FABRICATION OF POLYSTYRENE CORE-SILICA...
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Fabrication of Polystyrene Core-Silica Shell Nanoparticles for Scintillation Proximity Assay (SPA) Biosensors Item Type text; Electronic Thesis Authors Noviana, Eka Publisher The University of Arizona. Rights Copyright © is held by the author. Digital access to this material is made possible by the University Libraries, University of Arizona. Further transmission, reproduction or presentation (such as public display or performance) of protected items is prohibited except with permission of the author. Download date 11/05/2018 05:34:49 Link to Item http://hdl.handle.net/10150/556865
Transcript of FABRICATION OF POLYSTYRENE CORE-SILICA...
Fabrication of Polystyrene Core-Silica Shell Nanoparticlesfor Scintillation Proximity Assay (SPA) Biosensors
Item Type text; Electronic Thesis
Authors Noviana, Eka
Publisher The University of Arizona.
Rights Copyright © is held by the author. Digital access to this materialis made possible by the University Libraries, University of Arizona.Further transmission, reproduction or presentation (such aspublic display or performance) of protected items is prohibitedexcept with permission of the author.
Download date 11/05/2018 05:34:49
Link to Item http://hdl.handle.net/10150/556865
FABRICATION OF POLYSTYRENE CORE-SILICA SHELL NANOPARTICLES FOR
SCINTILLATION PROXIMITY ASSAY (SPA) BIOSENSORS
by
Eka Noviana
____________________________
A Thesis Submitted to the Faculty of the
DEPARTMENT OF CHEMISTRY AND BIOCHEMISTRY
In Partial Fulfillment of the Requirements
For the Degree of
MASTER OF SCIENCE WITH A MAJOR IN CHEMISTRY
In the Graduate College
THE UNIVERSITY OF ARIZONA
2015
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STATEMENT BY AUTHOR
This thesis has been submitted in partial fulfillment of requirements for an advanced degree at the University of Arizona and is deposited in the University Library to be made available to borrowers under rules of the Library.
Brief quotations from this thesis are allowable without special permission, provided
that an accurate acknowledgement of the source is made. Requests for permission for extended quotation from or reproduction of this manuscript in whole or in part may be granted by the head of the major department or the Dean of the Graduate College when in his or her judgment the proposed use of the material is in the interests of scholarship. In all other instances, however, permission must be obtained from the author.
SIGNED: Eka Noviana
APPROVAL BY THESIS DIRECTOR
This thesis has been approved on the date shown below:
May 13, 2015 Craig A. Aspinwall, Ph.D. Date
Professor of Chemistry and Biochemistry
3
ACKNOWLEDGEMENTS
I am indebted to my advisor, Dr. Craig Aspinwall, who kindly let me join his group
and has provided never ending encouragement and support during my research in the
Aspinwall Lab. I could never thank you enough for your exceptional patience and
guidance. I must also thank all the members of the Aspinwall groupnwho have been great
colleagues, friends and family to me. I thank Isen who has been an outstanding lab mentor
and helped me a lot with the research. Thank you to Vanessa and Xuemin who have been
sharing the office room with me for the past 1.5 years. Thank you to Kofi, the happiest and
funniest person ever; Jinyang, the most sincere and hard-working person I have ever met;
and Mark, one of the nicest and most thoughtful people I have encountered. I also thank
Colleen, our optimistic SPA specialist, for all her insights, support and encouragement, and
to all 2013-batch lab fellows (Malithi, Kendall, Nipuna and Zeinab) for all the wonderful
times.
I also thank my thesis committee: Dr. Michael Heien and Dr. Jeffrey Pyun for their
invaluable feedback and suggestions on my thesis. I thank Dr. Jeanne Pemberton as well,
who initially served as one of thesis committee members and one of faculty recommenders
for my Ph.D. applications.
Finally, thank you Mom and Dad for your prayers and support. I would not have
been able to do this without you. Thank you to all my friends in Tucson who have been
cheering me up and making me never feel lonely when I was thousands miles apart from
my family, and all my friends in other parts of the world for all the magnificent inspiration
and support.
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DEDICATION
To my beloved parents, mentors, friends, and fellow members, without whom it was impossible to finish my thesis work
Indeed, Allah will not change the condition of a people until they change what is in themselves
-QS Ar-Ra’d 13:11
Anyone who stops learning is old, whether at twenty or eighty. -Henry Ford
The essence of intercultural education is the acquisition of empathy-the ability to see the world as others see it, and to allow for the possibility
that others may see something we have failed to see -J. William Fulbright
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TABLE OF CONTENTS LIST OF ABBREVIATIONS ....................................................................................... 7
LIST OF FIGURES ....................................................................................................... 8
LIST OF SCHEMES ..................................................................................................... 11
LIST OF TABLES ........................................................................................................ 12
ABSTRACT .................................................................................................................. 13
CHAPTER 1: INTRODUCTION ................................................................................. 14
1.1. Background ................................................................................................. 14
1.2. Literature Review ........................................................................................ 16
1.2.1. Introduction to Biosensors ............................................................... 16
1.2.2. Roles of Nanomaterials in Biosensing Applications ....................... 18
1.2.3. Nanoparticles for Optical Imaging in Biological System ............... 20
1.2.3.1. Polystyrene Nanoparticles (PS NPs) ................................ 23
1.2.3.2. Silica Coating of Nanoparticles ........................................ 25
1.2.4. Scintillation Proximity Assay (SPA) Biosensors ............................ 30
1.2.4.1. Radioisotopes in SPA ....................................................... 31
1.2.4.2. SPA Materials ................................................................... 35
1.2.4.3. Detection and Measurement in SPA ................................ 37
1.2.4.4. Applications of SPA Biosensors in Biology and
Medicine ........................................................................... 39
CHAPTER 2: MATERIALS AND METHODS ........................................................... 44
2.1. Materials ...................................................................................................... 44
2.2. Fabrication of Scintillant-doped PS NPs ..................................................... 44
2.3. Silica Coating of Scintillant-doped PS NPs ................................................ 45
2.4. Characterization of PS NPs ......................................................................... 46
2.4.1. Size and Morphology Determination by TEM ................................ 46
2.4.2. Size and Size Distribution Determination by DLS ......................... 46
2.4.3. Zeta Potential Measurements .......................................................... 46
2.4.4. Emission Spectra Acquisition ......................................................... 47
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2.4.5. Scintillation Response Measurements ............................................. 47
2.5. SPA Proof-of-Concept using DNA Hybridization ...................................... 47
2.5.1. Immobilization of ssDNA on PS Core-Silica Shell NPs ................. 48
2.5.2. Radiolabeling of Complementary ssDNA ....................................... 48
2.5.3. DNA Hybridization on PS Core-Silica Shell NPs ........................... 49
CHAPTER 3: RESULTS AND DISCUSSIONS .......................................................... 51
3.1. Fabrication and Characterization of Scintillant-doped Core-Shell NPs ...... 51
3.1.1. Morphology, Size and Size Distribution ......................................... 54
3.1.2. Surface Charge Characterization ..................................................... 57
3.1.3. Emission Spectra of Scintillating NPs ............................................. 60
3.1.4. Scintillation Response to Free 3H .................................................... 61
3.2. Scintillation Proximity Assay using DNA Hybridization ........................... 62
3.2.1. Immobilization of Oligonucleotides on PS Core-Silica Shell NPs . 63
3.2.2. Oligonucleotide Radiolabeling ........................................................ 68
3.2.3. DNA Hybridization on PS Core-Silica Shell NPs ........................... 73
CHAPTER 4: SUMMARY AND FUTURE DIRECTIONS ........................................ 77
4.1. Summary ..................................................................................................... 77
4.2. Future Directions ......................................................................................... 77
REFERENCES .............................................................................................................. 80
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LIST OF ABBREVIATIONS
AAPH : 2,2’-azobis(2-methylpropionamidine) dihydrochloride
APTS : (3-aminopropyl)triethoxysilane
CCD : charge coupled device
CPM : counts per minute
Dimethyl POPOP : 4-bis(4-methyl-5-phenyl-2oxyzolyl)benzene
DLS : dynamic light scattering
DMF : dimethyl formamide
DMSO : dimethyl sulfoxide
DNA : deoxyribonucleic acid; ssDNA : single-stranded DNA
EDC : N-(3-dimethylaminopropyl)-N′-ethylcarbodiimide
FITC : fluorescein isothiocyanate
MES : 2-(N-morpholino)ethanesulfonic acid
MPTS : (3-mercaptopropyl)trimethoxysilane
NHS : N-hydroxysuccinimide
NPs : nanoparticles
OEG : oligo(ethylene glycol)
PdI : polydispersity index
PMT : photomultiplier tube
PS : polystyrene
pTP : para-terphenyl
QD : quantum dots
SPA : scintillation proximity assay
TEM : transmission electron microscopy
TEOS : tetraethyl orthosilicate
THF : tetrahydrofuran
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LIST OF FIGURES
Figure 1.1. The principle of biosensor operation ........................................................... 17 Figure 1.2. Overlay of bright-field and epifluorescence images of live HeLa cells
