATRA). This was

modified from the Kharasch reaction [116], which used light or conventional initiators to

generate radicals [114]. Matyjaszewski [117, 118] and Sawamoto [119, 120] have separately

48

reported this new type of controlled/ “living” polymerization. ATRP has been used to make

numerous types of polymer topologies such as stars, combs, and branched shapes [116]. It can

also be used to make block, gradient, alternating and statistically-based polymers [116]. The

polydispersity is typically in the range of 1.1–1.5 [115]. In order to generate such narrow

polydispersity, the ATRP reaction must have: 1) a fast initiation compared to propagation, and 2)

minimal side reactions [116]. The following first discusses the mechanism of ATRP, and then

how each component (initiator, monomer, catalyst, solvent) of the ATRP reaction can affect its

success.

ATRP is capable of polymerizing various acrylate- and methacrylate-based monomers to

achieve a narrow polydispersity index with controlled molecular weight [117]. Controlled/

“living” polymerization is achieved by having a fast dynamic equilibrium between a higher

concentration of dormant species (Pn-X) (rate of deactivation is 107 M-1 s-1) than of active species

(Pn•) (rate of activation is 1 M-1 s-1, or at parts-per-million concentrations [114, 116]), and is

described by the following reactions:

Pn-X + Cu(I)/2L Pn + Cu(II)X/2L• (1.21)

Pn• + monomer Pn+m• or Pn• (1.22)

where Cu(I) abstracts the halogen atom (X) from the initiator or dormant polymer chain (Pn-X)

and causes a homolytic cleavage of the halide-carbon bond, leaving a radical at the terminal end

of the polymer chain. Copper(I) becomes oxidized to copper(II) upon complexing with the

halide and ligands (L), which are used to adjust the strength of the catalyst [115]. Wherever a

radical alkyl or polymer chain is available, an acrylate or methacrylate-based monomer can be

49

added across the double bond of an alkene at the end of this chain, thus propagating the radical to

the end of this new monomer. The increase in Cu(II) will deactivate the polymerization process

and revert the active species (Pn•) back to its dormant state (Pn-X) [114]. Cu(II) is then oxidized

back to Cu(I) [114]. This reversible reaction is key to the controlled polymerization in ATRP.

The ATRP reaction is oxygen sensitive. Therefore, the monomer/catalyst mixture is

usually purged with an inert gas. Although any residual O2 can be scavenged by the metal

catalysts [116], this may lower the amount of catalysts available for polymerization, hence

affecting the polymerization rate [116].

ATRP initiators are typically alkyl halides with activated substituents on α-carbon such

as aryl, carbonyl, or allyl groups [115]. R groups on initiators are present to stabilize the radical

[115]. The order of stabilizing groups is: CN > C(O)R > Ph > C(O)OR > Cl > Me [116]. α-

halobutyrates, which were used in this work, can become initiating radicals more readily than

propionates because the former radical is more stable upon the halogen abstraction step [116].

Tertiary groups, by the same reasoning, are better stabilizing groups than secondary groups,

which are preferred over primary groups [116]. The leaving group, X, should be as labile as that

in dormant species [115]. The order of bond strength is R-Cl > R-Br > R-I. Therefore, R-Cl

would be the least efficient initiator whereas R-I would be the most efficient [116]. However,

the resultant metal iodide complexes tend to be unstable and light sensitive [116]. It was found

that the use of bromoderivatives as initiators and dormant species as well as the use of transition

metal chlorides as catalysts tend to accelerate the initiation process [115].

ATRP can be done in neat conditions, in solution, or in a heterogeneous system such as

an emulsion or suspension [116]. If the polymer is insoluble in its monomer, a solvent can be

used [116]. The type of solvent may affect the structure of the catalyst, and its participation in

50

atom transfer [115]. In general, ATRP can be done in water and various organic solvents such as

toluene, ethyl acetate, acetone, dimethyl formamide (DMF), and alcohol [116]. To achieve a

successful ATRP reaction, there should not be any radical chain transfer to the solvent [116].

Moreover, any interactions between the solvent and catalyst which may reduce catalytic activity

should be avoided [116]. For example, it was reported that HX was easily eliminated from

polystyryl halides in a polar solvent [116]. This is a solvent-assisted side reaction. Another

issue to address is whether the solvent would change the structure of the catalyst [116].

There is a wide variety of transition metal catalysts that can be used in ATRP based on

copper, ruthenium, and nickel [116]. The choice of transition metal is dependent on the

following recommendations, according to Matyjaszewski [116]:

1. The metal center should have two readily accessible oxidation states.

2. The metal center should have adequate affinity toward a halogen.

3. The coordination sphere around the metal should be expandable upon oxidation to

accommodate a halogen.

4. The ligand should complex the metal relatively strongly.

5. The oxidized metal should be able to rapidly deactivate propagating polymer

chains and convert them into their dormant states.

Thus far, copper has been used extensively as it is versatile for numerous systems and it

is inexpensive [116]. About 80% of publications that report ATRP make use of copper [116].

Cu(I) assumes a tetrahedral or square planar configuration, and complexes with one tetradendate

or two bidendate ligands [116]. Cu(II) forms a trigonal bipyramidal structure with one

tetradendate or two bidendate ligands, such as 2,2′-bipyridyl (bpy) [116]. Figure 1.11 shows the

complexed structures of bpy and copper in its reduced and oxidized states:

51

Fig. 1.11: The ligand-copper structures at reduced and oxidized states in ATRP. Adapted from Matyjaszewski

[116].

Ligands are added to adjust the strength of the catalyst(s) by influencing the redox

potential and halogenophilicity of the metal center [115, 116]. Ligands are also used to

solubilize the transition metal in non-polar solvent [116]. They should bind strongly to the metal

and allow the coordination sphere to expand for metal oxidation and accommodation of a

halogen [116]. For copper-based ATRP, as used in the work of this thesis, nitrogen ligands such

as bpy offer the best performance [116]. Other ligands such as sulfur, oxygen, phosphorus

ligands are not as compatible due to undesired electronic effects and unfavourable binding

constants [116]. Not only are electronic effects important, but so are steric effects of ligands

[116]. Any increase in steric effects may reduce catalytic activity or efficiency [116]. The

activity of nitrogen-based ligands also decreases with the number of coordinating sites [116].

Notably, the activity is greater for bridged and cyclic ligand compounds than for linear ones

[116]. One must bear in mind that ligands may participate in side reactions, which may affect

the success of the ATRP reaction [116].

More reactive monomers are known to form less reactive radicals due to the resultant

unstable radical structure [116]. Although radicals do not have charge, it is possible that

substituent groups can influence the radical region to be nucleophilic or electrophilic [116]. One

52

must be cautious when using acidic monomers as they have been found to reduce the activity of

the catalyst by coordinating to the transition metal as well as protonating any nitrogen-containing

ligands [116]. The consequence of the latter is reduced ability of the ligand to complex with the

metal [116].

Temperature plays a major role in the production of radicals. As it increases, the radical

propagation rate constant increases, which also leads to an increase in ATRP rate [116].

However, this also means that chain transfer and side reactions will also increase [116].

Secondly, the solubility of a catalyst may be improved by raising the temperature. However, the

rise in temperature may induce its decomposition [116]. Overall, the temperature should be

chosen based on the type of monomer, catalyst, and the targeted molecular weight [116].

Termination reactions occur to an extent of no more than 5% in the initial polymerization

stage [116]. The small percentage is due to the presence of oxidized metal complexes or

deactivators [116]. Upon termination, the bromide group can be easily converted if required by

nucleophilic substitutions to numerous types of functional groups such as azide, amine, and

hydroxyl for further reactions [115]. In contrast to ATRP, further derivatization of the terminal

propagating species in SFRP and RAFT is more complicated as nitroxide and dithioesters are

more difficult to displace [114].

1.5.3 Application to Biosensors

Surface-initiated ATRP has been used in numerous interfacial applications such as the

modification of quantum dots, nanoparticles and microbeads, protein adsorption studies, and

improvement of biocompatibility by means of surface coatings [121]. ATRP is now being used

53

in the biosensor area to improve biosensing capabilities and minimize non-specific adsorption.

One example of the application of ATRP in the development of a DNA biosensor was

demonstrated by Lou et al. [122]. They designed a DNA sensor on gold substrates based on the

positive visual appearance of polymer, synthesized via ATRP, which can amplify a signal based

on DNA hybridization [122]. The strategy involves immobilizing thiol-modified oligonucleotide

probes onto gold surfaces via the sulfur-gold linkage. The surface was then exposed to target

sequences. Only the fully complementary target carried the initiator which was able to start

ATRP, whereas mismatched oligonucleotide sequences did not carry the initiator. ATRP was

performed in two stages. The first resulted in a linear polymer of poly(2-hydroxyethyl

methacrylate) (PHEMA) and the second ATRP reaction polymerized 2-hydroxyethyl

methacrylate (HEMA) on the existing PHEMA chain by immobilizing the initiator onto the

hydroxyl group of the first PHEMA chain. Consequently, only the oligonucleotide probe areas

that were introduced to the fully complementary targets had a thick film of polymer, which could

be visualized by eye. Thus, ATRP was used as an amplification strategy to signify the presence

of fully matched oligonucleotide sequences. In another paper by Lou and He [123], they

suggested that the presence of DNA accelerated the ATRP rate. This could be beneficial because

one of the ideal requirements of a biosensor has always been speed of response. It was suggested

that the negatively charged DNA backbone may concentrate more positively-charged copper

catalysts at the surface, thus enhancing the rate.

Researchers have turned to ATRP in the search for non-fouling surfaces. In a paper

reported by Bernards et al. [124], it was found that a mixture of two oppositely charged

monomers produced a film that was resistant to protein adsorption. Proteins such as fibrinogen,

lysozyme, and bovine serum albumin were tested by surface plasmon resonance and it was

concluded that statistical copolymer provided non-fouling properties for all three proteins. This

54

work also explores the effect of a mixed film grown by ATRP to reduce non-specific adsorption

of oligonucleotides.

Although ATRP promises narrow polydispersities (unless the polymer chains are cleaved

from the surface), the actual values cannot be readily determined for surface-initiated ATRP.

Nevertheless, the narrow polydispersity originates from the absence of chain breaking reactions.

The ATRP mechanism is well understood such that the type of initiator, catalyst, solvent, and

monomers can be chosen to optimize the reaction. Furthermore, this method has the potential to

grow oligomers in the presence of oligonucleotides. Therefore, in this work, ATRP was selected

to grow the oligomer portion of the mixed film in the presence of immobilized oligonucleotides.

1.6 Contributions of this Thesis

The goal of this project is to improve the selectivity of hybridization to better distinguish

SNPs when using a typical optical DNA biosensor. The discrimination between fully

complementary (FC) and SNP targets is not trivial even at the same temperature, ionic strength,

and pH conditions because of similar thermal stabilities [16] and a low resolution between them

on surfaces due to the presence of non-specific adsorption and interactions. The difference in

stabilities also depends on the location of the base-pair mismatch and length of the target

sequence [16].

There have been numerous strategies proposed to achieve improvement of selectivity.

Although the types of transducers may be different, the theme of signal amplification and an “all-

or-none” response are used to improve selectivity for fully complementary over SNP targets.

For example, Taton and co-workers have explored the reduction of silver ions on gold

55

nanoparticles which were tagged onto oligonucleotide targets to improve the selectivity for FC

over SNP targets [125]. According to Fig. 2 of Taton’s paper, the full width half maximum

(FWHM) was decreased by at least six fold when nanoparticle-based reporter probes were used

compared to fluorescein-labeled ones [125]. Inouye et al. presented an electrochemical strategy

involving the attachment of a ferrocene-modified nucleoside which would render hybridized

duplexes immobilized on gold surfaces conductive during a cyclic voltammetry experiment

[126]. There was weak or no oxidation signal of the ferrocene-nucleoside for the SNP targets

because the DNA wire was broken at the SNP site [126]. Li and co-workers presented a method

based on rolling circle amplification (RCA) to amplify the chemiluminescence intensity for FC

targets [127]. Zhong et al. used sandwich assays based on the presence or lack of a ligated DNA

sequence to distinguish between the two targets [128]. These strategies (except for the work by

Taton et al. [125]) are only useful if one needs to detect fully complementary targets in the

presence of SNP targets. However, in medical cases where both wild- (perfectly matched) and

mutant-type (SNP) genes must be distinguished from each other, such as the genes responsible

for spinal muscular atrophy (SMA), there must be adequate resolution between the signals that

represent each DNA target.

The method presented in this work can discriminate between different DNA targets by

increasing the slope of melt curves and increasing melting temperature differences. This can be

done by controlling the orientation and interactions of the hybridized duplexes on surfaces. Our

group has previously determined that selectivity of DNA hybridization is governed by its

environmental conditions such as local charge density, which is contributed by the presence of

DNA, in addition to temperature, ionic strength, and pH [129]. To achieve selectivity during a

denaturation experiment in which the charge density decreases due to denaturing DNA targets,

one must have control of the environment such that structural regularity of DNA hybrids remain

56

similar, hence retaining a narrow distribution of surface free energies [18]. The control of DNA

probe orientation with respect to the surface has largely been done on conductive surfaces where

an electric field can be applied, causing the negative backbone of DNA to extend perpendicularly

from the surface by electrostatic repulsion, resulting in optimal conformation to increase

hybridization efficiency [130]. Alternatively, mixed films composed of thiolated

oligonucleotides and a diluent such as thiolated oligoethylene glycol and alkanethiols which are

self-assembled on gold surfaces have shown enhanced hybridization efficiency [81, 131, 132]

and reduced adsorption [81, 131]. Lee and co-workers have demonstrated by using near-edge x-

ray absorption fine structure that nucleobases were more parallel to the surfaces in mixed DNA

and ethylene glycol films than in pure oligonucleotide films [131].

On optical substrates, however, direct immobilization of functionalized biomolecules is

not possible without a surface activation step. Then, immobilization of the two components

using the same chemistry may not yield the ratio desired to achieve uniform conformation of

immobilized oligonucleotides. The work of Piunno and co-workers proposed a photochemical

strategy to prepare mixed films on optical fibers and investigated the effect of having a diluent

on the melting temperature (Tm) [18]. This was the first study which demonstrated that the

presence of charged, non-nucleic acid oligomers altered the local environment of oligonucleotide

probes by decreasing the Tm [18]. Although the mixed film was successfully prepared, the

syntheses and characterization of photolabile protecting groups add a level of complexity to the

procedure since special care must be taken to prevent intermediates and reactants from absorbing

any light which can initiate the photochemical reaction before actual use [133].

There were numerous strategies that were attempted to make the mixed films for this

thesis. The first strategy was the photochemical approach as proposed by Piunno et al. [18], but

57

synthesis of the photolabile group proved to be too inconvenient, difficult, and time consuming.

The specific personal challenges involved isolating a pure product from each reaction step, doing

a purification step in a dark room and ensuring that the intermediates of the photoprotecting

group is not exposed to light. This strategy was not pursued. Another approach which was

attempted was the use of dendrimers, which are highly branched polymers that grow from a

single apex or core [107]. One can use them to space out molecules, as demonstrated by Hong

and co-workers [107], by immobilizing the multiple terminal groups to a surface. This yields

single core sites which are well separated from each other for subsequent immobilization of the

oligonucleotides [107]. Poly(amido amine) dendrimers were used but the multiple amido sites

and the branched character of the dendrimers led to very high levels of non-specific adsorption

of oligonucleotides. Another spacer molecule, a trialdehyde template [134], was synthesized to

serve as a temporary protecting group for three closely located amine sites by forming siloxy-

Schiff bases with commonly used aminosilanes [134]. Upon acid-catalyzed hydrolysis, the

central part of the molecule can be removed to yield three equally spaced amine sites on the

surface. This molecular template was abandoned when the “grafting from” technique which it

was intended to use with was also discarded; this technique will be discussed next.

The next approach was a “grafting to” strategy in which the entire amine-functionalized

oligomers (either synthesized or commercially purchased) such as poly(ethylene glycol) (PEG)

were mixed with oligonucleotides in solution and the mixture was introduced to 3-

glycidyloxypropyltrimethoxysilane (GOPS)-modified glass surfaces. It was determined that

there was a threshold concentration of amine-functionalized PEG (with similar or shorter lengths

as compared to oligonucleotides) relative to amine-functionalized oligonucleotides for the

competitive immobilization reaction. Once the loading concentration ratio between

oligonucleotides to PEG was beyond 1:5, the immobilization density of oligonucleotides

58

remained the same. This was probably because immobilization of entire polymers was not as

efficient as the immobilization of the oligonucleotides. Furthermore, the experimental

conditions may not have been optimal for the immobilization step. However, the hydrodynamic

volume that the polymers occupy may also impede subsequent immobilization reactions, as

explained by Jordan and co-workers [135]. Although the “grafting to” approach usually involves

one reaction step and that the polymers can be characterized by gel permeation chromatography,

nuclear magnetic resonance (NMR) and mass spectrometry, it is known that “grafting to”

compared to “grafting from” techniques do not yield highly dense polymer brushes on surfaces.

Thus, the “grafting to” strategy was also abandoned. A “grafting from” method involving the

surface-initiated polymerization of 4-styrenesulfonic acid was attempted, using 2,2,6,6-

tetramethyl-1-piperidinyloxy (TEMPO) via the nitroxide-mediated radical polymerization

(NMRP). However, the reaction needed to be done at a refluxing temperature of a mixture of

water and ethylene glycol, which might compromise the integrity of oligonucleotide-modified

surfaces. Furthermore, the reaction rate was too fast, was difficult to control, and yielded a very

thick film (> 200 nm).

In this thesis, a facile “grafting from” strategy for the development of a mixed film on

silica-based substrates composed of nucleic acid probes and non-nucleic acid oligomer is

proposed. The comparison of melt curves collected from pure oligonucleotide films and mixed

films was used to quantitatively determine improvements in selectivity of discrimination

between FC and SNPs. The immobilization of mixed films was carried out on an aminosilane-

activated layer on silica-based substrates by first immobilizing pre-synthesized amine-terminated

oligonucleotides through a heterobifunctional linker, followed by surface-initiated atom transfer

radical polymerization (ATRP) of 2-hydroxyethyl methacrylate, which is the non-nucleic acid

oligomer component. Surface-initiated living radical polymerization such as ATRP enables the

59

growth of polymers one monomer at a time from the surface, which can avoid any steric issues

associated with immobilizing whole polymer chains, allowing the growth of dense brushes [136].

The two separate reactions were achieved by protecting half of the amount of 3-

aminopropyltrimethoxysilane (APTMS) for the second step with benzaldehyde (BZ) [108].

The first chapter in the discussion section is about the characterization of various

compounds, such as the protected APTMS (BZ-APTMS), initiator for the ATRP reaction, and

PHEMA, by NMR. The efforts made in characterization ensured that the critical compounds

were pure, and were synthesized successfully. PHEMA was polymerized in solution and its

structure was confirmed by NMR, indicating that the experimental procedure for the reaction

worked and providing confidence that the procedure could be examined as a novel method for

the modification of surfaces.

The second chapter presents the strategy that was used to build mixed films on silica-

based substrates. Surface characterization techniques confirmed that surface-initiated ATRP

proceeded on silicon wafers. The signal-to-background ratios, immobilization densities and

hybridization efficiencies were determined and compared between surfaces with only

oligonucleotide probes and those with PHEMA brushes. The reusability of the mixed film

further demonstrated that the mixed film was functional when used for DNA hybridization and

denaturation. Finally, melt curve experiments clearly demonstrated that there was an increase in

selectivity for 3 bpm targets observed on the mixed film compared to the oligonucleotide film by

sharpening the melting transition and increasing the difference in melting temperatures of the

two targets. The increase in melting transition slope was attributed to a narrowing of distribution

of surface free energies due to the “matrix isolation” strategy.

60

In the third chapter, a comparison of surface characterization techniques led to the

conclusion that ToF-SIMS equipped with a bismuth cluster ion source offered a sensitive

approach for detection of a low density of oligonucleotides in the presence of excess PHEMA

brushes. The oligonucleotide density was determined by using fluorescence and a gold

nanoparticle-labeling strategy. The density confirmed the enhanced sensitivity of ToF-SIMS

combined with the use of a polyatomic cluster ion source. Thus, this section is significant for

evaluating which techniques can best characterize these mixed films.

In the fourth and final chapter, a contribution is made towards the detection of SNPs by

demonstrating some optimization of selectivity by mixed films compared to the oligonucleotide

films. Evidence about the homogeneity and structure of mixed films was obtained. The

fluorescence lifetimes of Cy3 labels attached to oligonucleotide probes on surfaces with PHEMA

were twice as long as those without PHEMA. The increase in lifetime values indicated an

increase in rigidity of the mixed film environment compared to the oligonucleotide film. Melt

curves were collected from various surfaces at three different ionic strengths. A combination of

increased differences in Tm values as well as increased steepness of melting transitions resulted

in enhanced SNP discrimination.

Cumulatively, these studies verified that the mixed aminosilane layer and the ATRP-

based growth of oligomer were successful in producing mixed films that integrated

oligonucleotides within polymer brushes. This mixed film strategy proved to be practical in

enhancing signal-to-background noise by reducing adsorption, and oriented oligonucleotide

probes to conformations favourable for hybridization. The presence of the oligomer portion

exhibited both stabilizing and destabilizing forces, depending on the degree of complementarity

of the target, to further increase the resolution between the two target populations by sharpening

61

the melting transition slope and increasing the melting temperature differences. The use of

polymers such as PHEMA is advantageous as it does not tend to degrade at high temperature and

is stable in extreme pH ranges without being hydrolyzed [137], and it is not sensitive to the

presence of salts as gold nanoparticles are unless a specific aging protocol is applied [138]. The

increased ability of the mixed film to discriminate SNP targets from FC targets makes it an

excellent construct suitable for numerous types of biosensors and also for microarray

applications.