labeled with orange QD and allowed to grow for the indicated period of time (scale bar = 5 µm). ............................................................................... 22
Figure 1.3. Comparison of signal photostability acquired by using Cy5- doped NPs and Cy5-fluorophores signaling in DNA sensors ........................................ 23
Figure 1.4. Functionalized PS NPs uptake by human macrophages, undifferentiated and PMA differentiated THP-1 cells. Cell viability was assessed by the XTT assay after 48 hours exposure to PS NPs. ........................................... 24
Figure 1.5. Transmission electron microscopy (TEM) images of poly(St-co-AA) latex particles with (a) 10 wt.-% and (b) 15 wt.-% acrylic acid; and poly(St-co-AA) latex particles with (c) 0 wt.-% and (d) 20 wt.-% 2-aminoethyl methacrylate hydrochloride. ..................................................... 25
Figure 1.6. Several common strategies for biomolecules immobilization onto the surface of silica NPs. .................................................................................... 27
Figure 1.7. Method for synthesizing silica NPs: (a) Stöber method and (b) reverse phase micro-emulsion .................................................................................. 28
Figure 1.8. Plot of silica-shell thickness as a function of TEOS concentration ............. 29
Figure 1.9. Schematic representation of SPA using antigen-coated latex particles in the absence (a) and in the presence (b) of antibodies. ................................. 30
Figure 1.10.Radiation energy spectrum for general β emitters (left) and 3H (right) ....... 33 Figure 1.11.Range spectrum of tritium (3H) and 125I in water. ....................................... 34
Figure 1.12.Illustration of SPA using (a) scintillant-doped beads and (b) scintillant- coated microplates ....................................................................................... 35
Figure 1.13.Emission spectra of several commercially available SPA beads. Imaging beads are made of PS or YOx matrices doped with Eu, whereas YtSi and polyvinyltoluene (PVT) beads are doped with Ce and diphenylanthracine, respectively. ................................................................................................. 36
Figure 1.14.Schematic diagram of the components of a PMT-based counter ................ 38 Figure 1.15.Inhibition curves of 4 different agonists of purified human estrogen
receptor α ligand-binding obtained from competitive binding assay using [3H]17β-estradiol. (○) 17β-estradiol, (�) 4-(1-adamantyl)phenol, (Δ) 4-tert-octyl phenol, (▲) 4-n-octylphenol. ................................................... 40
Figure 1.16.The schematic illustration of FlashPlate SPA for DNA polymerase (A) and the results of the assays for different concentration of double stranded-DNA precursors (B). .................................................................................... 41
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Figure 3.1. Structures of pTP (A) and dimethylPOPOP (B). The excitation and emission maxima are listed below each structure. The overlapping emission-excitation spectra for pTP and dimethylPOPOP (C) .................... 52
Figure 3.2. TEM images of PS core NPs (A) and PS core-silica shell NPs using TEOS/ MPTS (B) and PS core-silica shell NPs using TEOS/APTS (C). The bars on the bottom of the images indicate 500 nm lengths ................... 54
Figure 3.3. Size and size distribution of PS core NPs (A) and PS core-silica shell NPs (B) measured by Dynamic Light Scattering. Z-average diameters were 312±10 nm and 442±14 nm for PS core and PS core-silica shell NPs, respectively. Colored lines represent multiple measurement on the same particle sample .................................................................................... 56
Figure 3.4. Schematic representation of zeta potential: ion concentrations and potential differences as a function of distance from the surface of a charged particle suspended in a medium ..................................................... 58
Figure 3.5. ζ-potentials of PS core and PS core-silica shell NPs measured in 10 mM NaCl at pH 3-10. The error bars represent the standard deviation of three technical replicates ....................................................................................... 59
Figure 3.6. Emission spectra of PS core, scintillant-doped PS and PS core-silica shell NPs (λex = 270 nm). Spectra was collected using 0.05 mg/mL suspension of NPs in water ............................................................................................. 60
Figure 3.7. Scintillation responses of PS core, scintillant-doped PS and PS core-silica shell NPs to free 3H-sodium acetate as measured using 0.5 mg NPs in 1 mL sample (A) and after normalization to the number scintillating particles (B). The error bars represent the standard deviation of three sample replicates. ................................................................................ 61
Figure 3.8. Schematic illustration of SPA using DNA hybridization. When the tritiated complementary strand is annealed to the immobilized oligo-nucleotide, the energy from β emission excites the scintillants in the NP and light is then produced ............................................................................ 63
Figure 3.9. Immobilization of acrydite-ssDNA onto thiol-functionalized NPs probed using FITC-labeled complementary strand using (A) MES 0.1 M pH 5.4 for 6 h, (B) MES 0.1 M pH 5.4 for 24 h, (C) phosphate 0.1 M pH 7.4 for 24 h, and (D) phosphate 0.1 M pH 7.4 for 60 h. Results for control and sample were show n in left and right, respectively. Spectra colored in blue and red were collected prior to and after hybridization, respectively. Red spectra were corrected to the background spectra (i.e. blue spectra) to compensate particle loss during wash steps ............................................. 65
Figure 3.10. Estimating amount of immobilized oligonucleotide using FITC-labeled complementary strand. Fluorescence spectra of different concentrations of FITC-ssDNA were collected at λex 480 nm (A) and a calibration curve was established using the emission intensities at 520 nm (B). Emission spectra of ssDNA-immobilized NPs were obtained before and after DNA hybridization with FITC-ssDNA (C). Intensity of the after
10
hybridization spectrum was subtracted to the background intensity and concentration of hybridized probe was estimated using equation from the calibration curve (D) ............................................................................. 67
Figure 3.11. H-NMR spectra of NHS-acetate (A) and N-hexylacetamide in CDCl3 obtained after completion of first and second steps of EDC/NHS coupling reaction, respectively. Both of the compounds were extracted from the reaction mixture using DMF. A peak of 6 equivalent protons of DMSO is seen at δ 2.62 ppm. .................................................................. 70
Figure 3.12. Separation of radiolabeled oligonucleotide using Sephadex G-50 column. Peak 1 and 2 correspond to radiolabeled oligonucleotide and unreacted label, respectively ........................................................................................ 72
Figure 3.13. DNA hybridization on the developed SPA NPs. Specific activity of the 3H-labeled ssDNA was 57 nCi/nmol. Sample volumes were 5 mL. The error bars represent the standard deviations of three sample replicates ....... 74
Figure 3.14. Affinity binding studies for nonlabeled complementary, mismatch and random strands. Approximately 6 nmol of 3H-labeled complementary strand was added to saturate the binding sites on SPA NPs and establish 100% response. Sample volumes were 5 mL. The error bars represent the standard deviations of three technical replicates .......................................... 75
Figure 4.1. Conjugating bioreceptors onto NPs through: (A) zero-length crosslinking, (B) OEG tethered linkage, and (C) antibody-assisted immobilization ........ 78
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LIST OF SCHEMES
Scheme 3.1. Thermal decomposition reaction of AAPH radical initiator ........................ 53
Scheme 3.2. Mechanism of conjugation reaction between thiol-functionalized NPs and acrydite-oligonucleotide. ............................................................................. 64
Scheme 3.3. EDC/NHS coupling reaction for oligonucleotide radiolabeling .................. 69
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LIST OF TABLES
Table 3.1. Optimization of the step I of EDC/NHS coupling reaction condition. Concentrations of sodium acetate, EDC and NHS were 20 µM, 120 µM and 120 µM, respectively. The reaction length was 2 h. .......................................... 71
Table 3.2. Optimization of the step II of EDC/NHS coupling reaction condition in buffered solution. The first step of reaction was performed in MES buffer pH 5.3 (i.e. the optimized condition as shown at Table 3.1) .................................. 71
Table 3.3. Optimization of EDC/NHS coupling reaction condition in DMF-water combination. Step 1 and II were performed in DMF and phosphate buffer 0.1 M pH 8.0, respectively ............................................................................... 71
Table 3.4. Results of oligonucleotide radiolabeling using EDC/NHS coupling chemistry 73
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ABSTRACT
The development of analytical tools for investigating biological pathways on the
molecular level has provided insight into diseases and disorders. However, many biological
analytes such as glucose and inositol phosphate(s) lack the optical or electrochemical
properties needed for detection, making molecular sensing challenging. Scintillation
proximity assay (SPA) does not require analytes to possess such properties. SPA uses
radioisotopes to monitor the binding of analytes to SPA beads. The beads contain
scintillants that emit light when the radiolabeled analytes are in close proximity. This
technique is rapid, sensitive and separation-free.
Conventional SPA beads, however, are large relative to the cells and made of
hydrophobic organic polymers that tend to aggregate or inorganic crystals that sediment
rapidly in aqueous solution, thus limiting SPA applications. To overcome these problems,
polystyrene core-silica shell nanoparticles (NPs) doped with pTP and dimethyl POPOP
were fabricated to produce scintillation NPs that emit photons in the blue region of visible
light. The developed scintillation particles are approximately 250 nm in diameter (i.e. 200
nm of core diameter and 10-30 nm of shell thickness), responsive to β-decay from tritium
(3H) and have sufficient stability in the aqueous media.
DNA hybridization-based SPA was performed to determine whether the
scintillation NPs could be utilized for SPA applications. A 30-mer oligonucleotide was
immobilized on the polystyrene core-silica shell NPs to give approximately 7.6 x 103
oligonucleotide molecules per NP and 3H-labeled complementary strand was annealed to
the immobilized strand. At the saturation point, increases in scintillation signal due to
oligonucleotide binding to the NPs were about 9 fold compared to the control experiments
in which no specific binding occurred, demonstrating that the scintillation NPs can be
utilized for SPA. Along with the improved physical properties including smaller size and
better stability in the aqueous system, the developed scintillation NPs could be potentially
useful as biosensors in cellular studies.
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CHAPTER 1: INTRODUCTION
1. 1. Background
To respond to changes in their immediate environment, cells must be able to receive
and process signals or stimuli that come from the extracellular environment. This signaling
process involves extracellular messengers (primary messengers, e.g. hormones and
neurotransmitters), receptors, and intracellular messengers (secondary messengers).
Dysregulation of cell signaling components leads to diseases and disorders, including
diabetes,1 depression,2 and Alzheimer’s disease (AD).3 Thus, the capability to monitor
dynamic changes in signaling molecule concentrations within the cellular environments
will enable investigation of key signaling pathways for better understanding of the
mechanisms of diseases and disorders, and possibly allow for development of novel
therapeutics.
Several important small molecule carbohydrate messengers, such as glucose and
inositol phosphate, lack suitable properties for optical and/or electrochemical detection,
making identification and quantification of these messengers analytically challenging.
Multiple techniques have been developed for analysis of carbohydrates in biological
systems, such as electrochemical detection strategies including application of enzyme-
based electrodes4,5 and direct oxidation of carbohydrates at electrode surfaces6, polarized
photometric detection (PPD)7, and attachment of fluorescent tags to monosaccharide
building blocks.8 Nonetheless, some of these techniques rely upon a separation step such as
liquid chromatography and capillary electrophoresis to achieve a satisfactory degree of
chemical resolution, and cannot easily be utilized for cellular sensing since they do not
provide sufficient temporal and spatial resolution. While fluorescence labeling enables
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cellular imaging of carbohydrate analytes, this strategy may affect the physicochemical
properties of small biomolecules in cellular environment due to the comparatively large
size of the fluorescence reporters.
Radiolabels, on the other hand, may introduce smaller perturbations to analyte
properties and potentially enable more biologically relevant sensing. Radiolabeled
molecular probes have been employed in positron emission tomography (PET) and single-
photon emission computed tomography (SPECT) to study diseases at molecular levels.9,10
However, these techniques depend on the use of high-energy γ-rays which require very
cautious handling and relatively expensive equipment (both radioisotope generator and
imaging instrument). Utilization of less energetic β-emission has also been explored for
tomography purposes.11 Yet, the spatial resolution of this tomography technique is very
limited (i.e. in the order of mm) and thus, is not really useful for cellular imaging.
Scintillation proximity assay (SPA) was initially developed by Hart and Greenwald
in 1979.12 SPA uses radioisotopes to monitor the binding of analytes to SPA beads through
a biorecognition element covalently attached to the scintillating material. The beads contain
scintillants that absorb energy and emit light when radiolabeled analytes are in close
proximity (<1 µm). This technique is rapid, sensitive, and separation-free, thus serving as
an attractive alternative for cellular imaging of small biomolecules. However, many of the
benefits of SPA have not been conferred for this type of applications due to the size
constrains of commercially available beads (ca. 2-8 µm in diameter) which limit the
number of scintillating elements that may be incorporated into cells, therefore affecting the
spatial resolution of the technique. In addition, conventional SPA imaging beads are
usually made of hydrophobic polymeric materials, such as polystyrene, that tend to
aggregate in the aqueous systems.
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To enable SPA for cellular imaging, we fabricated scintillant-doped polystyrene
nanoparticles (ca. 100-250 nm) with thin silica shells (10-30 nm) which are compatible
with aqueous environments and have amenable surface chemistry for attachment of suitable
biorecognition elements.
1.2. Literature Review
1.2.1. Introduction to Biosensors
Sensors are devices that act as reporters of specific physical or chemical phenomena
of interest. These phenomena can range from a simple physical parameter such as
temperature to various complicated events such as biochemical processes occurring in a
living organism. With the rapid advances in medical technology, the ability to measure and
monitor different aspects in human physiology and biology such as intracellular and
extracellular concentrations of metabolites becomes requisite, as does developing enabling
sensing devices. Sensing devices that involve a biological component in their constructs are
referred to as biological sensors or biosensors.13 For example, a biosensor capable of
monitoring the plasma glucose is necessary for delivering effective treatment to a diabetic
patient.
In general, a biosensor comprises a biological or biologically derived sensing
element that is either integrated within or closely associated with a physicochemical
transducer that converts the biochemical signal into a quantifiable and processable
electrical signal. Typically, these biosensing elements are enzymes, multi-enzyme systems,
antibodies, membrane components, whole cells, or whole slices of mammalian or plant
tissues, which are responsible for the recognition of analyte(s) and contribute to the
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specificity and sensitivity of the final devices.13 For instance, glucose oxidase (an enzyme
that catalyzes the oxidation of glucose into gluconic acid and hydrogen peroxide) has been
used in the construction of amperometric glucose sensors.4,5,14 Figure 1.1 shows the
principle of how biosensors operate.15
Figure 1.1. The principle of biosensor operation. Reproduced from Vargas-Bernal et al. in Soundararajan, R.P., Pesticides-Advances in Chemical and Botanical Pesticides, 2012, pp 329-356.15
Biosensors can be classified into several categories based on their transducing
modes such as electrochemical sensors, piezoelectric sensors, thermal sensors and optical
sensors. Electrochemical biosensors rely on the detection of electroactive species resulting
from the interaction or reaction between analyte and the sensor bioelement. Based on the
parameter that is being measured, electrochemical sensors can be divided into
potentiometric sensors (measuring voltage) and amperometric sensors (measuring current).
Many potentiometric and amperometric biosensors have been developed for measuring
biological analytes.4,5,14,16,17 Piezoelectric effect occurs when a mechanical stress is applied
to the surface of a crystal and causes a corresponding electrical potential across the crystal
whose magnitude is proportional to the applied stress.18 Piezoelectric biosensors have been
employed to measure biochemical processes that involve mass change such as
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hybridization of a DNA strand to its immobilized complementary strand and binding of a
ligand to the corresponding immobilized receptor.19,20 Thermal biosensors exploit a
fundamental property of biological reactions, the absorption or the evolution of heat.
Metabolites such as glucose and urea have been determined using enzyme-immobilized
thermal biosensors.21 Optical biosensors work by collecting photons that are generated
upon interactions between analyte and bioelement of the sensors, and transforming the
photons into measurable electrical signals. Several examples of optical biosensors include
fluorescence bionsensors22, Förster resonance energy transfer (FRET) biosensors23,24,
surface plasmon resonance (SPR) biosensors25 and scintillation proximity assay (SPA)
biosensors.26,27
The miniaturization of biosensors into nanobiosensors has several advantages over
conventional micro and macrosized-biosensors including reducing the amount of sample
required for measurement and broadening their applications to cellular and in vivo studies.
Many optical nanobiosensors constructed using quantum dots, fluorescent proteins and
dye-doped nanoparticles have been reported for measurement of biological analytes in vitro
and in vivo.24,28
1.2.2. Roles of Nanomaterials in Biosensing Applications
Nanomaterials have found broad application in the field of analytical chemistry.
Due to their small size (typically in the range of 1-100 nm), they exhibit unique chemical,
physical, and electronic properties, which are beneficial for the construction of sensing
devices including biosensors. Nanoparticles with a high surface-area-to-volume ratio
(SA/V) and versatility for modification are suitable for immobilization of biological
elements needed for sensing applications. Various types of nanoparticles (NPs) including
19
metal NPs, oxide NPs, semiconductor NPs and polymeric NPs, have been widely employed
in electrochemical sensors and biosensors.4,17,29,30
In general, nanostructured materials can be fabricated by three main approaches:
top-down, bottom-up or a combination of both. In the top-down approach, bulk starting
materials are broken down into nanoscale structures using several methods including
photolithographic patterning, wet etching, plasma etching, laser processing, and
grinding.31–35 This approach can be used to produce NPs, nanotubes, nanorods and
nanowires of metal oxide, semiconductor, metal and polymer materials. Using the bottom-
up approach, nanostructures are formed by assembly of atoms or molecules by suitable and
controlled parameters and processes. Bottom-up synthesis can be done by vapor-phase
methods such as chemical vapor deposition (CVD) and plasma enhanced CVD36,37; or by
solution-phased methods such as sol-gel deposition, self-assembled monolayer and
microemulsion polymerization.38–40 Solution-phase bottom-up methods can be utilized to
fabricate polymeric and biological nanomaterials. Both approaches can also be combined to
produce more sophisticated nanomaterials such as anodized aluminum oxide (AAO)
nanoporous thin films.41
Attachment of biomolecules directly onto surfaces of bulk materials may result in
the loss of bioactivity. However, the immobilization of such molecules on the surfaces of
NPs can retain their activity due to the biocompatibility of NPs. For example, gold NPs can
immobilize proteins as the gold atoms form covalent bonds with the amine groups and
cysteine residues of proteins.42 Some proteins that have been successfully immobilized with
gold NPs include horseradish peroxidase, microperoxidase-11, tyrosinase and
hemoglobin.43–46 He and coworkers immobilized several heme proteins with SiO2 NPs
through layer-by-layer assembly.29 Other metal and metal oxide nanostructures such as Pt,
20
Ag, TiO2, ZrO2 can also be used for enzyme immobilization.47 Immobilization of glucose
oxidase in Pt NPs/graphene/chitosan nanocomposite film was reported for the construction
of glucose biosensors.4 DNA can be immobilized with NPs to assemble electrochemical
DNA sensors. Cai et al. immobilized mercaptohexyl-functionalized oligonucleotide onto
16-nm diameter gold nanoparticles and were able to obtain 0.5 nM of complementary
ssDNA for the detection limit of the DNA sensors.48
Nanostructures have also been useful for constructing many optical sensors for
biological imaging, measuring temperature and pH in living cells, and quantification of
biologically relevant analytes.49–51
1.2.3. Nanoparticles for Optical Imaging in Biological System
Among several modalities in molecular imaging such as magnetic resonance
imaging (MRI), X-ray computed tomography (CT) and single photon emission computed
tomography (SPECT), optical imaging has been a convenient approach in terms of
versatility to obtain or design probes or contrast agents and relatively low cost of
instrumentations. Optical probes that have been used for molecular imaging range from
organic dyes, fluorescent proteins, quantum dots (QDs) to dye-functionalized structures
such as dye-labeled oligonucleotides or peptides and dye-doped nanoparticles.49,52–56 QDs
and dye-doped nanoparticles are small enough to allow incorporation of a high number of
detectable tracers into the sample and are amenable for modification for targeting purposes,
thus making them excellent probes for optical imaging.