62

Chapter 2

2 Experimental

2.1 Materials Redistilled benzaldehyde (BZ, >99.5%), 4-nitrobenzaldehyde (98%), (3-

aminopropyl)trimethoxysilane (APTMS, 97%), redistilled N,N′-diisopropylethylamine (99.5%),

2-bromoisobutyryl bromide (98%), N-hydroxysuccinimide (NHS), triethylamine (99%), 2-

hydroxyethyl methacrylate (HEMA, 97%), ethylene glycol methyl ether methacrylate (EGMEM,

99%), ethylene glycol methacrylate phosphate (EGMP), tris(2-carboxyethyl)phosphine

hydrochloride (TCEP), styrenesulfonic acid (SSA), Cu(I) chloride, Cu(II) bromide, 2,2′-

dipyridyl, sodium bicarbonate, sodium sulfate, sodium dodecyl sulfate, sodium carbonate,

sodium chloride, sodium orthophosphate were from Sigma–Aldrich (Oakville, ON). Tris

hydrochloride was from EM Science (Gibbstown, NJ, USA). (Sulfosuccinimidyl 4-[N-

maleimidomethyl]cyclohexane-1-carboxylate) (sulfo-SMCC) was from Fisher Canada (Nepean,

ON). Ammonium hydroxide (30%) and hydrogen peroxide (30%) were from EM Science

(Gibbstown, NJ, USA). An inhibitor present in the HEMA was monomethyl ether hydroquinone

(MEHQ). HEMA was purified by using MEHQ inhibitor remover (Sigma–Aldrich, Mississauga,

ON), which was packed in a column. All salts were dissolved in Millipore purified water (Milli-

Q water, 18 MΩ⋅cm). All solvents including methanol (MeOH), dichloromethane (DCM),

deuterated chloroform (CDCl3), diethyl ether, dimethylformamide (DMF), dioxane, toluene, and

silica gel for chromatography were from Sigma–Aldrich (Oakville, ON). Argon and nitrogen

gas were from BOC Canada Limited (Oakville, Canada).

63

Substrates included silicon wafers from International Wafer Service (Limerick, PA,

USA), and glass microscope slides (3 in. × 1 in. × 1 mm) from Fisher Scientific (Pittsburgh, PA,

USA).

Table 2.1 lists the oligonucleotide sequences from Integrated DNA Technologies

(Coralville, IA). A small aliquot of the disulfide form of each thiol-modified oligonucleotide

were first reduced by TCEP and purified through a Sephadex G-25 DNA grade column from GE

Healthcare (Baie d'Urfé, Québec, Canada).

2.2 Instrumentation 1H-NMR and 13C-NMR spectra were recorded on a Bruker Avance III 400 MHz (Bruker

BioSpin, Billerica, MA, USA) and a Varian Unity Inova 500 MHz NMR spectrometers (Varian,

Palo Alto, CA, USA). UV/Vis absorption spectra were measured using a Libra S22 spectrometer

(Bichrom Ltd., Cambridge, UK) and a HP 8452 A Diode-Array Spectrometer (Hewlett Packard

Corporation, Palo Alto, CA, USA). Solution phase fluorescence spectra were measured using a

QuantaMaster PTI spectrofluorimeter and Felix Software (Photon Technology International,

Lawrenceville, NJ, USA) equipped with a xenon lamp.

Time-of-flight secondary ion mass spectrometry (TOF-SIMS) experiments (Chapter 3.2)

were done at Surface Interface Ontario, University of Toronto. The instrument used was an

ION-TOF TOF-SIMS IV (Muenster, Germany), which was operated in static mode. The ion

source was a 69Ga-liquid metal ion gun (LMIG) source with a potential of 25 keV and a

maximum current of 2.5 pA. A spot area of 500 μm × 500 μm was sampled. Both negative and

positive ion modes were investigated and a mass range from 0–500 amu was scanned for the

64

SIMS spectra. TOF-SIMS mass spectra were obtained using the “Bunch mode” which provided

a high resolution determination of mass [139].

Other TOF-SIMS experiments (Chapter 3.3) used the same ToF-SIMS spectrometer but

used a Bin+ cluster ion primary source with a potential of 25 keV and a maximum current of 1

pA. A spot area known to contain oligonucleotides of 500 μm × 500 μm was sampled. Both

negative and positive ion modes were investigated and a mass range from 0 to 750 amu was

scanned for collection of SIMS spectra. The raw TOF-SIMS data were processed by IonSpec

Application software, version 4.5.0.0 by © ION-TOF GmbH (1996-2002).

The X-ray photoelectron spectroscopy (XPS) analysis (Chapter 3.2) was done using a

Leybold Max-200 spectrometer (Leybold-Haraeus, Cologne, FRG) at Surface Interface Ontario,

University of Toronto. The X-ray source was non-monochromatic Mg Kα (1253.6 eV), and

target samples were 1 cm × 1 cm in dimensions. The excitation voltage was 12.5 kV and the

emission current was 20 mA. A pass energy of 192 eV was used for low resolution survey and

elemental composition analysis, whereas a pass energy of 48 eV was used for high resolution

scans. Angularly dependent XPS spectra were recorded at take-off angles of 90°, 45°, 30°, and

20°. The software used to process the XPS data was SpecsLab v. 1.8.2 (© 1993–1998) Specs

GmbH.

Other XPS experiments (Chapters 3.2 and 3.3) were done using a Thermo Scientific

Theta Probe at Surface Interface Ontario, University of Toronto. The X-ray source was

monochromatic Al Kα (1486.68 eV) and was focused to a spot of 300 μm. The excitation

voltage was 12 kV and the emission current was 6 mA. A pass energy of 200 eV was used for

low resolution survey and elemental composition analysis, whereas a pass energy of 30 eV was

65

used for high resolution scans. XPS spectra were recorded at a take-off angle of 30°, 50°, 70°.

The low resolution spectra of C1s, N1s, and P2p were collected. For high resolution spectra, only

those of C 1s were collected. The XPS peak positions, curve fitting, and integrated intensities

were processed by the Avantage v.4.17 software by Thermo VG Scientific. All the spectra were

normalized against C 1s at 285.0 eV.

The atomic force microscope images of silicon wafer and glass surfaces were obtained

from a Digital Instruments Nanoscope Atomic Force Microscope (Santa Barbara, CA, USA),

courtesy of Dr. Shirley Tang from the University of Waterloo. The imaging was done in air

under tapping mode. The AFM tip was made of silicon nitride with a spring constant of 40 N m-1

and a nominal radius of less than 10 nm. The software used to process the images was

Nanoscope IIIa (Santa Barbara, CA, USA).

Scanning transmission electron microscope (STEM) imaging was done by a Hitachi HD-

2000 (Hitachi Instruments Inc., San Jose, California, USA) and SEM imaging was done by a

Hitachi S-5200 SEM (Hitachi Instruments Inc., San Jose, California, USA) at the Centre for

Nanostructure Imaging, University of Toronto. For STEM imaging of the Au nps-

oligonucleotide conjugates, a drop of the solution was placed on a TEM grid (carbon film on

copper grid) (Pelco International, Redding, CA, USA). The excess solution was dried with

Kimwipes at the edge of the grid and air-dried. The electron gun voltage was set at 200 kV and

the sample was magnified to 700,000× under (Z-contrast) ZC mode. For scanning electron

microscope (SEM) imaging of Au nps-oligonucleotide conjugates on fused silica slides, the

surfaces were cut to a size of 1.1 cm × 0.5 cm and they were adhered to SEM stubs by

conductive carbon paint (SPI Supplies, West Chester, PA, USA). They were also carbon coated.

The electron gun voltage was set at 5.0 kV and the sample was magnified to 250,000× under

66

secondary electron (SE) mode. The software used to view the STEM and SEM images was

Quartz PCI (Quartz Imaging Corporation, Vancouver, BC, Canada).

Thickness measurements were done using a Rudolph Research ellipsometer (Auto EL III

Rev 0.508) with a He–Ne laser emission at 632.8 nm which was incident at an angle of 70.00°.

The phase (Δ) and amplitude (Ψ) values that were measured were used to compute the thickness

and refractive index values by the software that was developed by McCracken [140]. The

refractive indices used for silane-coated and polymer-coated surfaces were 1.46 and 1.5,

respectively.

Wetting angle measurements were done using a monocular light microscope which had

been placed on its spine, with the sample between the objective and light source, clamped in an

external x-y-z adjustable stage.

Fluorescence intensities originating from glass slides were collected with a Versarray

Chipreader 5 µm System (Bio-Rad, Hercules, California, USA) equipped with 532 nm and 635

nm laser sources and two detection channels.

The lifetime of the Cy3 emission was measured from labeled oligonucleotides (Sequence

2, Table 2.1) which were co-immobilized with and without PHEMA onto microscope glass cover

slips (size 1 ½, Corning). The experiments were performed on a custom-built hyperspectral

confocal microscope described previously [141]. In brief, a femtosecond laser (Tsunami HP,

Spectra Physics, USA) was tuned to 960 nm (FWHM = 13 nm) and was frequency-doubled in a

nonlinear crystal to produce a narrow excitation spectrum centered at 480 nm. This linearly

polarized beam passed through a 1.4 NA/100x plan-apochromat objective (Carl Zeiss Canada)

and illuminated the sample at intensities in the range of 100 W/cm2. The emitted fluorescence

67

emission was collected through the same objective and was spatially and spectrally filtered using

a 50-μm pinhole and high-quality long-pass and band-pass filters (Semrock, Rochester, USA) to

remove out-of-focus fluorescence and the Rayleigh scattering from the surface. Further, the

fluorescence was divided into two components with polarization parallel and perpendicular to the

polarization of the excitation beam using a broadband polarizing cube beamsplitter (Newport,

Irvine, USA). Each beam was tightly focused onto an avalanche photodiode that featured a low

dark noise, high sensitivity and picosecond timing (PD5CTC, Optoelectronic Components,

Kirkland, Canada). Each time a photon was detected, the detector output an electric pulse that

was registered by a counting module (PicoHarp300, PicoQuant Gmbh, Germany). The photon-

by-photon data contained information about the emission rate, the excited-state lifetime, and

anisotropy.

2.3 Procedures

2.3.1 Preparation of Silicon, Fused Silica, and Glass Substrates

Substrates were cleaned by the RCA1 and RCA2 methods [142]. They were first

immersed in a 80 °C mixture of 1:1:5 of 30% NH4OH: 30% H2O2:water (v/v/v) (Milli-Q water

was used in all preparations and washings) for 5 minutes. They were then rinsed with water. In

the second step, they were immersed in a mixture of 1:1:5 of concentrated HCl: 30% H2O2:

water for another 5 minutes at 80°C. The substrates were rinsed with water and sonicated in

methanol for 15 min twice. They were finally rinsed in dichloromethane and ether and stored in

an oven at approximately 150 ºC.

68

2.3.2 Synthesis of Benzaldehyde (BZ)-Capped APTMS [Benzylidene-(3-trimethoxysilanyl-propyl)-amine]

The procedure as reported by Hicks [101, 143] was followed and Fig. 2.1 shows the

reaction. The reagents were added in a nitrogen-filled glovebox. Benzaldehyde was mixed with

APTMS at a 1:1 mole ratio and dissolved in toluene. Molecular sieves were added. The reaction

was stirred and refluxed under Ar for 24 h. The molecular sieves were taken out and the toluene

was removed by a rotary evaporator. The product was used without purification.

Characterization was done by 1H and 13C-NMR. NMR samples were dissolved in CDCl3 and

analyzed at 200 MHz. 1H data: δ 0.70, 1.80, 3.55, 7.33, 7.62, 8.21. 13C data: δ 6.79, 24.05,

50.29–50.80, 64.17, 128.04, 128.55, 130.50, 136.34, 161.11.

Fig. 2.1: Synthesis of BZ-APTMS.

2.3.3 Immobilization of APTMS or BZ-APTMS

In a nitrogen-filled glovebox, substrates were suspended in a mixture of

APTMS/toluene/N,N′-diisopropylethylamine at a 30:100:1 v/v/v ratio. For immobilizing both

BZ-APTMS and APTMS, the ratio was 15:15:100:1 v/v/v (BZ-APTMS/APTMS/toluene/N,N′-

69

diisopropylethylamine). The reaction was shaken and heated at 80-90 °C under argon for 24 h.

The samples were rinsed in toluene, 1:1 v/v toluene:methanol, and were then sonicated in

methanol for 15 min twice. They were further rinsed with dichloromethane and diethyl ether.

The samples were stored in a dessicator under vacuum until required. The immobilization of

mixed BZ-APTMS and free APTMS yielded a surface as depicted in Fig. 2.2.

Fig. 2.2: Immobilization of 1:1 ratio of APTMS and BZ-APTMS on silica-based substrates.

2.3.4 Immobilization of Heterobifunctional Linker, Sulfo-SMCC

Approximately 0.5–1 mg of fresh sulfo-SMCC linker was dissolved in 1.5 mL of 1×

phosphate-based saline (PBS, 1.0 M NaCl, 50 mM Na2HPO4, pH 7.0) buffer. About 10–20 μL

of this solution was deposited onto the substrates and was trapped with a glass cover slip for 30

minutes. The surfaces were rinsed with water and dried with a stream of nitrogen gas. This step

produced a surface as depicted in Fig. 2.3.

70

Fig. 2.3: Immobilization of sulfo-SMCC onto silica substrates. R = O-Si(OCH3)2-(CH2)3-

2.3.5 Immobilization of SH-SMN Probe

An aliquot of the stock oligonucleotide probe material (Table 2.1, Sequence 1) in its

disulfide form was cleaved to its free thiol form by using 2 μL of 20 mM TCEP and purified

using a Sephadex G-25 column with elution by water. The purified material was diluted in 1 ×

PBS to prepare a 1 µM solution and 2 µL of 20 mM (Chapter 3.2) or 300 µM TCEP (Chapter

3.4) was added prior to immobilization. The probe and a reference sequence (Table 2.1,

Sequence 2 or 3) were deposited on the surface as 2 µL spots overnight in dark, moist chamber.

Substrates were rinsed the next day with water and dried with a stream of nitrogen. This step

resulted in a surface depicted in Fig. 2.4 and 2.5.

71

Table 2.1: Oligonucleotide sequences used.

Sequence # Sequence Notes

1 5΄-SH-C6-ATTTTGTCTGAAACCCTGT-3΄ Thiol-modified SMN probe

2 5΄-SH-C6-ATTTTGTCTGAAACCCTGT-Cy3-3΄ Thiol-modified Cy3-labeled SMN

probe

3 5΄-SH-C6-ACAGGGTTTCAGACAAAAT-Cy3-3΄ Thiol-modified Cy3-labeled SMN

target

4 5΄- ATTTTGTCTGAAACCCTGT-Cy3-3΄ Cy3-labeled SMN probe

5 5΄-Cy3-ACAGGGTTTCAGACAAAAT-3΄ Fully complementary target

6 5΄-Cy3-ACAGGGTTTTAGACAAAAT-3΄ 1 Base-pair mismatch target

7 5΄-Cy3-ATAGGGTTTCGGACAAAGT-3΄ 3 Base-pair mismatch target

Underlined bases indicate mismatches.

Fig. 2.4: Confocal image (Cy3 channel) of a glass slide that was spotted with Sequence 1 and the reference spots

(green spots) used Sequence 2.

72

Fig. 2.5: Immobilization of oligonucleotides onto mixed APTMS and BZ-APTMS modified silica substrates. R =

O-Si(OCH3)2-(CH2)3-. R1 is sequence 1, 2, or 3 from Table 2.1 (SH-C6 portion is included in this figure).

2.3.6 Hydrolysis of Benzylimine

The substrates were immersed in a mixture of 1:1:1 H2O:MeOH:1 M HCl and shaken for

6 h. The substrates were then sonicated in water, and then in MeOH for 15 minutes each. The

benzaldehyde group was thus removed (Fig. 2.6).

73

Fig. 2.6: Hydrolysis of remaining BZ-APTMS sites. R = O-Si(OCH3)2-(CH2)3-. R1 is sequence 1, 2, or 3 from

Table 2.1 (SH-C6 portion is included in this figure).

2.3.7 Synthesis of Bromoisobutyryl NHS Ester Initiator

The procedure was adapted from Lou et al. [144] and the reaction is shown in Fig. 2.7.

2.7 mL of bromoisobutyryl bromide was added to 100 mL of diethyl ether that was already

chilled in an ice bath. In a separate round-bottom flask, 1 g of N-hydroxysuccinimide (NHS) and

1.8 mL of triethylamine were added to 20 mL of 1,4-dioxane. The NHS solution was added

dropwise to the bromoisobutyryl bromide solution over a 30-minute period and was then stirred

at room temperature for 1 h. The precipitate that formed was filtered using a Buchner funnel and

diethyl ether was used to rinse the precipitate. The filtrate was extracted 3 times with saturated

NaHCO3 (50 mL each time), and three times with water (50 mL each time). The organic layer

74

was dried with Na2SO4, which was then removed by filtration. The filtrate was concentrated in a

rotary evaporator. The crude product was separated by column chromatography using 100%

dichloromethane (silica gel, dichloromethane, Rf value = 0.2) and then 100% MeOH/DCM. 1H

NMR data: δ 2.06 (s, 6H), 2.86 (s, 4H). 13C NMR data: δ 25.646 (methylene), 30.737 (methyl),

50.875 (tertiary), 167.524 (NHS carbonyl), 168.566 (bromoisobutyryl carbonyl).

Fig. 2.7: Synthesis of the bromoisobutyryl NHS ester initiator.

2.3.8 Immobilization of Bromoisobutyryl NHS Ester onto APTMS-Modified Silicon Wafers

For every 10 mL of 1 M NaHCO3/Na2CO3 (pH 9.0) which was added to APTMS-

modified substrates, 1 mL of 20 mg mL-1 of the bromoisobutyryl NHS ester in DMF was also

added. The reaction was shaken for 30 min at room temperature. Substrates were sonicated in

the NaHCO3 buffer, water, and methanol for 15 minutes for each sonication step. The

immobilization of the initiator is shown in Fig. 2.8.

75

Fig. 2.8: Immobilization of initiator onto remaining APTMS sites. R = O-Si(OCH3)2-(CH2)3-. R1 is sequence 1, 2,

or 3 from Table 2.1 (SH-C6 portion is included in this figure).

2.3.9 ATRP of HEMA on Bromoisobutyryl-Immobilized Silicon Wafers

HEMA was first purified by eluting it dropwise through a column containing the MEHQ

inhibitor remover. In a 250 mL air-free flask, 20 mL of the purified HEMA and 20 mL of water

were degassed by purging the mixture with Ar and stirring for 20 min. CuCl, CuBr2, and 2,2′-

dipyridyl were mixed in the solid state at a molar ratio of 1:0.3:2.9 (CuCl is at 1 mmol) and was

added to the monomer mixture following the first purge cycle (see Fig. 2.9). This was further

purged for 30 min with Ar gas. The catalyst/monomer mixture was then transferred to an Ar-

purged flask containing the initiator-immobilized substrates. The reaction was shaken at room

76

temperature under Ar for 2–3 h. Figure 2.10 shows the surface-initiated structure of polymerized

HEMA (PHEMA).

Fig. 2.9: Purging the monomer/copper catalyst mixture with Ar gas. There was an Ar gas inlet which was inserted

through the septum to ensure the atmosphere inside the flask always contained Ar. A long stainless steel needle was

used to deliver Ar into the solution directly. A rubber septum was used to loosely plug the flask with the needle

being inserted outside of it. Excess gas could escape through the gap caused by the rubber septum and needle.

For kinetic studies of ATRP (Chapter 3.2), the initial time was set as the point when the

copper/monomer mixture was added. There was also a flask with Si wafers that did not contain

any initiator on the surface to serve as a control. Wafers were removed from the reaction flasks

at certain time intervals and the wafers were subsequently sonicated in MeOH, and then rinsed in

DCM and ether. The apparent thicknesses were then measured by ellipsometry.

77

Fig. 2.10: ATRP of HEMA on initiator-immobilized sites. R = O-Si(OCH3)2-(CH2)3-. R1 is sequence 1, 2, or 3

from Table 2.1 (SH-C6 portion is included in this figure).

2.3.10 Hybridization of Target Oligonucleotides with Probes on Aminosilane Surface or in Mixture with PHEMA on Glass Substrates

Various oligonucleotide targets (Sequences 5–7, Table 2.1) were introduced to glass

substrates modified with a pure oligonucleotide film (Fig. 2.5) or mixed film (Fig. 2.10). They

were heated in a block heater for 5 min at 90-95 °C so that the targets could hybridize with

surface-immobilized probe molecules from a temperature above the relevant Tm value. A 20 µL

aliquot of each hot target solution at 1 μM in 1 × PBS (Chapter 3.2) or 0.1× PBS (Chapter 3.4)

was immediately deposited onto an area of the glass slide which encompassed 8 oligonucleotide

probe spots (see Fig. 2.11). The other target was deposited onto another area of the same glass

78

slide with 8 oligonucleotide spots. Each target solution was separately trapped with a glass cover

slip and the slides were incubated in a moist chamber for 1 h at room temperature. The slides

were then gently washed with water and dried with a stream of nitrogen gas.

Fig. 2.11: Hybridization of various targets (Sequences 5–7) to immobilized probe spots.