QDs are made from combinations of II-VI, III-V, and IV-VI group semiconductors
such as CdSe, CdS, and CdTe using a number of methods.57,58 The QDs are
photochemically stable and have tunable emission.59 In addition, QDs already have nm-
21
sized dimension and are intrinsically fluorescent, and hence can be readily conjugated with
biologically relevant molecules for imaging applications. For example, Jaiswal and
coworkers reported the use of avidin-conjugated QDs to image mammalian and D.
discoideum cells.45 Moreover, QDs can also be incorporated into larger NP structures such
as polystyrene NPs to yield organic-inorganic hybrid particles that could be used in
biological probing.60
Organic dyes (or fluorophores) are commonly large molecules. Most dyes emit
light in the visible region. However, many new fluorophores that operate in the near
infrared (near-IR) zone have been developed and offer advantages over traditional visible
light dyes. These near-IR dyes provide higher sensitivity (since cellular or tissue
components produce minimal autofluorescence in the near-IR region), give stronger tissue
penetration which is desirable for in vivo imaging and are less energetic (i.e. less toxic to
the living cells).56,61
Organic dyes can be integrated to the nanostructures by entrapping them inside the
particles, covalently binding them to the particle precursors prior to the assembly of NPs,
or immobilizing them on the surface of the NPs.54,55,62–65 Organic dyes can be entrapped
inside the NPs during the nucleation of the nanostructures or by swelling the polymeric
particles. Reisch et al. incorporated rhodamine B octadecyl ester (R18) into poly(D,L-
lactide-co-glicolide) nanoparticles (PLGA NPs) during the formation of NPs by nano-
precipitation.62 Fabrication of nigrosine-doped NPs, which involved particles swelling in
the mixture of organic and aqueous solvents to allow dye uptake followed by evaporation
of the organic solvent to shrink back the particles, was reported by Zhang and coworkers.55
Direct coupling provides another approach via covalent binding to the particle matrix.
Yang et al. covalently linked fluorescein isothiocyanate (FITC) to 3-
22
aminopropyltrimethoxysilane (APTMOS) prior to the assembly of silica NPs.64
Immobilization of organic dyes on the surface of NPs has also been demonstrated using
Ag-SiO2 and Au-SiO2 core-shell particles.54,65
In the field of biological imaging, NPs offer advantages to deal with many
limitations of conventional organic dyes such as poor photostability, low quantum yield,
and insufficient in vitro and in vivo stability.66 Jaiswal et al. demonstrated that QD
bioconjugates could label HeLa cells over an extended period of time (Figure 1.2).49 The
studies also showed that labels were still retained by the cells even after continuous growth
for 12 days. A superior photostability of dye-doped nanoparticles over free organic dye was
also reported by Zhou and coworkers.54 When used as transducer components in sensor
platforms, signal intensity of dye-doped NPs sensors only decreased to 95% of the initial
intensity after nine continuous scans while it dramatically dropped to 76% for the sensors
with fluorescent dye molecules (Figure 1.3). In addition, NPs are versatile for modification
Figure 1.2. Overlay of bright-field and epifluorescence images of live HeLa cells labeled with orange QD and allowed to grow for the indicated period of time (scale bar = 5 µm). Reproduced from Jaiswal et al., Nat. Biotechnol., 2003, 21, 47-51.49
23
and can be fabricated using suitable materials to improve the biocompatibility. For
instance, NPs can be fabricated using biocompatible and biodegradable materials such as
PLGA.62
Polymeric organic and inorganic materials are typically used to fabricate dye-doped
particles. Some examples include polystyrene, polyvinyl toluene, PLGA, poly(methyl
methacrylate), and silica.55,62,64,67 Polystyrene is one of the most popular polymers for
entrapping dyes especially organic soluble dyes due to its hydrophobicity and inexpensive
price.
1.2.3.1. Polystyrene Nanoparticles (PS NPs)
Polystyrene latexes exhibit many properties that are desirable for biosensors. PS
particles have narrow size distributions and the particle size can be easily controlled.68 PS
NPs are also stable under physiological conditions and hence can be traced in vitro and in
vivo over a longer period of time and can avoid many additional problems caused by the
Figure 1.3. Comparison of signal photostability acquired by using Cy5- doped NPs and Cy5-fluorophores signaling in DNA sensors. Reproduced from Zhou et al., Anal. Chem., 2004, 76, 5302-5312.54
24
polymer degradation .68,69 PS NPs have been successfully used to entrap hydrophobic
fluorescent dyes and quantum dots for both optical imaging for diagnostic purposes and
developing strategies for therapeutic treatment.60,68–70 Furthermore, PS NPs are widely used
as a biocompatible drug carrier model for in vitro and in vivo experiments due to their
characteristics of being immunogenic and less toxic to the cells.69,71 Lunov and coworkers
studied the internalization of 100 nm functionalized PS particles by human macrophages,
as well as by undifferentiated and PMA-differentiated monocytic THP-1 cells, and the
results showed that PS NPs are nontoxic (Figure 1.4).
Figure 1.4. Functionalized PS NPs uptake by human macrophages, undifferentiated and PMA differentiated THP-1 cells. Cell viability was assessed by the XTT assay after 48 h exposure to PS NPs. Reproduced from Lunov et al., ACS Nano, 2011, 5 (3), 1657-1669.71
PS particles are fabricated using micro-emulsion or mini-emulsion polymerization
techniques with or without surfactant stabilizers with the sizes ranging from tens of nm to
several µm in diameter.72,73 In micro-emulsion, the dispersed domain diameter varies
approximately from 1 to 100 nm (typically 10-50 nm), while in mini-emulsion it varies
from 50 nm to 1 µm.74 During the polymerization process, one or more types of surfactant
may be used to stabilize the droplets and achieve particles with narrow size distributions.
Atik and Thomas reported the use of cetyltrimethylammonium bromide (CTAB) as a
primary surfactant and hexanol as a co-surfactant (i.e. surfactant that acts in addition to
25
primary surfactant) to obtain narrowly distributed latex particles with diameters of 20-40
nm.75 Cationic surfactants, such as dodecyltrimethylammonium bromide (DTAB),
tetradecyltrimethylammonium bromide (TTAB), cetyltrimethylammonium bromide
(CTAB) and octadecyltrimethylammonium chloride (OTAC) are also useful to form
ternary micro-emulsions (i.e. surfactant/oil/water systems) for polymerization studies
without cosurfactants.76–78 Holzapfel and coworkers used sodium dodecylsulfate (SDS), an
anionic surfactant, to prepare carboxyl functionalized PS particles and non-ionic
surfactants, i.e. Lutensol AT50, for amino functionalized particles synthesis.68 It was also
found that the particle size increased gradually with increasing carboxyl monomer
concentration and decreased with increasing amino monomer concentration (Figure 1.5).
Figure 1.5. Transmission electron microscopy (TEM) images of poly(St-co-AA) latex particles with (a) 10 wt.-% and (b) 15 wt.-% acrylic acid; and poly(St-co-AA) latex particles with (c) 0 wt.-% and (d) 20 wt.-% 2-aminoethyl methacrylate hydrochloride. Reproduced from Holzapfel et al., Macromol. Chem. Phys., 2005, 206, 2440-2449.68
C D
A B
26
Since the use of surfactant(s) may introduce impurities to the polymer products and
demand additional purification steps for surfactant removal, several efforts to develop
soap-free emulsion polymerizations have been explored. Yamada et al. used 2,2’-azobis
[N-(2-carboxy-ethyl)-2-2-methylpropionamidine] hydrate, an amphoteric initiator, to
produce µm-sizes PS particles up to 6 µm via soap-free synthesis.73 The use of cationic
radical initiators including 2,2’-azobis(2-amidinopropane) dihydrochloride (AIBA) and
2,2’-azobis (2-(2-imidazoline-2-yl)propane) dihydrochloride (AIBI) was also reported by
Fritz and coworkers for the surfactant-free PS NPs synthesis.69 By using cationic initiators,
a positively charged particle surface was able to be generated to provide adsorption
mechanism for oligonucleotides via electrostatic interaction.