2.3.11 Measurement of Immobilization Efficiency and Hybridization Yield (Chapter 3.2)

A set of various Cy3-labeled SMN probe (Sequence 2 in Table 2.1) concentrations (0.1,

0.5, 1.0, 5.0, 10.0, 20.0, 40.0, 60.0, 80.0, 100.0 nM) were prepared to construct a calibration

curve to determine the yield and efficiency. The DNA probe was dissolved in 1× PBS and 2 μL

of each concentration was spotted on a glass slide with the same chemistry as the surface in

question (oligonucleotide only or mixed film). The spots were air-dried. The integrated

fluorescence signals were averaged and were plotted against the theoretical number of probe

strands. A linear trend was used to fit a range of data points from the calibration slide that

included the fluorescence intensities in question. The number of immobilized probes and

hybridized targets were calculated from the calibration curve and Cy3 signals.

3 bpm

0 bpm

1 bpm

79

2.3.12 Acquisition of Melt Curves from Oligonucleotide Probe Films with and without PHEMA on Glass Surfaces

Glass substrates carrying hybridized DNA were immersed in various strengths of PBS

buffer (0.1×, 0.5×, and 1.0×). The temperature was increased at a rate of 0.4–0.6 °C min-1. At

each step of temperature increase, the glass slide was dried with a stream of nitrogen and was

scanned by the confocal Chipreader scanner at an excitation wavelength of 532 nm (see Fig.

2.12), after which it was returned to solution for further heating. For each scanned image, the

integrated intensity from any remaining Cy3-labeled targets at each spot on the glass surface was

measured using a LabVIEW program (National Instruments, Austin, TX, USA) written by W.

Russ Algar. Temperature correction for the decrease in fluorescence intensity at elevated

temperatures was applied as a monoexponential function. All fluorescence signals were

normalized by using the standard equation:

fssDNA = (IU-I)/(IU-IL) (2.1)

where I is the fluorescence intensity at each temperature, IU is the fluorescence intensity at the

upper baseline, IL is the fluorescence intensity at the lower baseline, and fssDNA is the fraction of

single-stranded target DNA that have denatured from the surface. The fssDNA was plotted with

respect to temperature. The melt curves were fitted with a sigmoidal function by using

OriginPro (version 7.5, OriginLab Corporation, Northampton, MA, USA).

80

Fig. 2.12: Confocal images (Cy3 channel) of hybridized targets (Sequences 5–7) on a mixed film glass surface and

their fluorescence intensities as temperature was increased. From left to right: 23°C, 49°C, 55°C, 63°C.

2.3.13 Conjugation of Thiolated-Oligonucleotides to Au Nanoparticles (Chapter 3.3)

133.6 µL of 688 nM of the Cy3-labeled thiolated target (Sequence 3, Table 2.1), which

was dissolved in water, was mixed with 450 µL of stock Au nanoparticles (68.1 nM, 5 nm in

diameter) in a microcentrifuge tube. The solution ratio of DNA to Au nps is 3:1. 10 µL of 20

mM of TCEP was added. The mixture was incubated overnight at room temperature. The next

day, 500 µL of 0.1× PBS was added to the mixture and the sample was vortexed briefly. The

mixture was incubated at room temperature for two more days.

The Au-DNA conjugates were centrifuged at 13,000 rpm for 25 min per cycle. A 200-µL

aliquot of the Au-DNA conjugate was centrifuged with 600 µL of 95% ethanol for each cycle. A

faint red oily film was deposited on the side of the microcentrifuge tube and the supernatant was

removed with a pipette. The subsequent aliquots of the sample and ethanol were added to the

same microcentrifuge tube containing the red oily film and the centrifuge process was repeated.

81

The entire purification procedure was repeated about four times in total. The deeper red oily

film from the final centrifugation step was dispersed in 0.3× PBS.

2.3.14 Dissolution of Gold Nanoparticles by KCN and Construction of a Calibration Curve for Calculation of Surface Density of Oligonucleotides on Gold Nanoparticles

A typical etching experiment involved dilution of 20 μL of the stock Au nps-DNA

conjugate solution in 60 μL of 0.1× PBS. The fluorescence spectrum of each diluted conjugate

sample (30 μL aliquot) was measured at an excitation wavelength of 520 nm and the emission

was collected from 540–650 nm. 5.00 μL of 40 mM KCN was then added and the fluorescence

spectrum was immediately scanned. The peak fluorescence intensity at 562 nm was used to

calculate the number of immobilized oligonucleotides.

To construct the calibration curve for the determination of oligonucleotides adsorbed

onto Au nps, thiolated oligonucleotides of various concentrations (62.5 nM, 125.0 nM, 250.0

nM, and 500.0 nM) of the SMN sequence labelled with Cy3 were mixed with 30.00 μL of stock

Au nps (68.1 nM), 5.00 μL of 40 mM KCN, and 0.1× PBS to make up a total volume of 105 μL.

The blank solution contained all reagents except for the oligonucleotides. The fluorescence

intensity of each DNA concentration was measured at an excitation wavelength of 520 nm and

the emission was collected from 540–650 nm. The peak fluorescence intensity at 562 nm was

plotted against the oligonucleotide concentration.

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2.3.15 Hybridization or Adsorption of the Au nps-DNA Conjugates to Fused Silica Surfaces Modified with Oligonucleotide Films or Mixed Films for SEM Analysis (Chapter 3.3)

An aliquot of 20 μL of the Au nps-DNA conjugate solution was diluted in 60 μL of 0.1×

PBS. The diluted solution was first heated at 60–70°C for 5 min and was smeared across

surfaces with a cover slip on PHEMA-covered surfaces with and without oligonucleotide probes.

The surfaces were then rinsed with water and dried with a stream of nitrogen.

83

Chapter 3

3 Results and Discussion

3.1 Syntheses of Benzaldehyde-Protected Aminosilane, Initiator, and PHEMA for Assembly of Mixed Film

3.1.1 Introduction

Covalent linkages were chosen for immobilization of the mixed film. The silane

coupling agent APTMS was first immobilized onto silica-based substrates by way of

alkoxysilane condensation [89, 90]. The free amine can then be used for immobilization of

thiolated oligonucleotides onto silica-based substrates via the heterobifunctional linker, sulfo-

SMCC. However, instead of using APTMS alone, half of the volume of APTMS used for the

silane immobilization reaction was first reacted with benzaldehyde (BZ) to protect the primary

amine group. This protection step reduces the number of crosslinkages between neighbouring

APTMS molecules [99], allows spacing and control of orientation of APTMS with respect to the

surface [101, 108], and temporarily saves amine sites for immobilizing the second component,

i.e. the PHEMA oligomer.

To grow PHEMA, surface-initiated ATRP was used, which required the immobilization

of an initiator. The chosen initiator compound is bromoisobutyryl bromide. This compound and

its derivatives are in common use for initiation of ATRP [123, 145, 146]. To immobilize the

initiator, it was first activated by coupling it to N-hydroxysuccinimide (NHS). Activation with

NHS enhances the rate at which the primary amine can react with the carbonyl group in

bromoisobutyryl because NHS is a good leaving group [147], making bromoisobutyrate a more

active electrophile [123]. Once the remaining amine sites were deprotected by hydrolysis, they

were free to react with the activated bromoisobutyrate initiator. Hence, immobilization of the

84

initiator to aminosilanized surfaces can be achieved. BZ-APTMS and the bromoisobutyryl NHS

ester were characterized by 1H and 13C NMR.

3.1.2 Synthesis of Benzaldehyde-Protected APTMS (N-[(1Z)-phenylmethylene]-3-(trimethoxysilyl)propan-1-amine)

Fig. 3.1 shows the structure of BZ-APTMS and its 1H NMR spectrum. The peak at ~ 0.7

ppm is consistent with the methylene protons adjacent to silicon (Protons a). The peak at 1.8

ppm was that of the methylene protons that are one carbon away from silicon (Protons b). At 3.5

ppm, protons from the methoxy groups (Protons c) were in a matrix that was highly crosslinked,

hence a broader singlet was observed. Protons d were not observed and they may be buried in

the broad methoxy proton peak since they were reported to be located at 3.6 ppm [101]. From 7–

8 ppm, the two peaks represented the two different groups of benzylic protons (Protons e and f).

The allylic proton (Proton g) at the carbon adjacent to the nitrogen was at 8.2 ppm, which again

agreed with Hicks and Jones [101]. The peak at 2.4 ppm was due to toluene, which was used as

the solvent in the reaction [148]. All the peaks found in the appropriate regions of the spectrum

agreed with the chemical shifts listed by Hicks and Jones [101], and the peak integration

provided results that were consistent with the structure of BZ-APTMS.

85

δ, ppm

Fig. 3.1: 1H NMR spectrum of BZ-APTMS in CDCl3. The inset contains the structure of BZ-APTMS. Different

types of protons are labeled.

86

3.1.3 Synthesis of the Bromoisobutyryl NHS Ester Initiator (1-[(2-bromo-2-methylpropanoyl)oxy]pyrrolidine-2,5-dione)

Fig. 3.2 shows the structure of the activated bromoisobutyryl NHS ester and its 1H NMR

spectrum. The spectrum contained two singlets, one for the methyl groups in the

bromoisobutyryl portion and one for the methylene groups in the NHS portion. The singlet

representing the methyl groups was at 2.1 ppm. Since the methyl groups were on the same

carbon and experienced a similar environment, they only appeared as a single peak. The

methylene protons should be more deshielded than the methyl protons due to the neighbouring

electron withdrawing carbonyl groups; hence the peak was found at a slightly more downfield

chemical shift, which was at 2.9 ppm. These NMR data agreed with those presented by Lou and

He [123].

The integration ratio between the singlet at 2.1 ppm and the one at 2.9 ppm accounted for

the number of protons in each group. The ratio was 1.7:1.0 (peak at 2.1 ppm: peak at 2.8 ppm

[123], which scaled to a ratio of 6:3.5. This was consistent with expectations because the peak at

2.1 ppm should have 6 protons (from two methyl groups) and the singlet at 2.9 ppm should have

4 protons (two methylene groups). The peak at 7.3 was the residual solvent peak from

chloroform [148]. Thus, the 1H NMR spectrum indicated that the bromoisobutyryl bromide was

successfully activated into its NHS form.

The 13C NMR spectrum of the initiator was also obtained and it matched well with the

results presented by Lou and He (Fig. 3.3) [123]. The assignment of the different carbons is also

shown in the 13C NMR spectrum. The corresponding chemical shifts are listed in Table 3.1. The

expected peaks should be located at 25.9, 30.9, 51.4, 166.2, and 168.9 ppm [123], which were in

agreement with the data in the table. The peak at 77 ppm is that of DMSO [148]. The NMR

data indicate that a pure form of the activated initiator was synthesized.

87

δ (ppm)

Fig. 3.2: 1H NMR spectrum of the activated bromoisobutyryl NHS ester initiator in CDCl3. The inset shows the

structure of the bromoisobutyryl NHS ester. The spectroscopically different protons are labeled.

88

δ, ppm

(a)

δ, ppm

(b)

Fig. 3.3: (a) 13C NMR (500 MHz) spectrum of the activated bromoisobutyryl NHS ester initiator in CDCl3. The

inset shows the structure of the bromoisobutyryl NHS ester. The spectroscopically different carbon atoms are

labeled; (b) high resolution data showing detail of the downfield region.

89

Table 3.1: Carbon peak positions of the bromoisobutyryl NHS ester.

Carbons δ (ppm)

a 25.5

b 30.4

c 50.8

d 167.4

e 168.5

3.1.4 NMR Characterization of PHEMA

To demonstrate that the polymerization procedure was functional, a solution-based ATRP

reaction was done. Figure 3.4 shows the 1H NMR spectrum of the resulting product in DMSO-d6

and the anticipated structure of the product. Table 3.2 lists each group of protons and their

chemical shifts. The chemical shifts and integrations were in accord with published data [149-

151]. The methyl peak (a) contained three protons and the hydroxyl (f) contained one. The

remainder of the peaks represented methylene groups and all were integrated to two protons.

The methyl protons (a) gave a doublet due to the tacticity of the polymer, as was also observed

for the methyl group in the carbon NMR spectra reported by Roman et al. and Verhoeven et al.

[152, 153]. The methylene protons (b) were located on the backbone of a polymer and were

expected to give a broad peak at 1.8 ppm. These NMR data showed that the ATRP reaction was

effective in solution, and provided the confidence that the procedure could be applied to surface-

initiated polymerization.

90

δ, ppm

Fig. 3.4: 1H NMR of PHEMA in DMSO-d6. Structure of PHEMA. The inset shows different types of protons,

which are labeled.

91

Table 3.2: Proton peak positions of PHEMA.

Protons δ (ppm)

a 0.8, 1.0

b 1.8

c 3.6

d 3.9

e 4.8

Acetone/Acetonitrile 2.1[148]

DMSO 2.5[148]

Water 3.4[148]

3.1.5 Conclusions

1H and 13C NMR demonstrated that BZ-APTMS, the bromoisobutyryl NHS ester

initiator, and PHEMA were successfully synthesized. BZ-APTMS would be mixed with

APTMS and used to immobilize the oligomer portion of the mixed film. The bromoisobutyryl

NHS ester would be immobilized to the remaining primary amine sites and used to initiate the

ATRP reaction. The successful synthetic scheme to produce PHEMA in solution was then

adapted for application for deposition onto surfaces.

92

3.2 Surfaces for Tuning of Oligonucleotide Biosensing Selectivity Based on Surface-Initiated Atom Transfer Radical Polymerization on Glass and Silicon Substrates

April K. Y. Wong and Ulrich J. Krull

Chemical Sensors Group, University of Toronto Mississauga, 3359 Mississauga Rd. N.,

Mississauga, ON, Canada L5L 1C6

Contributions: April Wong performed all measurements (except for XPS, ToF-SIMS, and AFM)

and data analysis and interpretation under the guidance of Ulrich Krull.

Previously published in Analytica Chimica Acta, 639 (2009) 1-12. Supplementary melt curve

data were added in this chapter for completeness.

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3.2.1 Abstract

Covalently immobilized mixed films of oligonucleotide and oligomer components on

glass and silicon surfaces are reported. This work has investigated how such films can improve

selectivity for the detection of multiple base-pair mismatches. The intention was to introduce a

“matrix isolation” effect on oligonucleotide probe molecules by surrounding the probes with

oligomers, thereby reducing oligonucleotide-to-oligonucleotide and/or oligonucleotide-to-surface

interactions. Thiol-functionalized oligonucleotide was coupled onto (3-

aminopropyl)trimethoxysilane (APTMS) via a heterobifunctional linker, sulfosuccinimidyl 4-

[N-maleimidomethyl]cyclohexane-1-carboxylate (sulfo-SMCC). Using a variety of monomers

such as 2-hydroxyethyl methacrylate (HEMA), oligomers were grown by surface-initiated atom

transfer radical polymerization (ATRP) from a bromoisobutyryl NHS ester initiator which was

immobilized onto APTMS sites that coated glass and oxidized silicon substrates.

Various surface modification steps on silicon substrates were characterized by

ellipsometry, wettability, atomic force microscopy, X-ray photoelectron spectroscopy, and time-

of-flight secondary ion mass spectrometry. Polymerized HEMA (PHEMA) in mixture with

oligonucleotide probes was evaluated for fluorescence transduction of hybridization. The

presence of PHEMA was found to provide a sharper melt curve for hybrids containing both fully

complementary and three base-pair mismatched (3 bpm) targets, and this surface derivatization

also minimized non-selective adsorption. The maximum increase in slope was improvement by

a factor of 2- to 3-fold. An increase of almost 2-fold in difference of melting temperatures

between fully complementary and 3 base-pair mismatched targets was achieved using PHEMA.

The results suggest that the presence of oligomers dispersed among DNA hybrids can improve

94

selectivity through what is believed to be a reduction of dispersity of interactions of probes with

targets, and probes within their local environment at a surface.

3.2.2 Introduction

The screening of nucleic acids remains an area of great interest as the medical and

environmental communities demand diagnostic technologies that are sensitive, selective,

reusable, and rapid to determine the presence of targets such as disease-causing genes, or the

presence of pathogenic bacteria in food/soil samples [154-156]. As a biological recognition

agent, oligonucleotide probes offer chemical stability, high affinity, versatility in terms of

sequence choice and length, opportunity for chemical modification, reusability, and facile

synthesis [154]. Such selective chemistry can be coupled to an electrochemical, piezoelectric, or

optical transduction system to achieve biosensing capability [154, 155]. The interest herein is the

further development of surfaces for immobilized oligonucleotide probes which can enhance

selectivity for the detection of multiple nucleotide polymorphisms.

The thermal melting temperature (Tm), the temperature at which half of any available

DNA duplexes have denatured, is characteristic of the stability of hybridization [157]. It is

known that for oligonucleotides in solution, every extra A-T base-pair provides about 2 °C

increase in Tm while every G-C base-pair contributes to a 4 °C increase [157]. Therefore, a

window of 4–8 °C typically exists for determination of 2–3 mismatches when operating in bulk

solution, and even less when considering single base polymorphisms (SNPs). Examples of SNP

studies have been reported by groups led by, for example, Mirkin and Corn [125, 158]. A

nanoparticle-enhanced SPR imaging-based detection strategy which was reported by Corn’s

95

group had a sensitivity of 1 pM [158]. An increase in SNP selectivity was demonstrated by

Mirkin’s group by using Au nanoparticle-based reporter probes [125]. The ability to

discriminate between closely related oligonucleotide sequences is also influenced by the

steepness of the melt curve.

Hybridization that occurs on surfaces rather than in solution tends to significantly

broaden melt curves as there is a wide diversity of physical conformations that can exist at an

interface, meaning that there is a distribution of hybridization energetics at a surface [159]. The

local environment of the probe oligonucleotides is a key factor that influences selectivity and

efficiency [160, 161]. In fact, the very process of hybridization on a surface when it occurs to a

substantial extent can dynamically change the surface charge density and the corresponding melt

curve [160, 161]. Orientation, steric conformation, and mobility of immobilized oligonucleotide

probes can be affected by the surface charge density due to probe-to-probe and probe-to-surface

interactions.

The present work explores a covalently immobilized mixed oligomer and oligonucleotide

film in order to ameliorate nearest-neighbour interactions of probes and to block interactions

with the substrate. The concept is one of “matrix isolation”, such that oligomers, immobilized

through one end, on average surround oligonucleotide probes so that probes are isolated from

one another and from the surface.

An earlier attempt by our team to develop a mixed oligonucleotide and oligomer film was

based on a photochemical approach, where partial activation of a linker which had been

protected by a photolabile group was done using ultraviolet (UV) irradiation [159]. Ethylene

glycol phosphate was immobilized on hexaethylene glycol linkers, and then the remaining

photolabile group was deprotected for immobilization of oligonucleotide probes. Melt curves

96

from the resulting mixed films tentatively suggested that the presence of an oligomer did exert a

useful effect on the Tm of the DNA hybrids, while still maintaining expected trends for

decreasing ionic strength and Tm differences between fully complementary hybrids and those

containing a single base-pair mismatch. However, the synthetic approach was not very practical

during sensor fabrication as the photolabile protecting group had to be synthesized and purified

in a darkroom to prevent photodegradation. Sensitivity to UV light leads to the potential for

photodegradation [162].

In this new study, we are exploring a different synthetic approach to achieve a matrix

isolation effect. In general, end-functionalized polymers can be directly grafted to a surface via a

specific chemical linkage. However, this “grafting to” method cannot yield a high surface

density of polymers [112], which is undesirable if one wants to build a highly controlled

environment for oligonucleotide probes. A more effective way to graft polymer brushes of high

density onto surfaces is called a “grafting from” method, in which immobilized initiators are

used to grow the polymer chains from the substrate one monomer at a time. One such “grafting

from” method can be achieved using atom transfer radical polymerization (ATRP). This is a

form of controlled/“living” polymerization that is capable of polymerizing various acrylate- and

methacrylate-based monomers to achieve a narrow polydispersity index (Mw/Mn <<1.5) with

controlled molecular weight [117]. Controlled/ “living” polymerization is achieved by having a

fast dynamic equilibrium between a higher concentration of dormant species (Pn–X) than of

active species (Pn·) usually at ppm concentrations [163], and is described by the following

reaction:

Pn-X + Cu(I)/2L Pn + Cu(II)X/2L• (1.21)

97

Pn• + monomer Pn+m• or Pn• (1.22)

where Cu(I) abstracts the halogen atom (X) from the initiator or dormant polymer chain (Pn–X)

and causes a homolytic cleavage of the halide-carbon bond, leaving a radical at the terminal end

of the polymer chain. Cu(I) becomes oxidized to Cu(II) upon complexing with the halide and

ligands (L), the latter being used to adjust the strength of the catalyst [115]. Wherever a radical

alkyl or polymer chain is available, an acrylate or methacrylate-based monomer can be added at

the end of this chain, thus propagating the radical to the end of this new monomer. The increase

in Cu(II) will deactivate the polymerization process and revert the active species (Pn•) back to its

dormant state (Pn–X) [163]. This reversible reaction is key to the controlled polymerization in

ATRP. Surface-initiated ATRP has been used in numerous applications such as the modification

of quantum dots, nanoparticles and microbeads, biosensing, protein adsorption studies, and

improvement of biocompatibility by means of surface coatings [121].

Here we report a strategy for creating mixed films using surface-initiated ATRP of 2-

hydroxyethyl methacrylate (HEMA) and other monomers on 3-aminopropyltrimethoxysilane

(APTMS)-modified glass and oxidized silicon substrates. Surface characterization methods were

used to determine the success of each reaction step. The resulting films provided an

advantageous biosensing surface for detection of hybridization of oligonucleotides by

fluorescence. The melt curves collected from mixed films showed sharper transitions than those

observed for immobilized films of oligonucleotides alone. Greater melting temperature

differences for 3 bpm mismatches were noted, and the mixed films ameliorated the problem of

non-selective adsorption.

98

3.2.3 Surface Characterization of Each Immobilization Step

3.2.3.1 Evidence of APTMS and BZ-APTMS on Si Wafers

The first step in the preparation of the immobilized oligomer film was to create a

template on the substrates using organosilanes such as (3-aminopropyl)trimethoxysilane, or

APTMS. It was selected because the primary amine group is compatible with forming imine

linkages (with benzaldehyde (BZ) for temporary protection of the primary amine) and amide

linkages (with sulfo-SMCC) for the subsequent immobilization of thiolated oligonucleotides via

the maleimide end of the sulfo-SMCC linker. Smith and Chen have recently pointed out the

vulnerability of aminosilane films to hydrolysis when exposed to water [164]. Although they

have tested a similar aminosilane on the same substrate, we have found no indication that

significant hydrolysis has occurred during our experiments. We have carefully prepared the

silane films in solution phase by using dry solvents and glassware. Our reaction conditions were

different in terms of reaction temperature, amount of silanes with respect to solvent, presence of

a base catalyst, rinsing and drying procedures. Our aminosilane films appear to be robust, and

treatment with an acidic solution for the hydrolysis of benzylimine step still yielded an intact

film as confirmed by ellipsometry.

The successful immobilization of free and benzaldehyde-protected APTMS was

determined by wettability, ellipsometry, and AFM. We have previously characterized 3-

glycidoxypropyltrimethoxysilane films on silicon substrates and we have used the same

experimental procedure to immobilize APTMS [92]. However, the APTMS immobilization

reaction time was reduced to less than 8 hours from 24 h, and the temperature was lowered to

about 80–100°C due to the lower boiling point of APTMS and its tendency to polymerize at the

glass joints.

99

The film thickness as determined by ellipsometry after the immobilization of APTMS

increased from an initial thickness of 18 ± 1 to 33 ± 3 Å. This corresponds to 15 ± 3 Å or 2

monolayers of APTMS. Kurth and Bein also reported a thickness of 11 ± 2 Å for vapor deposited

APTMS [165, 166]. Similar ellipsometry thicknesses were also obtained for the immobilization

of a mixture of free APTMS and BZ-APTMS (data not shown). The use of anhydrous reagents,

dried glassware and atmospheric conditions in this work minimized the formation of undesired

multilayered silanes.

The cleaned silicon substrates had an initial wetting angle of 17 ± 4° and this increased to

an angle of 61 ± 5° following the immobilization of BZ-APTMS. The increase in

hydrophobicity was due to a change from a hydrophilic silanol surface to one which was covered

with hydrophobic aromatic groups. The immobilization of APTMS alone also increased the

wetting angle to 50 ± 4°, which was consistent with data suggested by Kurth and Bein [165]. A

mixture of APTMS and BZ-APTMS provided a wetting angle of 57 ± 7°, which was found to be

between the wetting angles of BZ-APTMS and APTMS separately.

X-ray photoelectron spectroscopy (XPS) can determine the presence of specific bonds on

surfaces. The high resolution spectra of N 1s such as Fig. 3.5 suggest the presence of BZ-

APTMS and APTMS on Si wafers. The elemental identifications and quantifications are listed

in Table 3.3. According to Kristensen, the free amine peak in APTMS occurs at about 399.7 eV

[99]. This peak is not present in Fig. 3.5a, which was a cleaned Si wafer with no APTMS added.

The signal was present in Fig. 3.5b for samples that were treated with BZ-APTMS. However,

this peak at 399.87 eV could also be the C=N imine signal since it is known to be located from

398.7 to 399.7 eV [167], and at 398.9 eV [168], and 400 eV [169]. This may be the peak which

arose due to the formation of the imine bond between APTMS and benzaldehyde. Peaks beyond

100

400 eV may be due to hydrogen-bonded or protonated nitrogen and/or primary amine species

[99, 168, 169]. However, a portion of the counts contributing to the peak at 399.87 may also be

due to unreacted primary amines (free or hydrogen-bonded) [99, 169]. The origin of the peak at

398 eV is not clear for positive assignment. Figure 3.5c, which represents data from the

hydrolyzed APTMS sample, shows that the peak centered at 400 eV is still present. This

suggests that the free amines may be intact after the hydrolysis of the benzylimine bond. Further

confirmation of immobilized BZ-APTMS was found in the UV spectra of 4-nitrobenzaldehyde-

APTMS immobilized onto fused silica. The spectra revealed an absorption peak at ~282–288

nm, which represents the absorption by the imine bond, and this peak disappeared following

hydrolysis (data not shown).

1300

1400

1500

1600

1700

392393394395396397398399400401402403404405406407408409

Cou

nts

/ s

Binding Energy (eV)

N1s Scan #250 Scans, 7 m 32.5 s, 400µm, CAE 50.0, 0.10 eV

N1s

N1s Scan #2 A

N1s Scan #2 B

(a)

101

1300

1400

1500

1600

1700

1800

1900

392393394395396397398399400401402403404405406407408409

Cou

nts

/ s

Binding Energy (eV)


N1s

N1s Scan #2 A

N1s Scan #2 B

(b)

1200

1300

1400

1500

1600

1700

392393394395396397398399400401402403404405406407408409

Cou

nts

/ s

Binding Energy (eV)


N1s

N1s Scan #2 A

N1s Scan #2 B

(c)

Fig. 3.5: High resolution XPS spectra of N 1s and the peak locations of the various nitrogen-containing species with

their atomic percentages; (a) Cleaned Si wafer; (b) BZ-APTMS modified Si wafer; (c) hydrolyzed APTMS Si wafer.

102

Table 3.3: Elemental identifications and quantifications of the N 1s spectra.

(a) Peak BE At. %

N1s 398.12 29.27

N1s Scan #2 A 402.22 55.40

N1s Scan #2 B 400.65 15.33

(b) Peak BE At. %

N1s 399.87 56.16

N1s Scan #2 A 401.51 30.23

N1s Scan #2 B 398.05 13.61

(c) Peak BE At. %

N1s 400.02 49.49

N1s Scan #2 A 401.67 28.39

N1s Scan #2 B 397.97 22.12

The surface topology of the cleaned and chemically modified silicon wafers was

significantly different as determined by AFM (Fig. 3.6). Si nodules are evident on the cleaned Si

wafer [170], but were no longer present in the image of the APTMS-covered Si wafer (Fig.

3.6b). They were instead covered by an undulating film of a higher surface roughness. This

agreed with our previous observation of surfaces that were modified with 3-

glycidyloxypropyltrimethoxysilane [171]. Some heterogeneity in surface topology was expected

for the APTMS-modified surface. Kristensen et al. have summarized the various ways that

APTMS can adsorb onto silanol surfaces [99]. It was suggested that APTMS can react with the

surface via siloxane linkages, and is terminated either with a free amine or that the primary

amine can be located at the surface due to protonation either at a silanol site on the surface or by

103

other free silanols [99]. Further possibilities include a formation of siloxane bonds, as well as

hydrogen bonding between the primary amine and silanol on a surface. However, when BZ-

APTMS was immobilized, the surface roughness appeared to have reduced (Fig. 3.6c). This is

consistent with a reduction in intermolecular interactions between the free aminosilanes and the

surface, as explained by Kowalczyk, Horr and Chiang [96-98], leading to a more ordered film.

Hicks and Jones have previously shown that the protection of aminosilanes with benzaldehyde

causes a change in the distance between neighbouring silanes (using pyrene monomer and

excimer as markers in fluorescence spectroscopy) [143]. The protection of APTMS with

benzaldehyde can reduce interactions between APTMS molecules and the surface, as well as

provide a method to spatially immobilize APTMS to yield free primary amines away from the

surface. These characteristics would provide a strong foundation for building a reproducible

mixed oligonucleotide and oligomer film.

104

Fig. 3.6: AFM images of (a) cleaned bare Si wafer; (b) APTMS-modified Si wafer; (c) BZ-APTMS modified Si

wafer; (d) PHEMA grown on BZ-APTMS-coated Si wafer (~10 nm thick).

3.2.3.2 Evidence of Surface-Initiated ATRP on Si Wafers

Prior to polymerization, the initiator must be immobilized to the surface. The

bromoisobutyryl initiator was immobilized via the nucleophilic attack of the carbonyl group of

the butyryl portion of the molecule by surface amine groups. The succinimdyl group leaves

readily. The success of the immobilization of the initiator was marked by an increase in wetting

angle. Typically, the wetting angle was observed to be from 55 to 65° due to the presence of a

more hydrophobic hydrocarbon layer. There was not an appreciable increase in thickness seen in

the ellipsometry data due to the small size of the initiator. A study of thickness changes with

105

time as determined by ellipsometry (A.1) demonstrated that surface-initiated polymerization only

occurred on surfaces with the initiators present, with a rate described by the following equation:

)9697.0(4029.1198.20 2 =−= Rxy (3.1)

where y is the thickness in nanometers and x is the time in hours. The surfaces without the

initiators had the following relationship, without a discernable linear fit:

)0581.0(4993.20602.0 2 =+= Rxy (3.2)

The resultant brush-like topology of the polymerized surface can be seen in Figure 3.6d, and it is

evident that the surface morphology changed significantly from Fig. 3.6c. Further evidence of

the success of ATRP is seen in ToF-SIMS ion images in Fig. 3.7, in which samples with and

without initiator were immersed in an ATRP reaction mixture containing ethylene glycol methyl

ether methacrylate (EGMEM). The intensity for fragments of the monomer, polymer end group

(bromine) and APTMS are clearly more evident in samples immobilized with the initiator.

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Fig. 3.7: ToF-SIMS ion images of EGMEM on Si wafers grown via ATRP; (a) cleaned Si wafer; (b) initiator-

immobilized Si wafer.

Relative elemental compositions based on C 1s and Si 2p derived from low resolution

XPS spectra are shown in Fig. 3.8. The carbon signal decreased for both control (no initiator)

and derivatized surfaces as the angle increased, but the initiator-immobilized surface had a more

intense carbon signal and it did not decrease as much as the control sample over the angle scan

range. The carbon signal magnitude suggested much more organic overcoating for the initiator-

immobilized surface. The reduction of signal for the modified surface with decreasing angle was

only about 10% whereas the control sample decreased by 50%, indicating that the organic film

on the modified surface was much thicker than that of the control. A similar trend was observed

(a) Control wafer (b) Initiator-immobilized Si wafer

107

for the silicon signals. The control sample had a large increase in silicon signal (17.3%–42.3%)

starting from 20 to 90° take-off angle due to the presence of a very thin contaminant layer. The

initiator-immobilized surface had a much lower silicon signal at 90°, and only decreased

marginally at reduced angles, consistent with the surface being buried in a polymer layer.

0

10

20

30

40

50

60

70

80

90

10 20 30 40 50 60 70 80 90 100

take-off angle (degree)

Elem

enta

l com

posi

tion

%

July 11, 2006

C 1s

Si 2p

C 1s

Si 2p

Fig. 3.8: Change in elemental composition in an angularly-resolved XPS experiment of C 1s and Si 2p of Si wafers

that were and were not immobilized with the ATRP initiator using EGMEM as monomer. (•) no initiator, Si 2p;

(♦) initiator, C 1s; ( ) no initiator, C 1s; (▲) initiator, Si 2p.

Further XPS evidence is shown in Fig. 3.9 in which ethylene glycol methacrylate

phosphate (EGMP) was polymerized on Si wafers. The C 1s signatures for polar carbons are

evident in the XPS spectrum (Fig. 3.9c) after only 1 h, while Si 2p signals at 99.0 and 102.5 eV

were reduced in intensity (Fig. 3.9d). The accumulated evidence indicates that ATRP was

successful and that the increase in film thickness was not due to non-specific adsorption of

monomers.

108

Fig. 3.9: Low resolution XPS spectra of (a) C 1s at 0 h of ATRP; (b) Si 2p at 0 h of ATRP; (c) C 1s at 1 h of

ATRP; (d) Si 2p at 1 h of ATRP. EGMP was used as monomer.

3.2.4 Choice of Monomer

The surface characterization data were collected from surfaces that were polymerized

with HEMA, EGMP, and EGMEM. This set of reagents explored structures that were neutral

and hydrophilic, or charged, and the resulting polymers were able to swell in an aqueous

109

environment. Furthermore, these monomers contain ethylene glycol units or hydroxyethyl

groups, which may alleviate non-selective adsorption similarly to polyethylene glycol [172].

Rates of polymerization were compared using the mole ratio of 1:0.3:2.9 for CuCl:CuBr2:2,2΄-

dipyridyl and the same volume ratio of monomer to water (1:1, 20 mL each). This set of

conditions was chosen because Lou and He [173] found that these conditions provided the

optimum ATRP rate for PHEMA. Data from numerous trials indicated that HEMA was the best

candidate as it provided the most reproducible thickness control without significant adsorption.

HEMA polymerized controllably and in a “living” fashion, as evidenced by its linear growth, in

a reasonable length of time. The ATRP growth rate of PHEMA allowed production of a film of

thickness of tens of nanometers in an hour. EGMEM and EGMP, on the other hand, typically

varied in reaction rates and did not show a trend consistent with “living” polymerization (data

not shown).

3.2.5 Effect of Benzaldehyde-Capped Aminosilanes on ATRP Rate

The thicknesses produced using free and benzaldehyde-capped aminosilanes were

compared to determine whether a supposed organization of aminosilanes prior to ATRP would

cause a change in the ATRP rate. No significant change in polymerization rate was observed.

Furthermore, it was noted that the polymerization rates obtained from the BZ-APTMS films

were more reproducible than the ones collected from APTMS films (A.2). Rates of 17.2 and

17.8 nm h-1 were obtained for the BZ-APTMS surfaces. APTMS surfaces had rates of 12.8 and

20.2 nm h-1. AFM data indicated that BZ-APTMS surfaces (Fig. 3.6c) were smoother than the

APTMS surfaces (Fig. 3.6b). This may explain the higher polymerization rate reproducibility

observed for BZ-APTMS films. Such flat surfaces can be extremely advantageous in building

110

highly reproducible environments for the purpose of tailoring biosensing selectivity. The linear

growth rate can be used to estimate the length of polymerization time needed to achieve a desired

film thickness.

3.2.6 Analytical Performance for Biosensor Development

3.2.6.1 Immobilization of Oligonucleotide Probes

The oligonucleotide sequence used was that of the survival motor neuron gene, SMN1,

which is associated with spinal muscular atrophy (SMA) [7]. Only one sequence was studied but

it seemed to have behaved as predicted by theory. Other published work from our group

indicated that this sequence is representative and is unknown to have secondary structures. For

example, other members of our group have also evaluated our biosensing platforms using the

uidA gene [7, 47] and the lacZ gene [174] from Escherichia coli.

The SMN1 probe was purchased as a disulfide-modified sequence. Therefore, reduction

to thiols was required to allow its conjugation with the maleimide group in sulfo-SMCC. In our

experiments, the cleavage was done in water prior to purification in a Sephadex column as well

as prior to conjugation in 1× PBS (pH 7.0). The concentration of TCEP was in excess in both

cases. It is known that an excess of TCEP can directly react with maleimide, causing a reduction

in reactivity of the latter with thiolated compounds, such as thiol-modified oligonucleotides

[175]. It was also found to co-elute with reduced thiols in common gel filtration columns. In

order to minimize the amount of TCEP in the final thiol-modified oligonucleotide solution prior

to conjugation, Shafer et al. suggested using 0.5 equivalents of TCEP. Working at pH values of

6 and 7.3 also seemed to increase the reactivity between maleimide and a thiolated compound

(over 80% reactivity for both pH values) compared to that between TCEP and maleimide (40 and

111

50% reactivity) [175]. A neutral pH was chosen for our conjugation protocol, but an excess of

TCEP will be avoided in the future.

The order of immobilization of the various components was examined. ATRP was done

prior to oligonucleotide immobilization to avoid the acidic step required for the hydrolysis of the

imine bond and the basic step required for the immobilization of the initiator. However,

immobilization efficiency proved to be low (data not shown), and hybridization signals from

Cy3-labeled targets were not observed. Films were subsequently prepared in the order of

oligonucleotide probe immobilization, followed by ATRP and then DNA hybridization. ATRP

still occurred in the presence of DNA, which agreed with the data collected by Lou and He, who

even showed that the presence of DNA can accelerate the ATRP rate [173].

The oligonucleotide probes do not appear to be affected by the acidic treatment in the

benzaldehyde hydrolysis step; after this step, the Cy3 labeled oligonucleotide probes were still

present as seen in fluorescence images, and hybridization still occurred. The following

discussion compares results related to wettability, signal-to-background ratio, yield and

efficiency, and melt curve analysis between surfaces of oligonucleotides immobilized on

APTMS coating which did and did not undergo ATRP of HEMA. Reusability of the mixed film

surface will also be noted. Multiple probe and control areas were introduced onto slides to allow

for parallel processing of experiments.

3.2.6.2 Comparison of Mixed films versus Oligonucleotide Films

Upon rinsing the two different surfaces (Fig. 2.5 and 2.10) with water following

hybridization with oligonucleotide targets (Sequence 5 and 7, Table 2.1), there was an obvious

112

difference in wettability. Water spots were always observed on the pure oligonucleotide film

due to the difference in hydrophilicity in the oligonucleotide probe spots compared to the

surrounding, more hydrophobic BZ-APTMS and APTMS film. On the mixed oligonucleotide

and PHEMA film, however, the entire surface had the same wettability with no obvious water

beads. This provides evidence that ATRP has occurred within and outside of the oligonucleotide

probe spots since the pendant group in PHEMA is hydrophilic. In a separate experiment, the

wetting angle was observed to decrease as the polymerization time of PHEMA increased (data

not shown).

The ratio of signal versus background was compared between surfaces of oligonucleotide

probes that were immobilized on just APTMS or a mixture of BZ-APTMS and APTMS (Fig.

2.5), and those that were co-immobilized with PHEMA (Fig. 2.10). Signal intensity was based

on Cy3 fluorescence from labeled fully complementary targets (Sequence 5, Table 2.1). The

signal-to-noise ratio observed on APTMS surfaces was 2 ± 0.4, and the ratio on mixed APTMS

and BZ-APTMS surfaces was 2.1 ± 1.7, which was statistically identical. For the mixed

oligonucleotide and PHEMA film, however, a ratio of 8.5 ± 3.1 was observed. Adsorbed

oligonucleotides were not easily removed from pure APTMS coatings even in denaturing

conditions such as high temperature and 95% formamide in TE buffer. Adsorbed

oligonucleotide targets on APTMS and BZ-APTMS slides could be removed by heating in water.

A minimal amount of DNA adsorption was observed on PHEMA-coated slides, which agreed

with previous studies about various PHEMA substrates [176, 177].

A comparison of immobilization yield and hybridization efficiency on the two different

surfaces is presented in Table 3.4. Given the large deviation (no trials can be rejected), the

density of oligonucleotide probes within PHEMA-grown surfaces and on the bare aminosilane

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surfaces is similar, but the percent immobilization yield indicates a slightly higher yield of

oligonucleotides immobilized within PHEMA, even though the immobilization occurred before

ATRP and on the same aminosilane surface. However, hybridization efficiencies greater than

100% were determined for the pure oligonucleotide films. This can be attributed to a higher

level of adsorption observed on the mixed aminosilane surfaces. The mixed film, however, gave

reasonable hybridization efficiencies, with that of the fully complementary targets being much

higher than the 3 bpm targets.

Table 3.4: Densities, immobilization yield and hybridization efficiency of oligonucleotide probes and targets on

oligonucleotide film and mixed film surfaces.

Oligonucleotide films Mixed films

Density of OligoNT

(number of strands

per cm2)

% Immobilization yield

or % hybridization

efficiency

Density of OligoNT

(number of strands

per cm2)

%

Immobilization

yield or %

hybridization

efficiency

Probes 9.9 × 109 ± 8.1 × 1010 0.08 ± 0.60 % 3.7 × 1011 ± 1.8 × 1011 2.7 ± 1.1 %

0 bpm 4.6 × 1011 ± 3.3 × 1011 >100 % 2.7 × 1011 ± 3.5 × 1011 71 ± 1 %

3 bpm 2.7 × 1011 ± 2.1 × 1011 >100 % 5.8 × 1010 ± 4.0 × 1010 16 ± 0.8 %

OligoNT = oligonucleotides. Immobilization yield was calculated from the number of immobilized oligonucleotide probes

divided by the loading concentration of the probes. Hybridization efficiency was calculated from dividing the number of

oligonucleotide targets by the number of oligonucleotide probes.

PHEMA-modified surfaces were tested for reusability. Fully complementary targets

were introduced (Sequence 5 of Table 2.1) and then the hybrids were denatured, and new targets

were hybridized to the surface again. The process was repeated several times as depicted in

114

Figure 3.10. Hybridizations # 4 and 5 used three base-pair mismatch (bpm) targets (Sequence 7),

which resulted in the lower fluorescence intensity signals because of lower hybridization

efficiency. Aside from the first denaturation step, which could only remove about 50% of the

hybridized targets, the rest of the hybridized target signal could be removed, and that the

immobilized probes were able to hybridize again with new targets.

1-hyb 1-den 2-hyb 2-den 3-hyb 3-den 4-hyb 4-den 5-hyb0.00

1.00x107

2.00x107

3.00x107

4.00x107

Fluo

resc

ence

inte

nsity

(A.U

.)

Cycle

Fig. 3.10: Reusability of mixed films of oligonucleotides and PHEMA-modified glass slide for hybridizations. “1-

hyb” indicates cycle 1-hybridization, and “1-den” indicates cycle 1-denaturation. Note that 3 bpm targets were used

for hybridizations 4 and 5 whereas fully complementary SMN targets were used for hybridizations 1–3.