Unmodified PS particles are insoluble in water and tend to aggregate. However,
surface functionalization or co-polymerization with hydrophilic monomers and utilization
of cationic or anionic initiators to control the surface charge of PS nanoparticles have been
employed to overcome the aqueous incompatibility.68,69,73,75 Coating of PS NPs using
various hydrophilic materials such as silica, polyethylene oxide (PEO) and polyvinyl
acetate (PVAC) has also improved the hydrophilicity.79–81
1.2.3.4. Silica Coating of Nanoparticles
The use of silica as a coating material for inorganic and organic nanomaterials has
emerged due to its high stability (particularly in aqueous media), chemical inertness,
controlled porosity, optical transparency and versatility for subsequent functionalization for
various applications. The main factors supporting the excellent stability of silica sols are
that the van der Waals interactions are remarkably lower than those involving other NPs
and positively charged species can be tightly attached to the polymeric silicate layer at
27
silica-water interfaces under basic conditions.82 The chemically inert nature and
transparency over a wide range of the spectrum region (i.e. ultraviolet to near-infrared)
make silica favorable for the fabrication of many types of chemistry apparatuses and optics.
Silica-based materials including silica NPs and silica core-shell NPs are also amenable for
different strategies for immobilization of biomolecules for sensing applications. Figure 1.6
shows an illustration of common bioconjugation techniques for the attachment of biological
molecules onto the surface of silica-based NPs.
Figure 1.6. Several common strategies for biomolecules immobilization onto the surface of silica NPs. Reproduced from Yan et al., Nano Today, 2007, 2, 44-50.83
A number of approaches have been used to put silica coatings on NPs including sol-
gel synthesis (Stöber synthesis) and reverse micro-emulsion. Figure 1.7 shows a schematic
representation of the Stöber and reverse micro-emulsion techniques for preparing silica
NPs.84 In the Stöber method, hydrolysis and condensation of tetraethyl orthosilicate
(TEOS) is facilitated by base in the mixture of ethanol/water. In contrast, in the reverse
28
micro-emulsion procedure, TEOS is hydrolyzed at the micellar interface and enters the
aqueous droplet to form a silica nanoparticle within the micelle. As silica-coating
procedures generally involve surfaces with a significant chemical or electrostatic affinity
for silica, various modifications have been applied to the traditional Stöber synthesis to
facilitate coating silica on NPs that do not possess such surface properties. The use of
silane coupling agents has been reported for the fabrication of numerous metal-silica core-
shell particles.85–87 For example, Liz-Marzan and coworkers used (3-aminopropyl)-
trimethoxysilane as a surface primer to render gold colloids vitreophilic (glass-loving).85
The utilization of the amphiphilic, nonionic polymer poly-(vinylpyrrolidone) (PVP) as a
surface stabilizer prior to Stöber synthesis was also reported by Graf et al.88 This method
has been applied to silica-coat various colloidal particles such as small gold and silver
colloids (ca. 14-38 nm), large silver colloids (~532 nm), boehmite rods (~10 nm) and both
cationic and anionic polystyrene colloids (ca. 340-770 nm).88
Figure 1.7. Method for synthesizing silica NPs: (a) Stöber method and (b) reverse phase micro-emulsion. Reproduced from Taylor-Pashow et al., Chem. Commun., 2010, 46, 5832-5849.84
29
In reverse phase (also known as water-in-oil) micro-emulsions, micelles act as
nanoreactors that control the kinetics of particle nucleation and growth. This method allows
for careful control of size and size distribution of NPs, thus serving as a superior method
for making silica NPs smaller than 100 nm with very narrow size distributions compared to
the Stöber method.55 This approach can be used to coat relatively small NPs (ca. 10-50 nm)
and control the shell thickness with nm precision. Han and coworkers demonstrated reverse
micro-emulsion mediated synthesis for silica-coating gold and silver NPs.89 They were able
to make silica core-shell NPs with different sizes ranging from 24 nm to 80 nm from 12
nm-sized oleylamine-stabilized gold NPs.
For intracellular applications of core-shell particles, it is important to precisely
control the thickness of the silica shell since the cellular uptake of NPs is inversely
proportional to their size.90,91 The simplest way to control the shell thickness is by varying
the amount of silica precursor, such as TEOS. Kobayashi et al. reported the formation of
silica shells 28-76 nm thick on silver NPs for TEOS concentrations of 1-15 mM (Figure
1.8).92 In addition, the rate of silica-shell formation on NPs in sol-gel synthesis also
Figure 1.8. Plot of silica-shell thickness as a function of TEOS concentration. Reproduced from Kobayashi et al., J. Colloid Interface Sci., 2005, 283, 392-396.92
30
depends on the pH of the system. Park and coworkers observed that at basic pH (10.0),
there is no or little silica coated on the PS particles while at higher pH (12.5) the nucleation
of silica became rapid and not homogenous, thus leading to relatively rough silica shells.79
NPs with smooth surfaces and uniform thickness were able to be obtained at pH ca. 11 in
an isopropanol/water mixture.
Silica-based nanoparticles including silica core-shell NPs have found vast
applications in the field of bioimaging, biocatalysis, bioseparation and therapeutic
delivery.84,93–96
1.2.4. Scintillation Proximity Assay (SPA) Biosensors
Figure 1.9. Schematic representation of SPA using antigen-coated latex particles in the absence (a) and in the presence (b) of antibodies. Reproduced from Hart and Greenwald, Mol. Immunol., 1979, 16, 265-267.12
The concept of SPA was first introduced by Hart and Greenwald in 1979 as a
sensitive and convenient method of immunoassay.12 They demonstrated the direct detection
of rabbit antihuman albumin (RAHA) based on the interaction between tritiated latex
particles (LH) and scintillant-doped latex particles (L*) that both were covalently coated
31
with human albumin (HA). When RAHA was present in the sample, it became bound to the
antigens on the surface of the latex particles and brought LH and L* into proximity. The
energy from 3H β-emission excited the scintillants in L* and caused them to emit photons
that could be detected by a phototube (Figure 1.9).
In a more widely-used format, SPA sensors employ isotope-labeled analytes that
emit low-energy radiation, the energy of which can be easily dissipated into the aqueous
medium.97,98 Under these circumstances, the scintillant-doped beads produce no or low
signal when the analytes are not bound. A signal increase occurs upon binding of the
radiolabeled analytes. Unlike conventional radioimmunoassay (RIA), SPA does not require
wash steps, thus greatly reducing the amount of liquid waste generated. The technology
also lends itself to automation, which effectively reduces the assay time and labor.
The following sections will describe several integral components of SPA biosensors
including radioisotopes used for analyte labeling, material platforms and detection systems.
Several applications of SPA sensors in biomedical studies will also be presented.
1.2.4.1. Radioisotopes in SPA
Radioactive decay from an unstable atomic isotope (also referred to as
radioisotope or radionuclide) involves the emission of one or more types of subatomic
particles (e.g. α particles, β particles, Auger electrons and neutrons) or electromagnetic
radiations (e.g. γ-ray photons and x-ray photons) directly from either the nuclei or the
electron shells. Alpha particles, or He nuclei, consist of two protons and two neutrons.
These particles have relatively high decay energies (on the order of MeV), only travel a
few cm in air and are easily absorbed by thin layer of matter. Several α emitters include
241Am, 242Cm, 210Po, 239Pu and 226Ra.99 Beta particles are emitted from the decay of a
32
proton to produce a positron (positive electron) and a neutron, or from the decay of a
neutron to yield a negatron (electron) and an anti-neutrino.100 Energies associated with β
particles are slightly lower, ranging from keV to MeV. However, these particles can
penetrate up to several meters in air and up to few mm in water since they travel in much
higher velocity due to their low mass.99 Several beta emitters such as 3H, 125I, 14C and 35S
have found applications in the field of radioimmunoassays.101–104 Auger electrons are
atomic electrons that result from a physical process of filling inner-shell vacancies after a
core electron is removed due to irradiation with external X-rays or electron beams, or
emission of an internal conversion electron. Several radionuclides that emit Auger
electrons include 123I, 125I, 111In, and 119Sb.99 Neutrons are neutral particles that are only
stable in the nucleus of an atom and decay within minutes outside the nucleus. Unlike α
and β particles, neutrons are not emitted in any significant quantities from most of
radionuclides that undergo nuclear decay, except from 253Cf and 248Cm.99 Gamma and X-
rays are similar in that both are high-frequency ionizing radiations. However, X-rays are
produced from energy transitions of electrons whereas γ-rays originate from unstable
atomic nuclei. The energies of γ-rays are in the range of keV to MeV, while the energies of
X radiations are typically lower (in the order of eV to keV).