The melt curve experiments were done in 0.5× PBS and targets (fully complementary

(Sequence 5) and three base-pair mismatch (Sequence 7)) were hybridized with the surface

oligonucleotide probes prior to melt curve experiments, and representative melt curves are

shown in Fig. 3.11 and 3.12, respectively. As one can see, we have demonstrated that the shape

of the melt curve is similar to that of the curves seen in Piunno et al, which were collected from

115

DNA-immobilized substrate continuously exposed to buffer to ensure equilibrium conditions are

satisfied [159]. Therefore, the graphs shown in Fig. 3.11 and 3.12 reflect the melting process of

double-stranded DNA. Further proof is found in the control spots, where Cy3-labeled SMN

target sequences (Sequence 3) were immobilized using the same chemistry as the unlabeled

SMN probes (Sequence 1). Throughout the melting experiment, the fluorescence intensities

remained similar, but those from the Cy3-labeled targets (Sequences 5 and 7) decreased with

temperature.

20 30 40 50 60 70 80

0.0

0.2

0.4

0.6

0.8

1.0

f ssD

NA

Temperature ( oC)

Fig. 3.11: Melt curves of fully complementary targets collected from: (■) oligonucleotide film only; (▲) solution

(0.5 × PBS); (●) mixed film of oligonucleotides and PHEMA. “fssDNA” represents fraction of single-stranded DNA.

116

20 30 40 50 60 70 80

0.0

0.2

0.4

0.6

0.8

1.0

f ssD

NA

Temperature ( oC)

Fig. 3.12: Melt curves of 3 bpm targets collected from: (■) oligonucleotide film only; (▲) – solution (0.5 × PBS);

(●) mixed film of oligonucleotides and PHEMA. “fssDNA” represents fraction of single-stranded DNA.

Sharper slopes and greater Tm difference between the fully complementary target and

mismatched target are the two key factors that represent improvement of selectivity. The

average Tm and slope values are listed in Tables 3.5 and 3.6, respectively. The general trend is

that the Tm shifts to lower temperatures when hybridization occurs on a surface. In bulk

solution, the Tm for the fully complementary SMN hybrid is 62.5 ± 0.1°C and this value shifted

to an average of 49.3 ± 1.4°C on a mixed film surface (Fig. 2.10); there is a 13.2°C difference.

Using just pure oligonucleotide films (Fig. 2.5) yielded an average of 44.9 ± 0.8°C. Precision of

Tm values were much better on mixed film surfaces, consistent with a sharper transition. The

same trend was observed for 3 bpm targets. The correlation coefficients (CV) were less than

6.5% for the oligonucleotide films and less than 6.1% for the mixed films.

The differences in Tm values between fully complementary and 3 bpm targets were

compared between the pure oligonucleotide films (Fig. 2.5) and mixed oligonucleotide and

117

PHEMA films (Fig. 2.5). Overall, the Tm differences on oligonucleotide films were on average

2-fold smaller than those observed on the mixed film surfaces (Table 3.5), which was determined

at a 95% confidence level. The average Tm difference on pure oligonucleotide films was 4.5 ±

1.6 °C and on mixed films, it was 10.6 ± 0.9 °C, which is statistically identical to that obtained

from bulk solution.

Table 3.5: Tm values of fully complementary and 3 bpm SMN targets on mixed film and oligonucleotide film

immobilized on glass slides.

Triala Tm of fully complementary

targets (ºC)

Tm of 3 bpm targets (ºC)

ΔTm at 95% C.I.e

Oligonucleotideb film only

1 45.6 ± 1.7 42.8 ± 0.8 2.9 ± 1.9

2 44.1 ± 2.9 41.0 ± 2.4 3.1 ± 1.3

Avg 44.9 ± 0.8 41.9 ± 0.9 4.5 ± 1.6

Mixed filmc 1 52.7 ± 1.0 42.4 ± 1.3 10.3 ± 1.0

2 47.6 ± 2.0 38.5 ± 1.4 9.1 ± 1.7

3 46.9 ± 1.7 37.5 ± 1.5 9.4 ± 0.6

4 52.9 ± 1.7 38.8 ± 1.7 14.0 ± 0.6

5 46.5 ± 2.8 36.3 ± 1.4 10.2 ± 1.2

Avg 49.3 ± 1.4 38.7 ± 1.0 10.6 ± 0.9

Bulk solutiond 1 62.5 ± 0.1 (n = 1) 53.1 ± 0.1 (n = 1) 9.4 ± 0.1

a One trial represents one glass slide with all the usable spots (out of 8 spots) that exhibited a sigmoidal function with an R2 > 0.95 on the same slide. At least 3 replicates were used, unless otherwise noted.

b SMN probes were immobilized on a 1:1 ratio of APTMS and BZ-APTMS as depicted in Figure 2.5

c Mixed film composed of oligonucleotides and PHEMA immobilized on APTMS/BZ-APTMS-modified surfaces as depicted in Figure 2.10.

d The solution melt curve was collected in 0.5× PBS.

e The confidence interval (C.I.) at 95% was constructed for the difference in Tm values between fully complementary and 3 bpm targets.

± Standard deviations were reported. Avg: Average values. The averages of all experiments were reported with their standard errors.

118

An examination of the slopes of the denaturation profiles of the fully complementary

SMN targets on mixed film surfaces compared to films composed only of oligonucleotides

confirmed the sharper melting transition observed on surfaces that had undergone ATRP (Table

3.6). A smaller dx value corresponds to a sharper slope because dx represents the change in x

(temperature) for the largest change in y (fraction of ssDNA). The 3-fold increase in steepness

for the melt curves collected from mixed films that were hybridized with fully complementary

targets is very clear; it may suggest that the local environment of oligonucleotide probes is more

controlled and homogeneous for the mixed film surfaces. Another example of the increase in

steepness of slope, possibly due to PHEMA, is further reflected in comparisons of slope values

for the 3 bpm targets as seen in Table 3.6. There was a 1.5-fold increase in sharpness observed

for the 3 bpm targets when the melt curves collected from mixed films were compared to the

oligonucleotide films. The dx values obtained from the mixed films were equal to or smaller, i.e.

sharper melting transitions, than those obtained from bulk solution. The oligonucleotide films

had much greater variability in the dx values than the mixed films, particularly for melt curves of

fully complementary targets.

119

Table 3.6: dx values of fully complementary and 3 bpm SMN targets on mixed films and oligonucleotide films

immobilized on glass slides.

Triala dx valuee of fully complementary targets

(ºC)

dx value of 3 bpm targets (ºC)

Oligonucleotideb film only

1 5.8 ± 0.6 3.4 ± 0.8

2 7.5 ± 1.1 3.0 ± 1.3

Avg 6.6 ± 0.8 3.2 ± 0.2

Mixed filmc 1 2.3 ± 0.4 1.2 ± 0.5

2 2.5 ± 0.8 1.5 ± 0.1

3 2.8 ± 0.7 2.2 ± 0.6

4 2.1 ± 0.4 3.1 ± 0.6

5 4.2 ± 1.6 2.5 ± 0.4

Avg 2.8 ± 0.4 2.1 ± 0.3

Bulk solutiond 1 2.9 ± 0.1 3.0 ± 0.1

a One trial represents one glass slide with all the usable spots (out of 8 spots) that exhibited a sigmoidal function with an R2 > 0.95 on the same slide. At least 3 replicates were used, unless otherwise noted.

b SMN probes were immobilized on a 1:1 ratio of APTMS and BZ-APTMS as depicted in Fig. 2.5.

c Mixed film composed of oligonucleotides and PHEMA immobilized on APTMS/BZ-APTMS-modified surfaces as depicted in Fig. 2.10.

d The solution melt curve was collected in 0.5× PBS.

e dx value (obtained from sigmoidal curve fitting) represents the change in temperature for the greatest change in the fraction of ssDNA

± Standard deviations were reported. The averages of all experiments were reported with their standard errors.

Avg: Average values

Overall, the melting temperature and slope data both suggest that the melt curves reflect

similar general energetics for hybridization, but a narrower distribution of energetics for the case

of the mixed film coating. This may be due to the effectiveness of PHEMA coating in

120

controlling the local environment of oligonucleotide probes. These data are certainly very

encouraging, suggesting greater selectivity than observed for films that are composed of only

immobilized probes. It appears that the PHEMA matrix exerts a destabilizing force towards

targets that contain mismatches.

3.2.7 Conclusions

The use of a series of surface analysis methods revealed the success of each

immobilization step including silane and initiator immobilization and surface-initiated ATRP.

The use of BZ-APTMS was superior in the control of the reproducibility of the polymerization

rate. It provided a smoother surface for polymerization as well as served as a temporary

protecting group for immobilized amines for subsequent reaction steps. HEMA was shown to be

the most useful material for development of biosensing surfaces. It dramatically reduced non-

specific adsorption, allowed reusability, sharpened the melt curves for both fully complementary

and 3 bpm targets, and increased resolution of such targets. Further experiments will be done to

explore effects of solution ionic strength, mismatch location and tuning for SNP analysis. The

results of this work support the hypothesis that selectivity of hybridization can be improved by

narrowing the distribution of energetics experienced by immobilized DNA hybrids.

121

3.3 Bin+ Cluster Ion Sources for Investigation of a Covalently-Immobilized Mixed Film Composed of Oligonucleotides and Poly(2-hydroxyethyl methacrylate) Brushes

April K.Y.Wonga, R.N.S. Sodhib, and Ulrich J. Krulla*

aChemical Sensors Group, University of Toronto Mississauga, 3359 Mississauga Rd. N.,

Mississauga, ON, Canada L5L 1C6

bSurface Interface Ontario, Department of Chemical Engineering and Applied Chemistry,

University of Toronto, On, Canada M5S 3E5

Contributions: April Wong performed all measurements and data analysis and interpretation

under the direction of Ulrich Krull. Dr. Rana Sodhi and Peter Brodersen of Surface Interface

Ontario collected the XPS and ToF-SIMS data.

Submitted part of this chapter to Surface and Interface Analysis, special conference proceedings

issue for the SIMS XVII conference (September 2009).

122

3.3.1 Abstract

The covalent co-immobilization of short non-nucleic acid oligomers of poly(2-

hydroxyethyl methacrylate) (PHEMA), which are grown by surface-initiated atom transfer

radical polymerization, with oligonucleotides on glass surfaces has been demonstrated to

significantly improve selectivity of nucleic acid hybridization for applications involving

microarrays and biosensors [178]. Physical characterization is required to examine how the two

components localize within the same area on a glass surface. Glass slides coated with

immobilized spots of oligonucleotide probes of 20mer length at a density of ~3.7 × 1011

molecules cm-2, and PHEMA of nominal thickness of less than 10 nm, were analyzed using static

time-of-flight secondary ion mass spectrometry (TOF-SIMS). The samples were subjected to a

polyatomic bismuth cluster ion source for the TOF-SIMS experiment. In the positive ion

spectra, fragments of PHEMA and nucleosides were observed. In the negative ion spectra,

PHEMA as well as signature fragments from nitrogenous bases, phosphate and phosphite groups

were identified. Comparison of negative ion spectra indicated that the Bi5+ source provided

details that were either not apparent or at much reduced sensitivity when using Bi3+ and Bi+

sources. Bi3+ provided higher secondary ion yields for positive ion fragments, and more than

200-fold selectivity for ionization of oligonucleotide in comparison to PHEMA when in mixture.

3.3.2 Introduction

A reduction in the ensemble of degrees of freedom of immobilized oligonucleotide

probes can be achieved by forming a mixed film composed of oligonucleotide probes and

poly(2-hydroxyethyl methacrylate) (PHEMA) brushes, which are co-immobilized and grown

from the surface to maximize the surface density [178]. Melt curve data based on fully

123

complementary and three base-pair mismatched targets have demonstrated that the presence of

PHEMA can enhance selectivity, as evident from the sharpness of the DNA melting transition

and increases of the difference between the Tm values [178].

X-ray photoelectron spectroscopy (XPS), time-of-flight secondary ion mass spectrometry

(TOF-SIMS), atomic force microscopy (AFM) and ellipsometry have previously been used to

ensure that the surface-initiated polymerization reaction of PHEMA had proceeded on silicon

wafers [178]. However, structural details about how oligonucleotides and PHEMA co-exist on

surfaces are lacking. Such data would be useful to improve the design of immobilization, and

therefore to optimize selectivity. As one step towards this goal we have used TOF-SIMS to

investigate oligonucleotides that were co-immobilized with PHEMA brushes. An ion source that

produced monoatomic and polyatomic bismuth was investigated based on the ability of such a

system to generate increased secondary molecular and fragment ion yields [179, 180].

Furthermore, data from this TOF-SIMS experiment was contrasted with data from XPS. SEM

was used to estimate the density of immobilized oligonucleotide probes using oligonucleotide

targets which were labelled with Au nanoparticles (nps).

The XPS and TOF-SIMS experiments used surfaces containing APTMS (Surface A),

oligonucleotides only (Surface B), mixed oligonucleotides and PHEMA (Surface C), and

PHEMA only (Surface D). Oligonucleotides were deposited as 2 mm spots on predetermined

areas on Surfaces B and C. The success of the immobilization of oligonucleotides was verified

by detecting the fluorescence emission from Cy3-labeled oligonucleotides using a confocal

scanner. PHEMA was grown from the surface on Surfaces C and D. Surface-initiated

polymerization was confirmed using ellipsometry, which indicated an increase in the apparent

thickness on silicon wafer samples that were exposed to the ATRP reaction mixture. The total

124

change in thickness was between 3 to 6 nm (determined by ellipsometry). For characterization

by SEM, the Au np-oligonucleotide conjugates and PHEMA were immobilized across fused

silica surfaces.

3.3.3 XPS Characterization of Mixed Film on Glass Surfaces

The four sample surfaces were investigated by angularly-resolved XPS using 30° as the

take-off angle. It is known that the smaller the angle (<45°), the more sensitive the XPS

instrument is in terms of detection of electrons from species of the uppermost sample layer [181].

Figure 3.13 shows the high resolution C 1s spectra of all four surfaces. The C 1s signatures of

PHEMA are not present in Fig. 3.13a-b but are very evident in the spectra shown in Fig. 3.13c-d.

The peaks at 289.0 eV, 287 eV, and 286 eV are representative of the ester carbon, carbon-

oxygen, and carbon bonded to a carbonyl, respectively [182]. These peaks are only present on

the mixed and PHEMA-modified surfaces (Surfaces C and D), demonstrating that PHEMA was

grown from the surface via the ATRP reaction. These high resolution spectra agreed with

previously collected spectra of PHEMA films alone [178].

125

(a) b)

(c) (d)

Fig. 3.13: High resolution C 1s spectra of all surfaces (A-D) investigated; (a) Surface A: aminosilanes only; (b) Surface B:

oligonucleotides only; (c) Surface C: mixed film; (d) Surface D: PHEMA only

Signals associated with atoms indicative of oligonucleotides were not as clear. There are

two types of carbon species which are unique to oligonucleotides. One is the carbon which is

singly-bound to nitrogen (C-N or N-C-N). The XPS signal is located at 286-287 eV [183]. The

other is the amide carbon, which should be at 288 eV [183]. Figure 3.13b has C 1s signatures at

these binding energies. Compared to Figure 3.13a, the increases in signals observed in Figure

0

50

100

150

200

250

282284286288290292294296298

Cou

nts

/ s

Binding Energy (eV)

C1s hr100 Scans, 15 m 55.0 s, 300µm, CAE 30.0, 0.10 eV

Angle = 30 °

C1sC1s hr A

C1s hr B

C1s hr C

C1s hr D

10

20

30

40

50

60

70

80

90

100

281282283284285286287288289290291292293294295296297298299300

Cou

nts

/ s

Binding Energy (eV)


Angle = 30 °

C1s

C1s hr A

C1s hr B

C1s hr C

C1s hr D

20

40

60

80

100

120

140

160

180

200

220

282284286288290292294296298

Cou

nts

/ s

Binding Energy (eV)


Angle = 30 °

C1s

C1s hr A

C1s hr B

C1s hr C

C1s hr D

10

20

30

40

50

60

70

80

90

282284286288290292294296298300

Cou

nts

/ s

Binding Energy (eV)


Angle = 30 °

C1s

C1s hr A

C1s hr B

C1s hr C

C1s hr D

126

3.13b are marginal. Figure 3.13c also has C 1s signatures at 288 eV and at 286 eV. However,

the latter peak may be specific to polar carbon species from PHEMA, as it is also observed in

Fig. 3.13d. The atomic percentages of each nitrogenous carbon species for Surfaces A to C are

listed in Table 3.7. One can see that from visual inspection of the fitting and from the table, the

areas of the peaks of interest are very similar from one sample to the next. There is only a slight

increase in the atomic percentages of the carbon species at ~286 eV on Surface C, but the

increase could be contributed to the polar carbon groups originating from PHEMA.

Table 3.7: Atomic percentages of nitrogen-containing carbon species unique to oligonucleotides on Surfaces A to C collected at

a take-off angle of 30° measured by XPS.

Surface Trial Atomic percentage of

carbon singly-bonded

to nitrogen at 286-287

eV (%)

Atomic percentage of

amide carbon at 288

eV (%)

A 1 11.13 4.15

2 15.02 3.99

B 1 15.09 6.36

2 14.24 4.09

C 1 17.59 5.72

2 17.67 4.19

Surface A: aminosilanes only; Surface B: oligonucleotides only; Surface C: mixed film.

The C-H and C-C peaks at 285 eV are present on all surfaces, as expected, due to

adventitious carbon atoms [92] and aliphatic carbon atoms from oligonucleotides and PHEMA.

One other expected peak associated with oligonucleotides would be located at 289 eV, which is

the urea carbon [N-C(=O)-N] [183]. There is a peak at this latter binding energy, but this signal

127

could come from the ester carbon of PHEMA. This would be consistent with the apparent

dominance of PHEMA C 1s signatures. The P 2p and N 1s spectra were also collected, but the

total atomic percentages were extremely low (<4.5% for nitrogen and < 0.3% for phosphorus).

These percentages did not correspond to the theoretical relative abundances with respect to

carbon and oxygen in nucleic acids. These XPS spectra show that PHEMA is certainly detected,

but solid evidence for the presence of oligonucleotides from XPS data is lacking.

3.3.4 Estimation of Density of Surface Immobilized Oligonucleotide Probes by Using Au Nanoparticle-Tagged Complementary Oligonucleotide Targets

In the work conducted by May et al., XPS signals from DNA were stronger with respect

to the background level [183]. However, nucleobases were analyzed in powder form, which

most likely provided a greater amount of sample for analysis. To investigate whether a low

density of oligonucleotides was the reason why XPS was not sufficiently sensitive enough to

detect signals indicative of the constituents of DNA, the surface density of immobilized

oligonucleotide probes was determined. The method devised to determine the surface density

was based on use of oligonucleotide targets that were labelled with gold nanoparticles (Au np).

Hybridization to surface-immobilized probes was an indicator of probe availability as has been

demonstrated by Csáki et al. [184].

The successful preparation of Au np-oligonucleotide conjugates was confirmed by an

absorption peak at 260 nm for nucleic acids [19] and at 520 nm for the Au nanoparticle plasmon

band [138]. A TEM image after DNA conjugation shows the expected size for these Au

nanoparticles, and also the absence of aggregation (Fig. 3.14). Further evidence included

128

measurement of the fluorescence emission from Cy3-labelled oligonucleotides (λmax = 564 nm),

which were immobilized onto Au np. A similar experiment was demonstrated previously by

Demers et al. [138]. Fluorescence emission from dyes attached to oligonucleotides which are

immobilized onto Au np is generally not observed due to FRET and a strong absorption

capability of the excitation energy by nanoparticles [138]. Dissolution of the Au nanoparticles

by KCN can recover the fluorescence emission [138] since Au0 is essentially converted to

[Au(CN)4]- [185]. Gold ions no longer have the quenching ability of gold nanoparticles [138].

The disappearance of a conduction band inherent with a solid nanoparticle means the absence of

plasmon oscillations that dictate optical properties, hence the inability to absorb emission from a

closely located dye [186]. Fig. 3.15 shows that fluorescence emission of Cy3 of immobilized

probes on Au nanoparticles was not observed before the addition of KCN, but it was recovered

after solubilization of the gold nps. The dissolution of gold proceeds quickly and immediate

colour change from a faint red colour to light yellow within seconds was observed.

Fig. 3.14: TEM of 5 nm gold nanoparticles after conjugation with thiolated SMN probes.

129

540.00 560.00 580.00 600.00 620.00 640.000.00

1.00x105

2.00x105

3.00x105

4.00x105

5.00x105

6.00x105

7.00x105

Fluo

resc

ence

inte

nsity

(A.U

.)

Wavelength (nm)

Fig. 3.15: Fluorescence intensity of Cy3-labeled thiolated SMN probe immobilized onto 5 nm gold nanoparticles (-

- -) before addition of 40 mM KCN and (⎯) after addition of KCN. λex = 520 nm.

By using the peak fluorescence intensity at 562 nm recovered after the etching procedure

and subtracting it from the intensity before etching, the number of oligonucleotides immobilized

onto each Au nanoparticle was calculated. An equation was constructed from calibration

experiments:

9962.0,433504.6311 2 =−= RCI , (3.3)

where I is the peak intensity at 562 nm and C is the oligonucleotide concentration in nM. The

ratio calculated was two Au nps to one oligonucleotide. This finding is consistent with the

dimensions of the maximum diameter of a double-stranded DNA (2 nm) [19], and issues of steric

hindrance. Furthermore, sequence-specific adsorption of oligonucleotides to the Au

nanoparticles can also influence surface coverage [187, 188]. The data suggested that each

130

nanoparticle carried either one oligonucleotide or no oligonucleotides (on average). The number

of Au nanoparticles observed in hybridization experiments should therefore be equal to the

number of actual immobilized oligonucleotide probes.

The surface density of the immobilized probes was estimated by counting the number of

Au nanoparticles observed in a SEM image. Fig. 3.16 shows the distribution of Au nanoparticles

on PHEMA-covered fused silica slides with immobilized oligonucleotide probes (Fig. 3.16a) and

without (Fig. 3.16b) immobilized oligonucleotide probes. The SEM images again show that the

nanoparticles were of the expected diameter of 5 nm. Assuming that each Au np-oligonucleotide

target hybridized with an oligonucleotide probe immobilized on the surface, the resultant density

was about 9 × 1010 (± 8%) Au nanoparticle·cm-2. This theoretically corresponded to the same

density of oligonucleotide probes. With consideration of some adsorption (Fig. 3.16b), the

adjusted density was slightly lowered to 7 × 1010 (± 8%) oligonucleotide probes·cm-2. Structural

features observed in Fig. 3.16a were possibly due to PHEMA and the bright clusters could be

aggregates of Au nanoparticles.

131

a)

132

b)

Fig. 3.16: (a) SEM image of Au np-oligonucleotide targets hybridized to surface-bound oligonucleotide probes on

PHEMA-modified surfaces; (b) SEM image of Au np-oligonucleotide targets adsorbed onto PHEMA-modified

surfaces (no oligonucleotide probes were immobilized).

Previously, we have used fluorescently labeled oligonucleotides to determine the surface

density of immobilized oligonucleotide probes within the mixed film to be 3.7 × 1011 (± 48%)

oligonucleotide probes·cm-2. The density of oligonucleotide probes determined from the Au np

method was 7 × 1010 (± 8%) oligonucleotide probes·cm-2, which is about an order of magnitude

away from the value determined by the fluorescence labeling method. These values are

133

comparable to the density value of 1.8 × 1011 probes·cm-2 that was considered to be low in earlier

work produced from our group, which was calculated from the number of oligonucleotide probes

and the surface area of controlled pore glass provided [129]. The low density of immobilized

oligonucleotide probes could be the reason for the absence of signals from oligonucleotides

observed in the high resolution XPS spectra for C 1s when using mixed films.

3.3.5 ToF-SIMS Characterization of Mixed Films on Glass Surfaces

TOF-SIMS has previously been used to characterize surfaces immobilized with nucleic

acids [180, 189, 190] and PHEMA [191-193]. In this work, a mixture of components was

characterized by TOF-SIMS using a bismuth cluster ion beam. Ionized fragments originating

from each nucleobase, nucleoside, and nucleotide have been identified by May et al. [183], but a

Cs+ ion source was used in their work. Evidence for the presence of PHEMA came from the

pendant group and backbone of PHEMA, which are represented by the C4H5O2- and the C2H5O+

fragments, respectively [192].

The same four types of surface (Surfaces A–D) that were characterized by XPS were also

analyzed by TOF-SIMS equipped with a polyatomic bismuth cluster ion beam. The relative

peak intensities of selected secondary ion fragments from both oligonucleotides and PHEMA

were compared between the surfaces, and comparison included consideration of various bismuth

cluster ion sizes with all experiments operated using the same current. All of the raw spectra at

both polarities are shown in, A.3 (Bi+), A.4 (Bi3+), and A.5 (Bi5

+). Figure 3.17 shows where a

few fragments associated with DNA may originate in the positive ion mass spectra. Evidence for

the presence of PHEMA came from the 2-hydroxyethyl pendant group, which is represented by

the C4H5O2- fragment [192], for example. Figure 3.18 shows a proposed fragmentation

134

mechanism of commonly observed positive and negative fragments from the backbone and

pendant group of PHEMA. Fig. 3.19 shows the overlaid ToF-SIMS spectra of selected ion

fragments.

(a) (b)

Fig. 3.17: Possible origin of (a) [C3H5NO2+H]+; (b) [C2HNO2+H]+ fragments.

Fig. 3.18: Proposed fragmentation of PHEMA for selected positive and negative fragments.

135

(a)

(b)

(c)

136

(d)

Fig. 3.19: Overlaid ToF-SIMS spectra when Bi3+ was used for the following ion fragments: (a) PO3

- at 78.96 m/z;

(b) C4H5O2- at 85.03; (c) C2H2NO2

+ at 71.96; (d) C2H5O+ at 45.02 m/z. Top to bottom: Surface A to D. The file

labels are incorrectly printed due to an internal printing error. Peak intensities are in counts.

In the negative ion spectra, PO3- was selected as a negative fragment associated with

oligonucleotides since it provided the most intense signal, which agreed with the work by

Hellweg et al. [190]. Figure 3.20a shows the signal intensity of PO3- using various bismuth ion

sources on different surfaces. This fragment should only appear on surfaces that contain

oligonucleotides. The Bi5+ ion cluster source generated excellent signal-to-noise ratio for the

PO3- fragment from both Surfaces B and C (Surfaces A and D served as controls in this case).

The peak intensity of the fragment found on Surface C using Bi5+ was five-fold higher in

intensity compared to using Bi3+ on the same surface. This trend was also observed when the

PO2- fragment was compared between the four surfaces and different bismuth cluster ion sizes.

The same pattern, but to a smaller extent, can be echoed for the C4H4N5- fragment, which is a

deprotonated adenine ion. However, the selectivity for the detection of this fragment on the

mixed film surface compared to other control surfaces was not as high.

137

(a)

n = 1 n = 3 n = 50.0

2.0x105

4.0x105

6.0x105

8.0x105

1.0x106

Peak

Inte

nsity

(cou

nts)

Bin+ cluster ion type

SurfaceA SurfaceB SurfaceC SurfaceD

(b)

n = 1 n = 3 n = 50.0

1.0x105

2.0x105

3.0x105

4.0x105

5.0x105

6.0x105

7.0x105

Pea

k in

tens

ity (c

ount

s)



Fig. 3.20: Peak intensities of (a) PO3- from nucleotides; (b) C4H5O2

- from PHEMA under various types of Bin+ (n =

1, 3, 5) primary ion source collected from various surfaces (A-D). Surface A: aminosilanes only; Surface B:

oligonucleotides only; Surface C: mixed film; Surface D: PHEMA only.

138

Evidence for the presence of immobilized PHEMA was detected readily (Fig. 3.20b)

using the Bi+, Bi3+, and Bi5

+ cluster ion sources by monitoring the peak intensity of the C4H5O2-

fragment. The peak intensities of this fragment were statistically significant only from PHEMA-

containing surfaces, which were surfaces C and D. The intensities of ion fragments when Bi3+

and Bi5+ ion sources were used produced twice the intensities of those obtained when Bi+ was

used.

The fragment C2H2NO2+ in the positive ion spectra came from the nucleosides of either

deoxycytidine or deoxythymidine. This fragment was found only on Surfaces B and C which

was consistent with the presence of oligonucleotides on these surfaces. Fig. 3.21a shows that the

Bi3+ cluster ion source provided secondary ion intensities of 1.5 to 2-fold higher for both

Surfaces B and C compared to using the Bi5+ ion source. There was a 20 to 40-fold increase in

the magnitude of the signals when the Bi3+ ion source was used compared to signals obtained

using the Bi+ ion source. The same trend was observed for the C3H6NO2+ ion, another possible

fragment from a nucleoside.

In Fig. 3.21b, the high intensity of the C2H5O+ fragment indicated the presence of the

PHEMA pendant group on Surfaces C and D. The Bi3+ cluster ion sources provided the highest

peak intensity of the fragment on Surfaces C and D.

139

(a)

n = 1 n = 3 n = 50.0

1.0x104

2.0x104

3.0x104

4.0x104

5.0x104

6.0x104

7.0x104

Pea

k in

tens

ity (c

ount

s)



(b)

n = 1 n = 3 n = 50.0

1.0x105

2.0x105

3.0x105

4.0x105

5.0x105

Pea

k in

tens

ity (c

ount

s)



Fig. 3.21: Peak intensities of (a) C2H2NO2+ from nucleosides; (b) C2H5O+ from PHEMA under various types of Bin

+

(n = 1, 3, 5) primary ion source collected from various surfaces (A-D). Surface A: aminosilanes only; Surface B:

oligonucleotides only; Surface C: mixed film; Surface D: PHEMA only.

140

Table 3.8: List of secondary ion fragments and peak intensities found by using Bi3+ as a cluster ion primary source

in TOF-SIMS for each type of surface.

Peak intensities found for each type of surface

Fragments m/z

ratio

Control

(Surface A)

DNA only

(Surface B)

Mixed film

(Surface C)

PHEMA only

(Surface D)

Oligonucleotides

PO2- 62.96 1.2 × 104 2.9 × 105 1.1 × 104 4.8 × 103

PO3- 78.96 3.1 × 104 9.6 × 105 3.9 × 104 1.5 × 104

C4H4N5-

(Adenine-H)

134.05 6.3 × 102 3.5 × 104 6.3 × 103 8.6 × 103

[C2HNO2 + H]+

(dC or dT)

71.96 4.9 × 103 5.3 × 104 7.1 × 104 1.4 × 104

[C3H5NO2 + H]+

(nucleoside)

87.95 3.7 × 103 3.0 × 104 3.1 × 104 2.6 × 103

PHEMA

C2H3O2- 59.01 1.4 × 104 1.5 × 104 3.2 × 105 2.1 × 105

C4H5O2- 85.03 4.2 × 103 3.1 × 103 7.1 × 105 7.3 × 105

C2H5O+ 45.02 2.1 × 104 3.2 × 104 5.3 × 105 4.8 × 105

C4H5O+ 69.02 1.4 × 104 1.6 × 104 4.3 × 105 3.1 × 105

Table 3.8 shows the selectivity of Bi3+ as a primary ion beam to ionize fragments

associated with oligonucleotides and PHEMA on different surfaces. Fragments unique to

oligonucleotides were observed on Surfaces B and C and those specific to PHEMA were

141

detected from Surfaces C and D at an intensity that was one to two orders of magnitude greater

than the respective control samples. The peak intensities of most secondary ion fragments

associated with oligonucleotides decreased when collected from Surface C (mixed film), as

opposed to Surfaces B (positive control). The relative intensities of the PHEMA fragments

stayed statistically similar between the mixed film and positive control surfaces (C and D). This

demonstrates that TOF-SIMS equipped with a bismuth cluster ion can selectively and

simultaneously detect ion fragments that are unique to two different components which co-exist

on the same surface.

Bi+ as a primary ion source was found to be inefficient in generating high secondary ion

yields (i.e. peak intensity) for fragments that are unique to oligonucleotides. Polyatomic bismuth

cluster ion sources (Bi3+ and Bi5

+) provided the highest secondary ion intensities for both

negative and positive ion species, which agrees with literature [179, 180, 189, 190]. The

selectivity for each fragment between the mixed film surface and the negative control surfaces

was also higher when the polyatomic bismuth cluster ion sources were used. Polyatomic cluster

ion sources have a higher probability of providing higher secondary ion yields because of

increased number of collision cascades near the surface [179], and this might translate into

increased secondary ion intensities. The secondary ion yield, which is defined as the number of

secondary ions divided by the number of primary ions, is in turn dependent on the amount of

primary ion energy which is deposited near the surface, and this relies on the mass and size of

the primary ion projectile [179, 189]. This enhancement is typically observed for substrates that

have insulating properties such as Si as reported by Bernhardt et al. [194], and this condition

might be expected for glass substrates as used in this work. Silicon surfaces have were found to

have higher scattering ion yields of antimony and bismuth cluster ions compared to conductive

materials such as gold and graphite [194]. These surfaces have a higher work function, hence

142

there is a high probability for cluster ions to be neutralized on the surface [194]. This in turn

reduces the number of cluster ions that can ionize surface species. However, the collision energy

used in this work was almost 200-fold higher than that used by Bernhardt et al. [194]. Another

contributing factor for the increase in secondary ion yield would be the nonlinear effects of

polyatomic cluster ion sources [195], besides the effects caused by the increase in mass of the

projectile [189]. Hellweg et al. have found a total enhancement factor of 31.1 and 10.4 per Bi

atom for the secondary ion yield of PO3- when the Bi3

+ cluster ion was used (compared to the Bi+

ion) for the analysis of an oligonucleotide-immobilized surface [189]. This nonlinear increase

was also observed for Bi52+ and Bi2

+ primary ion sources [189].

For the analysis of negative ion fragments related to oligonucleotides, only the Bi5+ ion

source was able to substantially increase the ion intensities from the low oligonucleotide density

of Surface C. The Bi3+ ion source generated higher secondary ion intensities from Surface B but

it was not sensitive enough to generate the same secondary ion fragments from Surface C.

However, the converse was true for the positive ion fragments. The Bi3+ ion source yielded

higher ion intensities of positive ion fragments that originated from both oligonucleotides and

PHEMA than the Bi5+ ion source. Therefore, the mechanism of dissociation for each primary

cluster ion and its interaction with surface species must be fundamentally different.

Nevertheless, these results support the fact that there is an increase in secondary ion intensities

when polyatomic cluster ion sources are used as primary ion sources [179].

Although some unexpected results were obtained for some secondary ion species when

using Bi5+ bombardment, it can be concluded that in general the secondary ion intensities were

maximized for both polarities when Bi3+ was used. The secondary ion yield was generally

maximized for both positive and negative ion cases when Bi3+ was used. The Bi3

+ ion source

143

was also observed to produce higher secondary ion yield than Bi5+ in the study by Touboul and

co-workers [180]. They pointed out that the velocities of various sizes of cluster ions, at the

same given kinetic energy, are inversely related to the square root of the number of substituents

in the cluster ion [180]. The decreased velocity may lead to lowered secondary ion generation

[179, 180].

One interesting observation worth noting is that the negatively charged oligonucleotide

fragments such as PO2- and PO3

- had comparable magnitudes in peak intensities as the negatively

charged PHEMA fragments (see Fig. 3.20 and Table 3.8), and only an order of magnitude lower

for positively charged oligonucleotide fragment when compared to positively charged PHEMA

fragments (see Fig. 3.21 and Table 3.8). The number of repeating units in oligonucleotides was

19 bases and PHEMA was about 20 to 40 units of monomer, with each repeating unit providing

one PO3- or C4H5O2

-, respectively. However, the relative densities on the surface was about

1:200 oligonucleotides:PHEMA, assuming that there were as many as 7 × 1013 polymer chains

cm-2 [136] on the surface and the density of oligonucleotides was determined to be about 3.7 ×

1011 molecules cm-2. The similarity in the magnitudes of the observed signal intensities for the

characteristic fragments suggests a significant selectivity for ionization of oligonucleotides, and

points to an unusual matrix effect.

144

(a)

C2H5O_n=3 C4H5O _n=3 C2H5O_n=5 C4H5O _n=5 0.0

1.0x105

2.0x105

3.0x105

4.0x105

5.0x105

6.0x105

7.0x105

8.0x105

9.0x105

1.0x106

1.1x106

Peak

inte

nsity

(cou

nts)

PHEMA Fragment and Bin+ (n=1, 3, 5) cluster ion used

Control conc1 conc2 conc3

(b)

C4H5N4_n=3 C6H7N2O_n=3 C4H5N4_n=5 C6H7N2O_n=5 0.0

5.0x104

1.0x105

1.5x105

2.0x105

2.5x105

3.0x105

Pea

k in

tens

ity (c

ount

s)

Oligonucleotide fragment and Bin+ (n=1, 3, 5) cluster ion used

Control conc1 conc2 conc3

Fig. 3.22: Comparison of peak intensities of selected positive fragments from PHEMA and oligonucleotides

between the Bi3+ and Bi5

+ primary ion sources at various concentrations; (a) C2H5O+ and C4H5O+ fragments from

PHEMA; (b) C4H5N4+ and C6H7N2O+ from oligonucleotides. The control was a mixed BZ-APTMS and APTMS;

conc1 was 0.10 μM, conc2 was 0.48 μM, and conc3 was 1.02 μM (loading concentrations).

145

In a separate experiment, a mixed film sample was prepared containing three different

surface densities of oligonucleotides, while keeping that of PHEMA constant. Figure 3.22 shows

the peak intensities of selected fragments from PHEMA and oligonucleotides at various PHEMA

to oligonucleotide ratio. The fragments C4H5N4+ (m/z = 109) and C6H7N2O+ (m/z = 111)

originated from adenine and thymine [183]. The Bi3+ cluster ion source provided higher peak

intensities than the Bi5+ source, which agreed with the first set of TOF-SIMS data (Fig. 3.20 and

3.21) and with those observed by Touboul et al. [180]. The density of the immobilized

oligonucleotide probes was at least 3.9 × 1011 (± 19%) probes·cm-2 by using a loading

concentration of 0.10 μM. This resulted in a PHEMA:oligonucleotide ratio of 181:1 by using a

density value of 7 × 1013 polymer chains·cm-2 for PHEMA. However, the peak intensities of

fragments associated with oligonucleotides were still high, and reached magnitudes about 2 to 7-

fold less than the peak intensities of PHEMA-related fragments.

Moreover, changing the oligonucleotide loading concentration from 0.1 μM to 0.48 μM

and 1.02 μM did not result in any changes in the peak intensities of all fragments. The

difference in surface density between the three loading concentrations was confirmed by

obtaining a confocal image of the surface. The integrated fluorescence intensity ratio collected

from each Cy3-labelled oligonucleotide-spotted area of a different loading concentration was

1:5:10 (0.1 μM:0.48 μM:1.02 μM). Therefore, the relative ratios of the number of immobilized

oligonucleotide probes from each area were the same as that of the loading concentrations. Since

changing the number of immobilized probes did not change the secondary ion intensities of

fragments from both PHEMA and oligonucleotides components, the formation of fragments may

have reached a saturated level. Another observation was that fragments that originated from the

nucleobases were detected this time but not the fragments from the phosphate backbone.

146

Besides fragments from thymine and adenine, fragments from cytosine, C4H6N3O+ (m/z = 112),

and guanine, C5H3N4O+ (m/z = 135) [183], were also readily identified in the positive ion mass

spectra. This demonstrates that any slight changes in the matrix may influence the yield of

particular fragments.

The comparison of data collected from both XPS and TOF-SIMS demonstrated the

superior sensitivity of TOF-SIMS equipped with a polyatomic bismuth cluster primary ion to

detect fragments from DNA (present at a low density) within a polymer matrix. Although XPS

provides chemical information of surface species such as oxidation states, type of bonding, and

environment of each element, it is known not to be as sensitive as SIMS [196]. The detection of

fragment ions from oligonucleotides (19 mer) at a low density within a matrix of PHEMA of less

than 10 nm in theoretical length, reiterates the superior capability of TOF-SIMS to investigate

such a unique surface.

3.3.6 Conclusions

A novel biosensing surface composed of oligonucleotides and poly(2-hydroxyethyl

methacrylate) (PHEMA) brushes that co-existed as a thin mixed film on silica substrates was

investigated by XPS, SEM, and TOF-SIMS. XPS was not able to detect oligonucleotides but

could detect PHEMA readily. This was in part due to the lower sensitivity of XPS and a low

density of oligonucleotides immobilized on the surface. TOF-SIMS with a bismuth cluster ion

source could selectively detect fragments unique to each component of the mixture. The

secondary ion intensities were generally maximized for both positive and negative ion fragments

when Bi3+ was used, which can be related to a higher velocity compared to a larger cluster ion at

the same kinetic energy, besides the well known enhancement in secondary ion yield provided

147

by cluster ions and their non-linear effects. There is evidence of a selective ionization for

oligonucleotides since the secondary ion intensities of ion fragments coming from both

components were similar in magnitudes even though the ratio of the density of oligonucleotides

to that of PHEMA was calculated to be 1:200. Therefore, the use of polyatomic bismuth cluster

ion sources in TOF-SIMS offered a sensitive and selective tool to detect oligonucleotides in a

polymeric environment.

148

3.4 A Mixed film Composed of Oligonucleotides and Poly(2-Hydroxyethyl Methacrylate) Brushes to Enhance Selectivity for Detection of Single Nucleotide Polymorphisms

April K.Y.Wonga,c, Denys O. Marushchakb,c, Claudiu C. Gradinarub,c and Ulrich J. Krulla,c*

aDepartment of Chemistry, University of Toronto, Toronto, Canada

bDepartment of Physics, Institute for Optical Sciences, University of Toronto, Toronto, Canada

cDepartment of Chemical and Physical Sciences, University of Toronto Mississauga, 3359

Mississauga Rd. N., Mississauga, ON, Canada L5L 1C6

Contributions: April Wong performed all measurements, except for the lifetime data, which

were contributed by Denys Maruschchak and Claudiu Gradinaru. All other data analysis and

interpretation were done by April Wong under the guidance of Ulrich Krull.

Accepted in Anal. Chim. Acta (2009), doi:10.1016/j.aca.2009.12.001

149

3.4.1 Abstract

Preliminary studies of mixed films composed of oligonucleotides and poly(2-

hydroxyethyl methacrylate) (PHEMA) have recently been shown to enhance the selectivity for

detection of 3 base-pair mismatched (3 bpm) oligonucleotide targets. Evaluation of selectivity

for detection of single nucleotide polymorphisms (SNP) using such mixed films has now been

completed. The selectivity was quantitatively determined by considering the sharpness of melt

curves and melting temperature differences (ΔTm) for fully complementary targets and SNPs.

Stringency conditions were investigated, and it was determined that the selectivity was

maximized when a moderate ionic strength was used (0.1 to 0.6 M). Increases of ΔTm observed

when using mixed films were up to 3-fold larger compared to surfaces containing only

immobilized oligonucleotide probes. Concurrently, increases in sharpness of melt curves for 1

bpm targets were observed to be up to 2-fold greater for mixed films. The co-immobilization of

PHEMA resulted in a more homogeneous distribution of oligonucleotide probes on surfaces.