SPA makes use of the weak energy emission of certain radionuclides (i.e. the low-
energy β emitters). Beta emitters give a wide spectrum of energies that fall between 0 and
maximum energy (Emax) where the majority of emitted particles have the energy of
approximately ⅓ Emax (Figure 1.10). Some β emitters that have found applications in SPA
include 3H, 125I, 14C, 35S and 33P.12,26,27,105–107
33
Figure 1.10. Radiation energy spectrum for general β emitters (left) and 3H (right). Reproduced from L’Annunziata, in L’Annunziata, Handbook of Radioactivity Analysis, 2012, pp 1-162, and Kherani and Shmadya in Mannone, Safety in Tritium Handling Technology, 1993, pp 85-106.99,108
Tritium (3H) is widely used to label analytes in SPA. 97,98 The radionuclide has a half
life of 12.262 ± 0.004 y with a decay constant equal to 0.0565 per year or 1.791 x 10-9 per
second.109 The maximum β energy of 3H is 18.6 keV with the most frequent single β-
particle energy is 2-3 keV and an average energy close to 5.6 keV (Figure 1.10).108,110
Radiation from 3H travels a very small distance (i.e. the maximum and average ranges in
air are 6 mm and 0.5 mm, and the maximum range in water is only 6 µm).99,108 Compared
to other β emitters, 3H has the shortest travel distance in aqueous media and thus gives the
advantage of minimal background signal from non-proximity effects in SPA measurement.
Tritium has been successfully utilized in SPA for studying ligand-receptor binding,
measuring enzymatic activity and screening for drug candidates.26,111,112
125I is also commonly used for SPA and decays with a half-life of 60.25 d.70 It emits
a soft γ radiation, various internal conversion- and Auger-electrons with a maximum
energy of about 35 keV.113 Iodine-125 may also emit X-rays as a result of excess energy
from electron capture, internal conversion and subsequent changes in the electron orbitals.
Since their energies are slightly higher, 125I electrons can travel a few µm further in water,
34
compared to 3H beta particles (Figure 1.11). Similar to 3H, 125I has been applied in SPA for
monitoring ligand/receptor and antigen/antibody interactions and high throughput
screening for drug discovery.114,115
Radioisotopes with higher energy β-particle emission such as 14C, 35S and 33P are
also used to develop SPA platforms for different binding studies.105–107 14C, 35S and 33P
have maximum energies of 155 keV, 167 keV and 249 keV, respectively.99 The half-lives
of 35S and 33P are considerably shorter (87.4 d and 25.3 d, respectively) compared to 14C
(5730 y) to decay to the half of the initial isotope amount.99
Due to the fact that the specific activity of 14C is generally low, its use in SPA is not
typical. In addition, the path lengths of the radiations from 14C, 35S and 33P are quite long
(i.e. approximately 125 µm for 33P)116 and could potentially introduce high background
noise due to non-proximity effects at high concentrations. However, the use of SPA to
detect these isotopes may be advantageous for improving the detection of analytes present
at very low concentrations.
Figure 1.11. Range spectrum of tritium (3H) and 125I in water. Reproduced from Ertl and Feinendegen, Phys. Med. Biol., 1970, 15, 447-456.113
35
1.2.4.2. SPA Materials
SPA utilizes scintillant-doped beads or scintillant-doped polymer microplates
(Figure 1.12). SPA beads are approximately 2-8 µm in diameter and have scintillants
dispersed in the bead matrices. Commercial beads are available in a conjugated form with a
wide range of biomolecules such as antibodies, streptavidin, receptors, enzymes and small
molecules, such as glutathione.116 SPA plates containing plastic scintillator-coated wells to
which the biomolecules are bound are also commercially available in a 96- or 384-well
microplate format. The microplate format requires less than 400 µL working volume, thus
effectively reducing reagent consumption. SPA microplates are also amenable for
automation and hence, are well suited for high throughput applications.
A B Figure 1.12. Illustration of SPA using (a) scintillant-doped beads and (b) scintillant-doped polymer microplates.
In general, the materials used as matrices in SPA can be classified into inorganic
crystals and organic polymers (plastics). Inorganic materials used in commercial SPA
beads include yttrium silicate (YSi) and yttrium oxide (YOx). Inorganic matrices are denser
than organic polymers (i.e. 4.1g/cm3 and 5g/cm3 for YSi and YOx, respectively) and hence,
36
are settle rapidly and may cause dispensing issues.67 Ce ions are stably trapped within the
YSi crystal lattice to yield scintillation beads that emit violet and blue light (Figure 1.13).
The blue-emitting beads are most suitable in conjunction with photo multiplier tube
(PMT)-based scintillation counters. To perform faster scintillation measurements for
multiple samples, imaging beads that are made of Eu-doped YOx can also be employed in
a 96-, 384- or 1536-well microplate format. The imaging beads can emit a red light (Figure
1.13) that is best captured on a charged-couple device (CCD)-camera based detector. This
SPA imaging allows for simultaneous measurement for up to 1536 samples within
minutes.117 Eu-YOx imaging beads that emit light in the red region also overcome color
quenching issues from yellow and red colored samples.
Figure 1.13. Emission spectra of several commercially available SPA beads. Imaging beads are made of PS or YOx matrices doped with Eu, whereas YtSi and polyvinyltoluene (PVT) beads are doped with Ce and diphenylanthracine, respectively. Reproduced from PerkinElmer Inc., http://www.perkinelmer.com/resources/technicalresources/applicationsupportknowledgebase/radiometric/spa_bead.xhtml67
Two most popular organic polymer matrices in SPA beads technology are
polyvinyltoluene (PVT) and polystyrene (PS). These polymeric beads are less dense
compared to inorganic beads (i.e. 1.05 g/cm3 for PS), and thus remain in suspension better
37
and are more easily dispensed. However, due to their hydrophobicity, PS beads require
additional coating with a polyhydroxy film prior to surface functionalization with
biorecognition elements.116 Diphenylanthracine (DPA) that emits light in the blue region is
commonly doped inside PVT to produce plastic scintillators that work well with PMT-
based counters. Eu can also be co-polymerized in the PS matrix to make PS imaging beads.
Other polymer materials, poly(methacrylic acid) crosslinked with divinylbenzene (DVB),
have also been explored to prepare molecularly imprinted SPA beads whose sizes are
smaller than the commercially available beads.98 SPA microplates are fabricated by putting
a permanent coating of plastic scintillator inside the wells. Some of the microplates are also
designed to have hollow tubes, instead of the conventional wells, as solid supports for the
scintillator layer. Polystyrene is the main dispersant material for scintillating microplates
produced by PerkinElmer Inc. including ScintiPlateTM, FlashPlateTM, LumiPlateTM, and
Cytostar-TTM.
1.2.4.3. Detection and Measurement in SPA
Detectors for SPA must be very sensitive since scintillation signals are lower than
signals generated from other optical methods such as absorption and fluorescence
spectroscopy. Hence, photodetectors that enable signal amplifications such as
photomultiplier tubes (PMT), avalanche photodiodes and charged-couple device (CCD)
can be utilized for SPA. PMT-based counter and CCD camera-based imagers are widely
used for SPA detection and measurement. PMT-based counters employing standard bialkali
PMTs are well-suited for measuring signals from blue light emitting scintillation beads or
plates, whereas CCD camera imagers are optimal for capturing scintillation signals from
SPA beads that emit the red light.
38
Figure 1.14. Schematic diagram of the components of a PMT-based counter. Reproduced from L’Annunziata in L’Annunziata, Handbook of Radioactivity Analysis, 2012, pp 1021-1115.118
The components of a conventional PMT-based counter are schematically depicted
in Figure 1.14. In principle, photons emitted from the sample reach the adjacent PMTs
which then convert the optical signals into currents through a series of pulse amplification
at the dynodes of the photo-detectors. Two PMTs are generally used for measuring light
intensity from nuclear decay process. These PMTs allow for coincidence light detection
and pulse summation needed for detecting low-energy radioisotopes (e.g. 3H) and for
distinguishing a true nuclear event from instrument background. To be counted as a
scintillation event, photons must be detected in both PMTs within a coincidence resolving
time of approximately 18 ns.118 The output signals are typically expressed in counts per
minute (CPM) which represents the number of light flashes detected by the coincidence
detectors when the nuclear decays of the radionuclides are occurring. More recently
developed counters such as MicroBeta® and TopCount® are designed for faster sample
measurement as they can have up to 12 detectors that simultaneously measure 12 samples
at a time.119 Positioning the two coincidence PMTs above and below the sample could also
improve photon collection in low volume samples. In addition, a patented single PMT time
39
resolved counting method that discriminates between background and real counts by
analyzing the decay period of each scintillation event has been employed as well.
CCD camera-based imagers used in fluorescence microscopy are also applicable for
imaging scintillation events. Such imagers include LEADseekerTM from GE Healthcare and
ViewLux® from PerkinElmer.120,121 SPA imaging using CCD camera greatly reduces the
assay time compared to the counting system with PMT-based instruments. Typical plate
read times as rapid as 5 min can be easily achieved, and in some cases this could be
reduced to 2 min.122Another advantage of using SPA imaging for assays in microplates is
that signal decay due to measurement time across the plate in PMT-based readers, can be
avoided. This is particularly beneficial for SPA experiment employing beta emitters with
fast decays such as 33P.