Lifetime measurements of fluorescence emission from immobilized oligonucleotide probes

labeled with Cy3 dye indicated the difference in microenvironment of immobilized

oligonucleotides in the presence of PHEMA.

3.4.2 Introduction

Single nucleotide polymorphisms (SNPs) are sites found in deoxyribonucleic acid (DNA)

which contain a single base-pair mismatch [9]. It is defined as such when the more common

sequence variant occurs less than 99% of the time [197]. In the human genome, a SNP occurs on

average once in every 1000 bases [197]. As the most abundant human genetic variation, SNPs

are used as a genetic marker which can be traced from one generation to another [9].

150

Furthermore, the study of how SNPs can lead to the onset of diseases is also of great interest [9].

Rapid, selective, and sensitive tools are needed for the identification and detection of SNPs.

Established molecular biology techniques to screen SNPs involve amplification and separation

steps [7]. A number of sensor and microarray detection strategies have been reported based on

colourimetric [125, 128], electrochemical [126, 130, 198], and optical [7, 199] detection. Some

of these involve large scale parallel analysis of samples for SNPs, and there is also interest in

dedicated analysis to detect specific SNPs.

A common feature of all biosensors, regardless of the transducer type, is the obligatory

immobilization of single-stranded oligonucleotides onto a solid substrate [18]. Ultimately, these

oligonucleotide probes are exposed to DNA targets, which may or may not hybridize to the

probes depending on sequence complementarity. The key parameters that govern stringency of

DNA hybridization and denaturation at a surface are temperature, ionic strength, pH, and density

of oligonucleotides. Temperature, pH, and ionic strength affect interactions that keep the DNA

duplex stable, such as hydrogen bonding, charge repulsion of the phosphate backbones, and base

stacking [28, 29]. The extent, rate, and selectivity of hybridization are heavily dependent on the

environment that is experienced [18]. For interfacial hybridization and denaturation, factors that

affect the environment include oligonucleotide distribution and density, the presence of nearest

neighbour interactions, and of interactions between probes and the surface [18]. Density,

however, is also a dynamic function of the degree of hybridization [18]. Ideally, to control

density, one must control the structural environment around each oligonucleotide probe. This

would then provide similar energetics for each probe molecule, resulting in improved selectivity

for the detection of SNPs.

151

Melt curves are commonly used to evaluate selectivity (stability) of binding by

considering differences in melting temperature (ΔTm) between a fully complementary (FC) and a

SNP target [7, 125, 200]. In bulk solution, the ΔTm can be 4–5°C for a 20-mer oligonucleotide

[16, 201]. This difference may be smaller for interfacial denaturation [7]. Moreover, melt

curves collected from surfaces tend to be broader, reflecting the diversity of energetics at an

interface and providing even lower selectivity for SNP detection [7]. A higher selectivity is

indicated by a greater difference in melting temperature as well as a sharper melting transition.

An increase in ΔTm helps to distinguish between FC and SNP target populations at a given

temperature and ionic strength [18]. An example of sharpening of melting transitions has

previously been demonstrated by Taton et al. by implementation of gold nanoparticles [125].

Control of orientation and nearest-neighbour interactions of oligonucleotide probes and

hybridized DNA is desirable to improve selectivity of hybridization at interfaces of

electrochemical sensors. The application of electric fields on conductive substrates has been

used to alter orientation of immobilized probes [202] to improve the discrimination of SNPs

[130], and to enhance hybridization kinetics [203]. However, electrostatic “combing” is not

possible with glass and silica-based substrates. A “matrix isolation” design was proposed by

Piunno et al. which involved the co-immobilization of non-nucleic acid oligomers with

oligonucleotide probes [18]. They have shown that a mixed film composed of oligonucleotide

probes and ethylene glycol phosphate can lower the Tm by 5°C and that SNP detection was still

possible [18]. Another example of inclusion of non-nucleic acid oligomers was demonstrated by

Boozer et al. [81]. The conformation of immobilized oligonucleotides was controlled by self-

assembling thiolated oligoethylene glycol and oligonucleotides on a gold surface, resulting in

improved hybridization efficiency [81]. This experiment showed that the orientation of

152

oligonucleotide probes could be controlled to maximize hybridization efficiency. Polymeric

coatings have frequently been used on surfaces to prevent adsorption of biomolecules. Yalçin et

al. have further explored this concept to improve DNA hybridization efficiency by 50%

(compared to silanized surfaces) by taking advantage of the swelling properties to create a 3-D

polymeric platform for the immobilization of oligonucleotides in microarray applications [204].

Urakawa et al. have also used a gel-coated microarray to optimize the discrimination of SNPs

[205]. The use of polymers is becoming more widespread to optimize sensor designs.

The polymeric coating used by Urakawa and co-workers is usually deposited as a layer

before oligonucleotides are immobilized. In contrast, the mixed oligomer and oligonucleotide

film used in our work is similar to that described in Piunno et al. [18], and comprises

interspersed oligomer brushes with oligonucleotides. This new method involves the building of

oligomer brushes from the surface via surface-initiated atom transfer radical polymerization

(ATRP) [178]. ATRP is based on exchanges of halogen atoms between dormant species and

metal catalysts to control the dynamics of radical polymerization [117-120]. It is a controlled/

“living” polymerization, providing controlled molecular weight and narrow polydispersity [117-

120]. Other advantages include mild conditions, availability of aqueous-based ATRP and a wide

choice of acrylate and methacrylate-based monomers [116]. Surface-initiated ATRP also allows

the growth of dense brushes without issues of steric hindrance [112]. In a previous investigation,

DNA hybridization and denaturation were found to be functional when mixed into films of

poly(2-hydroxyethyl methacrylate, PHEMA) grown onto silicon and glass substrates [178]. It

was demonstrated that the selectivity for the detection of 3 base-pair mismatch (3 bpm) can be

improved by physically controlling the environment surrounding the oligonucleotide probes

using PHEMA brushes [178]. Not only were the mixed films reusable for a number of cycles,

153

but the melt curves were 3-fold sharper, and the melting temperature differences (ΔTm) increased

by 30% in comparison to films composed only of oligonucleotides [178].

This new report provides a detailed examination of conditions required to control the

sharpness of the melting transitions and ΔTm values to distinguish between SNP and FC targets

for PHEMA-oligonucleotide coatings on glass surfaces. Fluorescence intensity measurements

were used to reflect the extent of hybridization. Fluorescence lifetime data provided a further

means to evaluate differences in the environments experienced by probe oligonucleotides in the

PHEMA-oligonucleotides mixed films, and films composed only of oligonucleotides.

3.4.3 Selectivity of Various Targets on the Mixed films

The mixed films were first investigated for selectivity towards FC (Sequence 5, Table

2.1) and 3 bpm targets (Sequence 7, Table 2.1) versus non-complementary (NC) targets

(Sequence 4, Table 2.1). Figure 3.23 shows that the relative integrated fluorescence intensities

for the FC:3bpm:NC targets were 30.5:4.6:1.0. These results suggested that hybridization

occurred, and that the signals were not dominated by non-specific adsorption. Figure 3.24

indicates that the FC targets adsorbed minimally to the 1× PBS-treated area in the absence of

probes, whereas the signal intensities at the probe-deposited areas were 1000-fold higher. The

various targets (Sequences 5–7 of Table 2.1) hybridized to the areas that contained probes, and

resulted in a relative fluorescence intensity ratio of 15.5:5.2:1.0 for FC:1bpm:3bpm.

154

Fig. 3.23: Integrated fluorescence intensities of various targets introduced to oligonucleotide probes (Sequence 1).

Standard deviations are shown. FC = fully complementary (Sequence 4), number of trials (n) = 8. 3 bpm = 3 base-

pair mismatch (Sequence 6), n = 8. NC = non-complementary (Sequence 3), n = 3

Figure 3.24: Integrated fluorescence intensities of various targets introduced to areas with and without immobilized

oligonucleotide probes. Standard deviations are shown. The “no probe” areas were spotted with 1× PBS. n (No

probe+ FC) = 4, n (probe + all other targets) = 8

155

3.4.4 Fluorescence Lifetime to Identify Different Microenvironments in Oligonucleotide Films vs. Mixed Films

The excited state decays of Cy3 dyes that were end-labeled to single-stranded

oligonucleotides (Sequences 2, Table 2.1) were measured for samples in which the probes were

immobilized on surfaces with and without PHEMA. Table 3.9 lists the lifetimes and their

relative amplitudes obtained from bi-exponential fitting applied to both types of surfaces. Figure

3.25 shows the raw data and the fitting curves for some fluorescence lifetime experiments. The

instrument response was very fast (~50 ps rise time) and had a negligible influence on fitting the

decay, so that no deconvolution routine was applied. In both cases, a monoexponential decay

was not sufficient to adequately fit the decay, as judged from the chi-square value and the profile

of the fitting residuals.

The fast component of Cy3 fluorescence decay in oligonucleotide films had a lifetime of

0.8 ns, while the slower component had a lifetime of 2.5 ns. The shorter lifetime had twice the

weight of the longer lifetime (66% vs. 34%). In the mixed film the fast decay time was 1.5 ns,

while the slow decay time was 3.8 ns. Again, the fast component was dominant, which was 82%

vs. 18% for the slower component. Overall, it is evident that the lifetime of Cy3 was

significantly longer in the mixed film than that measured from the oligonucleotide film. In

addition, the intensity time trajectories show that the fluorescence of the PHEMA film was bright

and stable, while the signal from the oligonucleotide film was less intense and photobleached

rapidly.

156

Table 3.9: Fluorescence lifetime and steady-state anisotropy values obtained from intensity time trajectories

measured on a confocal microscope.

τ1 (ns) α1 (%) τ2 (ns) α2 (%) χ2 r

Oligonucleotide filma 0.80±0.03 66 2.54±0.05 34 1.60 0.23±0.03

Mixed filmb 1.55±0.02 82 3.78±0.09 18 1.65 0.14±0.01

The lifetime fitting program uses the Levenberg-Marquardt algorithm and a χ2 value close to 1 represents a good fit. a –fluorescence acquired in the first 20s of every time series and b – fluorescence from the entire time series (300s).

Figure 3.25: Fluorescence intensity decay of Cy3-ssDNA in different environments: upper decay corresponds to

the mixed film and the lower one to the oligonucleotide film. Raw data in black, bi-exponential fit curves in red and

green. See table 2 for the numeric results of the fitting analysis.

157

Free Cy3 dyes in solution have a very short lifetime value of 180 ± 10 ps as reported by

Sanborn et al.[206]. This is because cyanine dyes are known to photoisomerize, i.e. undergo

trans-cis isomerisation through the polymethine chain, from the first excited singlet state to the

ground state (see Fig. 3.26). This process is in competition with fluorescence [206]. The

activation energy barrier for this process is strongly dependent on the rigidity of the local

environment which the dye experiences [206]. The study by Sanborn et al. compared the

photoisomerization efficiency, fluorescence quantum yields and lifetimes of Cy3 with Cy3B, the

latter having a rigid backbone which prevented isomerisation [206]. Cy3B had a much longer

lifetime of 2.70 ± 0.01 ns than Cy3. Therefore, the rigidity of the environment was a

determinant of these parameters [206]. Without a flexible linker, isomerisation was impeded.

Consequently, the fluorescence process dominated, and the fluorescence quantum yield and

lifetime were increased.

Fig. 3.26: The structure of 5′-Cy3-labeled oligonucleotides. Cy3 can be photoisomerized from the trans to cis

conformation through the methine bridge. R is the oligonucleotide sequence

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When Cy3 is covalently attached to the 5′-end of ssDNA, both quantum yield and

lifetime increase in magnitude [206]. This increase is associated with base-stacking of the dye,

which can occur with two to three terminal nucleobases [206, 207]. The decay of 3′-Cy3-ssDNA

in solution was previously measured and fitted to a sum of two exponentials, 1.2 ns (84%) and

2.1 ns (16%) on our instrument (Gradinaru, unpublished results). The two populations of probe

molecules with covalently attached Cy3 dyes are proposed to be systems with Cy3 stacked with

oligonucleotide (longer component), and systems without stacking which yielded a lower

activation barrier for photoisomerization to occur (shorter component) [206, 208]. Single-

molecule anisotropy data acquired in our lab supports this hypothesis (Gradinaru, unpublished

results). The longer lifetimes detected in the mixed films compared to oligonucleotide films

indicate a more rigid environment for the Cy3 dye in both the stacked and unstacked

conformations in a PHEMA matrix. Fig. 3.27 shows the two possible stacked and unstacked

interactions of Cy3 with oligonucleotides in an oligonucleotide film vs. a mixed film.

Fig. 3.27: Proposed stacked and unstacked interactions of Cy3 with immobilized oligonucleotide probes (a) in an

oligonucleotide film; (b) in a mixed film.

159

The presence of PHEMA provided a much more homogeneous spatial distribution of

Cy3-labeled oligonucleotides than observed from films composed of only oligonucleotides.

Figure 3.28 shows two fluorescence images of Cy3-oligonucleotides immobilized on glass, with

and without PHEMA. The oligonucleotide film showed a highly non-uniform surface

distribution, with variations on a micrometer scale, whereas the mixed film showed a

homogeneous intensity across tens of micrometers or more. In addition, when embedded in the

PHEMA matrix the Cy3 fluorescence was much more intense and photobleached much more

slowly than in the absence of the polymer. The combination of the fluorescence lifetime and

imaging are consistent with the hypothesis that the oligonucleotide probes were surrounded by

PHEMA brushes in the mixed film.

a) b)

Fig. 3.28: Fluorescence images of (a) oligonucleotide-only film, and (b) PHEMA film on glass cover slips. The

intensity variations in the PHEMA image are mostly due to imperfect flattening of the excitation field in a wide-

field microscope.

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3.4.5 Comparison of Selectivity for SNP Detection between Oligonucleotide and Mixed films

Interfacial melt curves were collected from oligonucleotide-modified glass surfaces with

and without co-immobilized PHEMA. The extent of hybridization was determined using the

integrated fluorescence intensity from Cy3-labeled FC and SNP targets (Sequences 5 and 7,

Table 2.1) collected at each temperature during each melt curve experiment. The modification

of oligonucleotides with fluorophores such as Cy3 at the 5′-end has been shown by Moreira and

co-workers to increase Tm by 1.4 °C [209]. However, Cy3 does not have an influence on the

width of the melting transition and slope [209]. The same dye was used on both types of

surfaces. The melt curves which were collected from solution used unlabeled oligonucleotides.

These latter sets of data were intended as benchmarks for ΔTm and dx values.

The melt curves were sigmoidal in nature with a single, cooperative transition similar to

those obtained by Piunno et al. under equilibrium conditions [18]. To compare the effectiveness

of the mixed film relative to the oligonucleotide film to discriminate SNP from FC targets, the dx

and Tm values were obtained by fitting each melt curve with the sigmoidal function:

dxxx

e

AAAy0

1

)( 212 −

+

−+= (3.4)

where A1 and A2 are the lower and upper baselines, x is temperature, y is the fraction of single-

stranded DNA (fssDNA), x0 is the melting temperature, and dx is the change in temperature for the

greatest change in fssDNA. Each melt curve experiment used slides that contained at least 8

hybridized spots for each of the two targets, hence yielding 16 melt curves per run. Large ΔTm

and a small dx values are desired for a highly selective surface. A small dx value signifies a

161

steep slope. Only melt curves which exhibited a good sigmoidal fit with R2 ≥ 0.95 were included

in the data analysis.

In general, interfacial hybridization and denaturation are very different energetically

compared to these processes done in bulk solution [160, 210]. The Tm values collected from

solid-liquid interfaces are often lower than those observed from bulk solution because the probes

are anchored to a solid substrate, hence lowering the stability of duplexes [129]. Melt curves

from bulk solution have been reported to have melting transition ranges of about 10 °C [129]

while interfacial melt curve were reported to have melting transition that spanned from 12–20 °C

[129, 199]. In this investigation, melting transitions of oligonucleotide films were as small as

10°C, whereas the mixed PHEMA-oligonucleotide films showed transition ranges as small as

5°C. This is greatly improved from previous studies [129, 199]. A more accurate variable to

describe the steepness of the melting transition is to use the dx value. Table 3.10 lists the dx

values obtained from both oligonucleotide and mixed film surfaces in all ionic strengths. For

SNP targets in 0.5× PBS, the values ranged from 1.1–3.0 °C for the mixed films compared to

3.6–5.7°C for the oligonucleotide films, indicating that overall, the mixed films provided sharper

melting transitions. The average of dx values found for the mixed films for both targets were

statistically the same as those obtained from bulk solution (1.9°C).

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Table 3.10: Comparison of dx values in all ionic strengths studied.

0.1× PBS 0.5× PBS 1.0× PBS

Type of surface Trial dx value of FC

targets (°C)

dx value of SNP targets

(°C)

dx value of FC targets

(°C)

dx value of SNP targets

(°C)

dx value of FC targets

(°C)

dx value of SNP

targets (°C)

1 3.4 ± 1.5 1.4 ± 0.7 2.0 ± 0.5 3.6 ± 1.3 5.7 ± 0.9 4.4 ± 0.6 Oligonucleotide film only

2 2.2 ± 0.5 1.8 ± 0.4 2.4 ± 0.8 3.8 ± 1.0 4.0 ± 1.4 5.5 ± 0.9

3 - - 7.5 ± 1.1 5.7 ± 0.8 4.6 ± 0.2 3.1 ± 0.8

4 - - - - 3.8 ± 1.6 2.3 ± 1.4

Avg 2.8 ± 0.6 1.6 ± 0.2 4.0 ± 1.8 4.4 ± 0.7 4.5 ± 0.4 3.8 ± 0.7

Mixed film 1 3.2 ± 0.5 0.8 ± 0.4a 1.7 ± 0.4 1.1 ± 0.8 3.8 ± 0.5 2.0 ± 0.8

2 2.1 ± 1.1 0.9 ± 0.6 2.8 ± 0.7 3.0 ± 0.4 3.8 ± 1.1 2.2 ± 0.6

3 2.1 ± 0.4 3.0 ± 0.2 3.4 ± 1.2 2.7 ± 0.9

4 4.2 ± 1.6 2.9 ± 0.4 - -

Avg 2.6 ± 0.6 0.8 ± 0.1 2.7 ± 0.6 2.5 ± 0.5 3.7 ± 0.1 2.3 ± 0.2

Bulk solution 1 2.5 ± 0.1 2.1 ± 0.1 2.6 ± 0.1 1.9 ± 0.1 2.9 ± 0.1 2.5 ± 0.1

± The standard deviations of at least 3 replicates were reported. The averages of all experiments were reported with their standard errors.

a Two replicates.

Avg: average values

The total ionic strength for each salt condition was: 0.11 M (0.1× PBS) , 0.56 M (0.5× PBS), and 1.12 M (1.0 × PBS).

163

Table 3.11: Comparison of ΔTm values in all ionic strengths studied.

0.1× PBS 0.5× PBS 1.0× PBS

Surf

ace

Tri

al Tm of

FC targets

(ºC)

Tm of SNP

targets (ºC)

Δ Tm (ºC) at 95% C.I.a

Tm of FC

targets (ºC)

Tm of SNP

targets (ºC)


Tm of FC

targets (ºC)

Tm of SNP

targets (ºC)


1 38.1 ± 1.7

34.6 ± 1.6

3.4 ± 2.5

47.7 ± 0.7

43.0 ± 2.0

4.7 ± 2.6 49.6 ± 0.5

45.1 ± 2.1

4.5 ± 5.6 A

2 35.1 ± 1.5

34.2 ± 0.9

0.9 ± 2.0

42.4 ± 0.8

39.5 ± 3.7

2.9 ± 9.5 47.8 ± 1.7

44.8 ± 3.5

3.0 ± 5.3

3 - - - 44.1 ± 2.9

40.9 ± 1.8

3.2 ± 2.9 47.4 ± 2.9

44.6 ± 1.8

2.8 ± 4.4

4 - - - - - - 45.4 ± 1.7

43.7 ± 1.6

1.7 ± 2.3

Avg 36.6 ± 1.5

34.4 ± 0.2

2.2 ± 1.3

44.7 ± 1.6

41.1 ± 1.0

3.6 ± 0.6 47.6 ± 0.9

44.6 ± 0.3

3.0 ± 0.6

B 1 35.1 ± 1.3

30.8 ± 1.4 b

4.3 ± 4.7

49.1 ± 2.0

43.6 ± 1.8

5.5 ± 2.8 46.2 ± 0.4

42.6 ± 1.5

3.6 ± 1.4

2 36.4 ± 0.7

30.8 ± 0.9

5.6 ± 1.1

46.9 ± 1.7

36.9 ± 3.4

10.0 ± 2.9

47.3 ± 1.6

43.3 ± 1.8

4.0 ± 2.6

3 - - - 52.9 ± 1.7

37.2 ± 1.6

15.7 ± 1.7

50.4 ± 3.3

44.9 ± 2.1

5.5 ± 3.3

4 - - - 46.5 ± 2.8

36.9 ± 1.9

9.6 ± 3.2 - - -

Avg 35.7 ± 0.6

30.8 ± 0.0

5.0 ± 0.7

48.8 ± 1.5

38.6 ± 1.7

10.2 ± 2.1

48.0 ± 1.3

43.6 ± 0.7

4.4 ± 0.6

Bulk solution

1 52.9 ± 0.1

46.4 ± 0.1

6.5 ± 0.1

62.5 ± 0.1

53.3 ± 0.1

9.2 ± 0.1 65.9 ± 0.1

57.5 ± 0.1

8.4 ± 0.1

Surface A: Oligonucleotide films; surface B: Mixed films.