1.2.4.4. Applications of SPA Biosensors in Biology and Medicine
Since the principle of SPA is detection upon analyte binding to its corresponding
biorecognition element-functionalized scintillation beads or plates, this technique has been
extremely useful for studies that involve binding interactions.12,26,105,111 Immunoassays
using SPA sensors have been reported in a number of studies.27,106,123 For example,
Udenfriend and coworkers demonstrated the measurement of tissue enkephalins, serum
thyroxin (T4) and urinary morphine using antibody-functionalized beads and 125I-labeled
antigens under inhibition mode.27 The results showed that SPA is comparable to
conventional RIA and the SPA technique was so sensitive that urine samples required
dilution prior to analysis.
40
SPA has been used heavily to provide information on the ligand-receptor
interactions.26,124,125 Figure 1.15 shows an example of receptor-binding assay done by Nose
et al. using purified human estrogen α ligand-binding domain and [3H]17β-estradiol in the
presence of 4 different unlabeled ligands.26 The inhibition curves depict the competitive
binding of various agonists to the receptor. Percentage of B/Bo expresses the ratio of
radioactivity measured in the presence of unlabeled agonists to radioactivity in the absence
of the agonists (i.e., no competing ligands in the solution). The relative strength of
interaction among several ligands can easily be determined based on the position of each
ligand plot. A ligand with a corresponding plot located on the left has a stronger interaction
with the receptor than those on the right. This relative competitiveness for receptor binding
can also be expressed using the IC50 term. The IC50 is calculated as the concentration
needed to inhibit 50% of the labeled ligands binding to the receptor. Similar binding
studies were performed by Dechavanne and coworkers to obtain correctly folded and
biologically active proteins in E. coli from 96 different refolding conditions.125
Figure 1.15. Inhibition curves of 4 different agonists of purified human estrogen receptor α ligand-binding domain obtained from competitive binding assay using [3H]17β-estradiol. (○) 17β-estradiol, (�) 4-(1-adamantyl)phenol, (�) 4-tert-octylphenol, (▲) 4-n-octylphenol. Reproduced from Nose et al. Toxicol. Lett, 2009, 33-39.26
41
Figure 1.16. The schematic illustration of FlashPlate SPA for DNA polymerase (A) and the results of the assays for different concentration of double stranded-DNA precursors (B). Reproduced from Earnshaw and Pope, J. Biomol. Screen. 2001, 6, 39-46.126
Several kinetic studies of enzyme activities have been also performed using SPA
biosensors.105,126 Earnshaw and Pope used FlashPlate SPA to characterize the activities of
DNA polymerase, primase and helicase.126 Figure 1.16 shows an assay format for
determining the activity of DNA polymerase. Single stranded DNA was immobilized onto
scintillant beads that were fixed inside the well of the microplate (i.e. FlashPlate). A shorter
complementary strand (DNA primer) was annealed to the immobilized strand and then
polymerase was added along with the labeled precursors for DNA elongation. As more
labeled precursors were incorporated into the growing double stranded DNA by
polymerase, more energy from β decay was transferred to the scintillants to produce
42
photons. The activity of DNA polymerase was observed under different concentrations of
deoxyribose adenosine triphosphate (dATP) precursor for up to 30 min (Figure 1.16). The
results of this assay were comparable to those obtained from the conventional filtration-
based polymerase assay. In another enzyme SPA, Jeffery et al. used γ-33P-labeled inorganic
orthophosphate to obtain kinetic analysis on adenosine triphosphatase (ATPase). 105
ATPase catalyzed the hydrolysis of 33P-labeled ATP to produce radioactive inorganic
phosphates that reacted with ammonium molybdate. The reaction generated radioactive
phosphomolybdate anions which were adsorbed onto SPA beads. To minimize the non-
proximity signals due to the longer radiation path length of 33P, a solution of 3.5 M CsCl
was added to float the SPA beads to the surface of the suspension prior to the
measurements.
Since microplate SPA can be used for measuring hundreds of samples within
minutes (i.e., approximately 40 min using PMT-based counter for a 384-well microplate
and 2-5 min using CCD camera-based imager), this technique has a great potential for high
throughput screening of drug candidates. Huss and coworkers reported the application of
SPA to screen 49 plant metabolites for inhibition of cyclooxygenase-2 (COX-2) catalyzed
prostaglandin E2 (PGE2) production.111 Tritiated PGE2 tracer was added to compete with
unlabeled PGE2 produced by COX-2 catalyzed reaction for binding to antiPGE2-
functionalized SPA beads. As the reaction inhibitors were added, the amount of unlabeled
PGE2 was suppressed and more labeled PGE2 bound to the beads to produce signals. Of the
49 metabolites that were screened, eugenol, pyrogallol and cinnamaldehyde were found to
inhibit COX-2 with the IC values in the µM regime.111 Several other works that involved
ligand screening using SPA include screening for inhibitors of various kinases127,128 and for
PPARγ modulators.129
43
SPA biosensor technology could potentially be applied for molecular studies such
as detection of analytes in a single living cell if the sensors are small enough to allow
adequate amount of tracers becoming incorporated into the cell, compatible and non-toxic
to the cell.
44
CHAPTER 2: MATERIALS AND METHODS
2.1. Materials
The primary scintillant p-terphenyl (pTP), secondary scintillant 4-bis(4-methyl-5-phenyl-
2oxyzolyl)benzene (dimethylPOPOP) and styrene were obtained from Acros Organics
(Geel, Belgium). Tetraethylorthosilicate (TEOS), (3-mercaptopropyl)trimethoxysilane
(MPTS), 2,2’-azobis(2-methylpropionamidine) dihydrochloride (AAPH), N-(3-
dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride (EDC HCl), N-
hydroxysuccinimide (NHS), fluorescein isothiocyanate (FITC) and Tween-20 were
obtained from Sigma Aldrich (St. Louis, MO, USA). Acrydite functionalized DNA
oligomer: 5’-/Acryd/AAA CCC TGT AAA CCC TGT AAA CCC TGT AAA-3’ and amine
functionalized DNA oligomers: 5’-/5AmMC6/TTT ACA GGG TTT ACA GGG TTT ACA
GGG TTT-3’ (complementary), 5’-/5AmMC6/TTT ACA GAG TTT AGA GGG TTG
ACA GGG ATT (mismatch), and 5’-/5AmMC6/AGC ACT GAG AGA AGA GTG TTG
ACA CAT ATT-3’ (random) were obtained from Integrated DNA Technologies, Inc.
(Coralville, IA, USA). Tritiated sodium acetate (1.54 Ci/mmol) was obtained from Perkin
Elmer (Boston, MA, USA).
2.2. Fabrication of Scintillant-doped PS NPs
PS core particles were prepared using a surfactant-free emulsion polymerization as
previously described with several modifications.100 Styrene was filtered through an alumina
column to remove the stabilizer and weighed to approximately 3.0 g. The styrene was then
combined with 50 mL degassed nanopure water (18.3 MΩ.cm) in a 250 mL round-
bottomed flask equipped with a magnetic stir bar. A gentle stream of Ar was flowed
45
through the flask for 15 min before the flask was sealed using a rubber cap. Polymerization
was initiated by adding 0.3 mL of 10 mg/mL AAPH solution into the water/styrene
mixture. The mixture was stirred quickly and maintained at 80 ± 5°C in a water bath for 6
h. Unreacted styrene was removed using a rotary evaporator and PS NPs were rinsed
several times with water, isopropanol and water again. The PS NPs were suspended in
water and the concentration of NPs in the stock suspension was estimated by freezing and
then lyophilizing 1 mL of the suspension overnight, and weighing the dry particle residue.
To incorporate scintillants into PS NPs, the NPs were swollen in a mixture of
aqueous-organic solvents using a method described by Benhke et al.130 with modifications.
Approximately 500 mg of PS core NPs was combined with 80 µmol pTP and 8 µmol
dimethylPOPOP in 50 mL mixture of water-tetrahydrofuran (43:7). The mixture was
shaken at 300 rpm for 1 h at room temperature and the NPs were then rinsed several times
with water. The dye-doped PS NPs were suspended in water and the concentration of NPs
in the stock suspension was estimated as described above.
2.3. Silica Coating of Scintillant-doped PS NPs
Thiol functionalized silica shells were added to scintillant-doped PS NPs using
modified Stöber synthesis.100 A 14 mg sample of particles was suspended in a 500 mL
round-bottomed flask containing 240 mL mixture of water-isopropanol (1:5). The mixture
was stirred briskly using a magnetic stir bar at room temperature and 7.5 mL 28% NH4OH
was added to the flask. After several minutes, 2 mL of TEOS-MPTS (9:1) was added
dropwise to the flask. Stirring was continued for 1 h after which the NPs were rinsed
several times with ethanol followed by water. The core shell NPs were suspended in water
and the concentration of NPs in the stock suspension was estimated as described above.
46
2.4. Characterization of PS NPs
The characterization of the NPs includes size determination using Transmission
Electron Microscopy (TEM) and Dynamic Light Scattering (DLS) techniques, zeta
potential measurement, emission spectra acquisition and measurement of scintillation
response towards 3H sodium acetate.