Avg: average values. ± The standard deviations of at least 3 replicates were reported. The averages of all experiments were reported with their standard errors.

a The confidence interval (C.I.) at 95% was constructed for the difference in Tm values between FC and SNP targets.

b Two replicates.

The total ionic strength for each salt condition was: 0.11 M (0.1× PBS), 0.56 M (0.5× PBS), and 1.12 M (1.0× PBS).

164

The ΔTm values collected from oligonucleotide films and mixed films in all ionic

strengths are listed in Table 3.11. In 0.5× PBS, the ΔTm values observed for each trial from the

oligonucleotide surface are much lower than the differences observed for the mixed film (at a

confidence level of 95%). The largest ΔTm found for oligonucleotide films was 4.7 ± 2.6 ºC. At

the same ionic strength, the largest ΔTm found for mixed films was 15.7 ± 1.7 ºC, and exceeded

the performance observed for the bulk solution experiment by 6.5 ºC. The average ΔTm for

mixed films was more than three times as large as the average ΔTm found for the oligonucleotide

films. The 2- to 3-fold increase in ΔTm seen when using the mixed films is due to a decrease in

the Tm for the SNP targets and an increase in the Tm of FC targets; i.e. the mixed film structure

stabilizes FC hybrids and destabilizes SNPs, relative to films composed of only oligonucleotides.

In general, the melting temperatures decreased as the salt concentration decreased and the

SNP targets had lower Tm than the FC targets, as was expected. The coefficient of variation

(CV) (%) for the Tm values in each experiment was under 10%. The CV between experiments

for the same target was under 7%. The CVs of the dx values for each experiment were larger due

to the magnitudes of the standard deviations with respect to the data values. The dx values

between experiments were generally reproducible within 1–2 °C.

The increase in the sharpness of melt curves was also evident in 1× and 0.1× PBS. The

former ionic strength yielded dx values for the SNP targets ranging from 2.0–2.7 °C for the

mixed films versus 2.3–5.5 °C for the oligonucleotide films. Using FC targets, the mixed films

showed a more consistent pattern of sharper melting transitions than the transitions seen for

oligonucleotide films. In 0.1× PBS, the dx values for the SNP targets collected from the mixed

films were twice as sharp as those from oligonucleotide films and bulk solution. The dx values

165

for the FC targets between the two surfaces were not significantly different. The average ΔTm

value associated with the mixed films in 0.1× PBS was more than twice as large as that from the

oligonucleotide films. For the 1× PBS case, the average ΔTm value was generally similar

between the mixed films and the oligonucleotide films at the 95% confidence level. Figure 3.29

is a collection of representative melt curves that reflect the trends shown in Tables 3.10–3.11.

The mixed films provided better resolution between the FC and SNP targets compared to the

oligonucleotide films 0.1× and 0.5× PBS (Figures 3.29a–d). The mixed films either had sharper

melting transitions and/or larger ΔTm values to achieve SNP discrimination, whereas the

oligonucleotide films had significant overlapping populations of FC and SNP targets at any

given temperature and ionic strength. The melt curves of Figs. 3.29b and 3.29d indicate the

advantage for SNP analysis by using the PHEMA-oligonucleotide mixed film in combination

with moderate ionic strength.

OLIGONUCLEOTIDE FILMS MIXED FILMS

(a) (b)

166

(c) (d)

(e) (f)

Figure 3.29: Individual melt curves of (•) SNP targets and (■) FC targets that represent average Tm and dx values

collected from oligonucleotide (left column) and mixed films (right column) in increasing PBS strengths (0.1, 0.5,

1×).

(a) DNA film in 0.1 × PBS: Tm (SNP) = 34.6 ± 0.3, Tm (FC) = 38.3 ± 0.7, dx (SNP) = 1.3 ± 0.3, dx (FC) = 3.0 ± 0.6;

(b) mixed film in 0.1 × PBS: Tm (SNP) = 30.7 ± 0.2, Tm (FC) = 36.7 ± 0.5, dx (SNP) = 0.6 ± 0.1, dx (FC) = 2.0 ±

0.5; c) DNA film in 0.5 × PBS: Tm (SNP) = 41.5 ± 1.0, Tm (FC) = 41.5 ± 0.6, dx (SNP) = 3.3 ± 1.0, dx (FC) = 2.1 ±

0.5; (d) mixed film in 0.5 × PBS: Tm (SNP) = 36.7 ± 0.4, Tm (FC) = 47.3 ± 0.5, dx (SNP) = 2.9 ± 0.4, dx (FC) = 3.0

± 0.5; e) DNA film in 1 × PBS: Tm (SNP) = 44.4 ± 0.9, Tm (FC) = 47.7 ± 1.1, dx (SNP) = 3.6 ± 0.9, dx (FC) = 4.6 ±

1.1; f) mixed film in 1 × PBS: Tm (SNP) = 43.9 ± 0.3, Tm (FC) = 48.5 ± 0.8, dx (SNP) = 1.7 ± 0.3, dx (FC) = 3.6 ±

0.7.

The increase in sharpness of melt curves associated with the mixed films appears most

prominently for mismatched duplexes, and this is the case for SNPs as well as previously

reported 3 bpm targets [178]. The sharpness observed for mismatched systems may be due to

167

the sterics associated with “breathing” events at terminal ends and mismatched sites. Fig. 3.30

illustrates this concept. The “breathing” mechanism is well known in which segments in these

regions repeatedly denature and re-anneal [27]. The PHEMA brushes tend to magnify the

sharpness of the melting transition more so for the 3 bpm targets [178] compared to the SNP

targets, as would be expected if the interactions were dominated by physical occupancy

considerations. The hanging pendant groups of 2-hydroxyethyl may achieve higher chain

mobility as temperature increases. The mismatched sites could then be destabilized by these side

chains.

Furthermore, the data shows that the destabilization effect is related to salt concentration,

suggesting an electrostatic contribution is also significant. The local salt concentration may be

amplified due to molecular crowding by PHEMA brushes (see Fig. 3.30). At 1× PBS, there was

a large amount of salt to counteract repulsion arising from negative charges of the DNA

backbone and destabilizing forces from PHEMA brushes; all Tm values for both FC and SNP

were relatively high. At 0.1× PBS, there may have been insufficient salt to suppress the

repulsion, hence both FC and SNP targets denatured at a lower temperature. At 0.5× PBS, the

effect of uncharged PHEMA played a more dominant role in the destabilization of the SNP

targets, but did not compromise the stability of the FC targets. The trend in destabilization of the

duplexes can also be explained by the swelling behaviour trend of PHEMA. It was found by

D’Agostino et al. that the swelling ratio decreases with increasing NaCl concentration [211].

The physical occupancy of PHEMA and hence steric repulsion would play more of a dominant

role at lower ionic strengths.

168

Fig. 3.30: Interpretation of melting transition width and Tm differences between different targets hybridized to the

mixed film.

A further consideration about the origins of the changes of duplex stability is related to

water activity in the presence of PHEMA. Spink and Chaires found that increasing the

concentration of polyethylene glycol (PEG) 3400 increased the melting temperature of duplexes

in bulk solution [212]. Here we observed some stabilization for FC targets for immobilized

PHEMA-oligonucleotide films. However, the effects of immobilized PHEMA tended to

destabilize base mismatches. Spink and Chaires reasoned that the presence of PEG caused a

change in the osmotic environment, which can affect the stability of DNA and its helical

structure by changing the number of water molecules that can bind to DNA [212]. It is known

that the amount of water that is uniquely bound to DNA can influence DNA stability because

bound water is thermodynamically different from bulk water [212]. Water molecules first adsorb

to the sugar-phosphate regions but it is also known that some are associated with base-pairs (4–5

water molecules per base-pair) [212]. Such bound water can be released during denaturation,

causing changes in molar volume and entropy. The addition of co-solutes such as PHEMA with

its pendant sidegroups can affect water activity, particularly when duplex disruption due to a

169

mismatch is present. The co-immobilization of PHEMA brushes should be able to contribute

both stabilizing and destabilizing effects on double stranded DNA, depending on the presence of

mismatches.

The broad nature of melt curves collected from oligonucleotide films can also be

attributed to adsorbed oligonucleotides and/or other intermediate structures [160]. The

minimization of adsorption of oligonucleotides on the mixed films contributed to the sharpness

of the melting transition [178]. The signal-to-background ratio was about 2:1 for oligonucleotide

films but was 100:1 for the mixed films. The mixed film apparently blocked adsorption-prone

sites by interspersing of PHEMA chains. One additional point worth noting is that the high

signal-to-background ratio obtained from mixed films was not the result of stringent washing

conditions such as using formamide and SDS washes, and sonication, as is typically done in

hybridization experiments [205].

3.4.6 Conclusions

A comparison has been made of the selectivity and physical environment of

oligonucleotide films with mixed films of oligonucleotides and PHEMA brushes. The resolution

to distinguish between SNP and FC targets by means of temperature was clearly enhanced for

the mixed films. A greater separation of Tm values by as much as 3-fold was observed, and a

sharpening of melting transitions reached as much as 2-fold. The fluorescence of Cy3 attached

to single-stranded oligonucleotide probes showed a bi-exponential decay on the nanosecond

timescale. Both fast and slow lifetime values increased substantially in the mixed films

compared to oligonucleotide films. Cy3 is known to be a molecular rotor and our results indicate

that in the presence of PHEMA the dye experiences a much more rigid environment. The

170

oligonucleotide probes are most likely surrounded by PHEMA brushes at a molecular level,

which is instrumental in improving the selectivity for SNP detection.

171

Chapter 4

4 Summary This investigation demonstrated that a mixed film composed of oligonucleotides and

PHEMA oligomers can be immobilized on silica-based substrates. The use of a mixed BZ-

APTMS and APTMS led to a smoother surface, as shown by AFM data, compared to a surface

derivatized with APTMS alone. The surface-initiated growth of HEMA monomers via ATRP on

silica-based substrates was demonstrated. The initial mixed silane layer provided similar ATRP

reaction rates as on the APTMS film, as shown by ellipsometry data, but the rates were more

reproducible on the mixed aminosilane surface. The smoother mixed aminosilane surface

became an excellent base film layer for further immobilization of both oligonucleotides and

PHEMA to yield reproducible and functional biosensing surfaces.

Hybridization events on the mixed film were functional as proven by using Cy3-labeled

targets of various degrees of complementarity. Comparison of adsorption levels and melt curve

data was made between mixed films and oligonucleotide films. There was an increase in signal-

to-background ratio of 4-fold for mixed films compared to the oligonucleotide films. The mixed

film was also reusable for at least 5 cycles of use. Melt curve data using 3 bpm targets

demonstrated a 2-fold increase in ΔTm and 3-fold increase in sharpness of melting transitions

when PHEMA was co-immobilized with oligonucleotide probes.

A comparison of the ability of XPS and ToF-SIMS equipped with a bismuth cluster ion

source to detect both oligonucleotide and PHEMA components on a glass substrate was done.

XPS could not detect atomic signatures of oligonucleotides but ToF-SIMS could. The bismuth

cluster ion source could also enhance the signal even though the oligonucleotide probes were

172

surrounded by an excess of PHEMA brushes. The oligonucleotide density was estimated by

fluorescence data and a gold nanoparticle-labeling strategy to be on the order of 7×1010 to

3.7×1011 oligonucleotide probes cm-2. ToF-SIMS was determined to be the more sensitive

surface analysis technique to characterize the mixed oligonucleotide and PHEMA films.

The enhancement of detection of SNP targets by the mixed film compared to the

oligonucleotide film was shown. The signal-to-noise ratio was found to be 100-fold higher for

the mixed film. Numerous melt curves were collected from each surface at three different ionic

strengths to determine the effects of salt on the ability of PHEMA brushes to change ΔTm and

steepness of the melting transition. Moderate ionic strength was found to allow PHEMA brushes

to increase ΔTm by 3-fold and steepness of melt curve slope by 2-fold. Increases in ΔTm are due

to an increase in stability for fully complementary targets and a decrease in stability for partially

matched targets. The sharpness was improved due to the isolation effect of PHEMA, leading to

a reduction of intermediate states that often broaden the melting transition ranges. This effect

was amplified when there are base-pair mismatches because there is a lower number of degrees

of freedom for melting, hence increasing the cooperativity for melting. At moderate ionic

strength, the increase in stability for FC targets may have been due to effective charge screening,

but this screening was not adequate when there were base-pair mismatches due to the

destabilizing effect of the steric repulsion of PHEMA and the increased bulkiness of mismatched

duplexes. The lifetimes of Cy3 emission from Cy3-labeled oligonucleotide probes on surfaces

with PHEMA were about twice as long as those immobilized on surfaces without PHEMA. The

increase in lifetime values indicated an increase in rigidity of the mixed film environment for the

oligonucleotide probes compared to the oligonucleotide film. This was consistent with the

intention that the oligonucleotide probes should be surrounded by PHEMA brushes, resulting in

173

a narrower distribution of energetics of DNA hybrids observed in the melt curves collected from

mixed films.

These collective results verified that the mixed aminosilane base layer and the ATRP-

based growth of the oligomer portion were successful in producing the mixed film. This mixed

film strategy has proven to be practical in enhancing signal-to-noise ratio by reducing adsorption

and reducing the degrees of freedom favourable to hybridization. Furthermore, the increased

ability of the mixed film at low to moderate ionic strength to discriminate SNP targets from fully

complementary targets by narrowing the melting transition width and increasing the melting

temperature differences makes it an excellent sensing layer candidate for numerous biosensor

and microarray applications.

174

Chapter 5

5 Future Directions Thus far, this work has only focused on the performance of mixed films which were

composed of one neutral monomer, and which was polymerized from a 1:1 mixed protected and

unprotected (3-aminopropyl)trimethoxysilane (APTMS). Further changes in the experimental

parameters can be examined to understand how to increase the steepness of the melting transition

slope and difference in melting temperatures. For example, a negatively charged monomer such

as 3-sulfopropyl methacrylate can be used to examine the effect of a constant net negative charge

around oligonucleotide probes. The ratio of protected and unprotected APTMS can be varied,

which should theoretically change the oligonucleotide-to-oligomer ratio. Alternatively, the

loading concentration of oligonucleotides can be varied to change the oligonucleotide-to-

oligomer ratio. Furthermore, different oligomer and oligonucleotide lengths can be used. One

important experiment that should be done is to test the mixed films on other sequences to

determine if it can also decrease the melting transition range and increase the melting

temperature differences. The location of the SNP site can be varied, and the most pronounced

effect would probably be observed for mismatched sites which are centrally located or closer to

the surface, but not at the terminus. Not only does ionic strength matters but the choice of buffer

ion types for use in the denaturation process might affect how PHEMA brushes could change the

melting transition width and melting temperature differences. For example, ions of different

sizes and charges of various degrees of polarizability should be studied since different ion

properties may affect the stability of DNA to a different extent [213]. Lastly, real samples

should be tested to see the mixed film surface can detect complementary sequences from a

175

complex matrix. These changes may affect the overall performance of the mixed film to

discriminate fully complementary targets from targets containing a one base-pair mismatch.

The lifetime studies of Cy3 fluorescence were promising in that data from such

experiments can be used to probe the dynamics of the mixed film environment. A more detailed

examination may encompass fluorescence anisotropy, i.e. the rotational freedom of fluorophores

by following the intensities of the perpendicularly and parallel polarized fluorescence emission.

However, a different fluorophore may have to be chosen because Cy3 labels on oligonucleotides

are too sensitive to temperature and local rigidity, and have been shown to provide high steady-

state anisotropy values [206]. It would be more prudent to collect time-resolved anisotropy data

instead and/or choose another dye [206]. Fluorophores that are categorized as “molecular rotors”

such as meso-substituted boron-dipyrrin (BODIPY) dyes are very sensitive to the local viscosity

[214]. Alexa and fluorescein dyes have also been used in anisotropy measurements [215]. If

Cy3 is still chosen as the physical probe, according to Sanborn and co-workers, single-stranded

DNA are better probes for studying lifetime and anisotropy data because isomerization of Cy3 is

suppressed [206]. It has been shown that single-stranded DNA-Cy3 stacking interactions are

reduced when they undergo hybridization, hence resulting in a lower barrier for Cy3

isomerization [206]. Besides using single-stranded probes, one can change the flexibility of the

probe by changing the sequence. For example, an oligonucleotide composed of just thymine is

known to be very flexible as opposed to one composed of just adenine, which is very rigid due to

the opportunity for more base-stacking interactions between the purine bases [216]. The use of a

single-stranded, flexible oligonucleotide sequences may give more information about the

dynamics of the mixed biosensing environment.

176

The vision of an ideal DNA biosensing surface is to achieve minimal adsorption, high

reusability, a high selectivity, and a high sensitivity as well as being reagentless and labeless.

The mixed film as a novel DNA biosensing surface was shown to be reusable and selective with

minimal adsorption levels, but it must be further modified to become an “ideal biosensor”. An

idea for the next step to improve sensitivity would be to immobilize the mixed oligonucleotide-

polymer film on a 3-D polymeric coating as demonstrated in the work by Yalçin and co-workers

[204]. This coating was shown to increase binding sites and lower steric hindrance, which led to

a higher hybridization efficiency compared to silanized surfaces [204]. The swellable property

of this polymer provides more rotational freedom and less steric hindrance for the immobilized

probes because they are lifted away from the surface [204]. The co-immobilization of oligomers

would still serve to reduce nearest neighbour interactions, and to reduce interactions between

probes and the surface.

The intention to create the mixed films on silica-based substrates is to apply this

biosensing surface to optical fiber surfaces such that TIR can be used to excite surface-localized

fluorophores. The first detection scheme may involve just the detection for Cy3-labeled targets,

for example. The dye can be attached to the target strands once they have been isolated and

amplified from a real complex sample. The use of fluorescence to transduce DNA

hybridizations means that the use of labeled reagents cannot be avoided, but labels can also be

incorporated in such a way that the sample itself need not be labeled. For example, Wang and

Krull have demonstrated a self-contained optical fiber biosensor by attaching an intercalator,

thiazole orange, to the immobilized probes. When DNA hybridization occurs with unlabeled

target sequences, the dye emission was enhanced due to intercalation [46]. The sensor was

regenerable as well [46]. Alternatively, the dye can be covalently attached to PHEMA (either to

the monomer or polymer) so that it can be permanently attached to the mixed films. Lastly, one

177

can use a sandwich assay in which reporter strands containing a fluorophore are used; they can

bind to immobilized probes on optical fibers after the unlabeled target sequences have bound to

immobilized probes, as described in Algar and Krull [47]. The strategy for immobilizing the

mixed films would have to be altered. The initiator for the surface-initiated polymerization

would have to be attached to biotin such that it can be immobilized onto a neutravidin-coated

surface. Another way would be to mix thiolated oligonucleotides with thiolated-initiator such

that there are more initiators immobilized onto the quantum dot-coated surface, without using

neutravidin. The fluorophores attached to these reporter strands should experience a “matrix

isolation” environment in the mixed films to yield melt curves with narrow distribution of

energetics and increased melting temperature differences between fully and SNP targets.

178

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189

Appendices A.1. Kinetic study of the growth of PHEMA via ATRP on surfaces modified with and without

initiator.

y = 20.198x - 1.4029R2 = 0.9697

y = 0.0602x + 2.4993R2 = 0.0581

-20

0

20

40

60

80

100

120

140

160

0 1 2 3 4 5 6 7

time (h)

appa

rent

thic

knes

s (n

m)

(♦) surfaces with initiator; (■) surfaces without initiator.

190

A.2. Comparison of ATRP rates on mixed BZ-APTMS and APTMS and free APTMS surfaces.

y = 177.59x - 9.8536R2 = 0.986 (BZ-APTMS 1)

y = 172.23x - 17.401R2 = 0.9441 (BZ-APTMS 2)

y = 202.01x - 14.575R2 = 0.9697 (free APTMS 1)

y = 128.08x + 12.973R2 = 0.9824 (free APTMS 2)

-200

0

200

400

600

800

1000

1200

1400

0 1 2 3 4 5 6 7

time (h)

ellip

som

etry

thic

knes

s (A

ngst

rom

s)

BZ-APTMS 2BZ-APTMS 1free APTMS 1free APTMS 2Linear (BZ-APTMS 1)Linear (BZ-APTMS 2)Linear (free APTMS 1)Linear (free APTMS 2)

191

A.3. Overlaid ToF-SIMS (a) positive and (b) negative spectra obtained by using Bi+ on various

surfaces (top to bottom: mixed APTMS and BZ-APTMS, oligonucleotides only, mixed

oligonucleotides and PHEMA, PHEMA only).

(a)

192

(b)

193

A.4. Overlaid ToF-SIMS (a) positive and (b) negative spectra obtained by using Bi3+ on various



(a)

194

(b)

195

A.5. Overlaid ToF-SIMS (a) positive and (b) negative spectra obtained by using Bi5+ on various



(a)

196

(b)

197

A.6. Overlaid ToF-SIMS positive spectra obtained by using Bi3+ on surfaces containing various

ratios of oligonucleotides to PHEMA (top to bottom: control, conc1, conc2, and conc3. Conc1

was 0.10 μM, conc2 was 0.48 μM, and conc3 was 1.02 μM (loading oligonucleotide probe

(sequence 2) concentrations).

201

A.7. Overlaid ToF-SIMS positive spectra obtained by using Bi5+ on surfaces containing various

ratios of oligonucleotides to PHEMA (top to bottom: control, conc1, conc2, and conc3. Conc1

was 0.10 μM, conc2 was 0.48 μM, and conc3 was 1.02 μM (loading oligonucleotide probe

(sequence 2) concentrations).

A MIXED BIOSENSING FILM COMPOSED OF OLIGONUCLEOTIDES … · students: Anthony Tavares, Rhys Crasto,...

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