2.4.1. Size and Morphology Determination by TEM
PS core NPs and PS core-silica shell NPs were imaged without staining using
TecnaiTM G2 Spirit Transmission Electron Microscope (FEI Company, USA) operated at
100 kV accelerating voltage.
2.4.2. Size and Size Distribution Determination by DLS
A very dilute suspension of PS core NPs and PS core-silica shell NPs in 5 mM
phosphate buffered saline, pH 7.0 (PBS) was transferred into disposable folded capillary
cell and analyzed using Zetasizer Nano (Malvern Instruments Ltd, UK). The measurements
were performed in quintuplicate.
2.4.3. Zeta Potential Measurements
Zeta potential measurements were performed in a zeta dip cell using Zetasizer Nano
(Malvern Instruments Ltd, UK). PS core NPs and PS core-silica shell NPs were dispersed
in 10 mM NaCl and suspension pH was varied from 3 to 10. Zeta potentials of the samples
were calculated using Smoluchowski approximation (1.5) as the solution for the Henry
equation. All measurements were performed in triplicate.
47
2.4.4. Emission Spectra Acquisition
Emissions of PS NPs, scintillant-doped PS NPs and PS core-silica shell NPs were
scanned in the range of 300-500 nm using QuantaMasterTM spectrofluorometer (Photon
Technology International Inc., USA) with excitation wavelength set at 270 nm (i.e.
wavelength for maximal absorption of polystyrene). Each measurement was performed
using 1 mL of 0.05 mg/mL NPs suspension in water, placed in a semi-micro quartz
fluorescence cuvette.
2.4.5. Scintillation Response Measurements
Scintillation responses of PS NPs, scintillant-doped PS NPs and PS core-silica shell
NPs were determined by subjecting 1.0 mL of 0.5 mg/mL NPs suspension in water to
different activities of 3H acetic acid (i.e. 0, 0.5, 1.0, 2.0, 5.0 and 10.0 µCi). Tween-20 was
added to the suspension to a concentration of 0.1% w/v to prevent particle aggregation
and/or sedimentation. All samples were prepared in triplicate and measured three times
each using Beckman LS 6000IC Scintillation Counter (Beckman Coulter Inc., USA).
2.5. SPA Proof-of-Concept using DNA Hybridization
To demonstrate that SPA can be performed using the developed core-shell NPs, a
30-mer oligonucleotide or single-stranded DNA (ssDNA) was immobilized onto NPs and
the3H-labeled complementary strand was annealed to the immobilized probe. To evaluate
binding selectivity, three different ssDNA strands (i.e. complementary, mismatch by 4
bases, and random) were tested under SPA inhibition mode.
48
2.5.1. Immobilization of ssDNA on PS Core-Silica Shell NPs
Single-stranded DNA was immobilized onto the core-shell NPs using a similar
method to that described by Nuhiji et al.131 with some modifications (described later in
Results and Discussion). A 50 mg sample of core-shell NPs was dispersed in 1 mL
phosphate buffer 0.1 M pH 7.4 and was reacted with 80 nmol acrydite-functionalized 30-
mer ssDNA for 72 h at room temperature with brisk stirring. Particles were rinsed several
times with the phosphate buffer and resuspended in the same buffer. Control NPs were
prepared using the same procedure by omitting the ssDNA from the reaction mixture.
To probe ssDNA immobilization on the NPs, the immobilized DNA was annealed
to an FITC-labeled complementary strand and fluorescence spectra were collected prior to
and after DNA hybridization using an excitation wavelength of 480 nm. A 0.5 mg sample
of ssDNA-functionalized NPs was dispersed in 1 mL annealing buffer (100 mM phosphate,
50 mM NaCl and 10 mM EDTA; pH 7.4) and was mixed with 1 nmol FITC-labeled
complementary strand. The mixture was heated at 70°C for 5 min and allowed to cool
down to room temperature for 40 min while being shaken at 300 rpm. The NPs were rinsed
several times using annealing buffer with 0.1% Tween-20 and were suspended in 1 mL of
same annealing buffer prior to fluorescence spectra acquisition. The amount of
immobilized DNA was estimated by subtracting the emission spectrum of particles after
FITC-ssDNA annealing to that before the annealing process and comparing the subtracted
intensity to FITC-ssDNA calibration curve.
2.5.2. Radiolabeling of Complementary ssDNA
Complementary ssDNA was tritiated by conjugation with 3H-sodium acetate using
EDC/NHS coupling chemistry. To optimize the coupling reaction, a model reaction using
49
non-tritiated sodium acetate and hexylamine was performed under several different
conditions (described later in Results and Discussion). The reaction intermediate and/or
final product was isolated and yield percentages were determined using 1H-Nuclear
Magnetic Resonance Spectroscopy (Avance III 400 MHz NMR Spectrometer, Bruker
Corp., USA) with dimethylsulfoxide (DMSO) as an internal standard. The optimized
procedure was then applied to the ssDNA radiolabeling.
100 nmol of 3H-sodium acetate (corresponded to 154 µCi activity) was reacted with
10 µmol EDC and 10 µmol NHS in 30 µL dimethylformamide (DMF) by stirring the
solution for 3 h at room temperature. 100 nmol of amine-functionalized oligonucleotide
was dissolved in 90 µL phosphate buffer 0.1 M pH 9.5 and mixed with the 3H-sodium
acetate/EDC.HCl/NHS solution. The mixture was stirred briskly for 3 h at room
temperature and then separated using a Sephadex G-50 column (length = 11 cm, internal
diameter = 6 mm) upon reaction completion. Column fractions were subjected to
scintillation counting and specific activity of radiolabeled ssDNA was estimated as follows:
Specific activity of 3H-ssDNA = !"#$ !" !!!!!"#$ !"#$
!"#$% !"#! x
!"# !!"!"" !"#$
2.5.3. DNA Hybridization on PS Core-Silica Shell NPs
A 5 mg sample of ssDNA-functionalized core-shell NPs were dispersed in 5 mL
annealing buffer containing 0.1% Tween-20 in a 7 mL-scintillation vial. The NPs were then
titrated with increasing amounts of 3H-labeled complementary ssDNA (i.e. 0, 0.2, 0.5, 1, 2,
3, 4, 5 and 6 nmol). DNA was hybridized by heating the samples at 70°C (i.e.
approximately 10° above its melting temperature (Tm) which is 59.8°C) for 5 min and
allowing them to cool to room temperature for 40 min while shaking at 300 rpm prior to
50
each scintillation measurement. As a control, non-functionalized core-shell NPs were also
subjected to similar hybridization procedure. All samples were prepared in triplicate and
measured three times each.
The affinity of immobilized ssDNA to three different 30-mer ssDNA strands (i.e.
complementary, mismatch by 4 bases, and random) was also tested using SPA inhibition
mode. A 5 mg sample of ssDNA-functionalized NPs were saturated with 3H-labeled
complementary strand (approximately 6 nmol of 3H-ssDNA) and titrated with increasing
amounts of non-labeled complementary, mismatch or random strands (i.e. 0, 1, 2, 3, 4, 5
and 6 nmol). Prior to each scintillation measurement, samples were heated at 70°C for 5
min and allowed to cool to room temperature for 40 min while shaking at 300 rpm. Results
were expressed as percentages of 3H-ssDNA bound to NPs which is the ratio of
scintillation count after each addition of non-labeled ssDNA to that in the absence of non-
labeled strand (i.e. 100% binding sites are occupied with 3H-labeled ssDNA).
51
CHAPTER 3: RESULTS AND DISCUSSIONS
3.1. Fabrication and Characterization of Scintillant-doped Core-Shell NPs
Polystyrene particles have been widely used as a SPA solid support/matrix due to
their inexpensive price, well-studied characteristics, and established fabrication methods.
Polystyrene (PS) itself shows luminescent characteristics of two emission peaks with
maxima at 290 nm and 330 nm when excited at its first absorption band (240-270 nm),
which makes it a scintillator itself.132 However, due to the low quantum yield of PS,
incorporation of primary scintillants and even secondary scintillants (wavelength shifting
agent) with higher quantum yields is necessary to enhance scintillation signals. In this
work, pTP and dimethylPOPOP were specifically used as primary and secondary
scintillants, respectively, to produce PS NPs that emit light in the blue region of the visible
spectrum. The structures of pTP and dimethylPOPOP are shown in Figure 3.1. Both
scintillants have high quantum yields (i.e. Φf are equal to 0.93 for both pTP and
dimethylPOPOP in cyclohexane)133 and the excitation spectrum of dimethylPOPOP greatly
overlaps with the emission spectrum of pTP (Figure 3.1C), thus enabling significant
resonance energy transfer from primary scintillant to secondary scintillant.
Primary and secondary scintillants are typically added to the liquid scintillation
cocktail in the range of 2-10 g/L and 0.5-1.0 g/L respectively.134 In scintillation particles,
however, the concentrations of the primary scintillant are relatively high (i.e. about 16.0 to
40.0 wt % of the dry solid weight) whereas the concentrations of secondary scintillant are
generally in the range of 0.001-0.2 wt %.135 Since the emission of β-particles is isotropic,
the portion of absorbed energy by the matrices of scintillation particles would be much
lower than that by the solvent in liquid scintillation cocktails. Therefore, having a higher
52
number density of scintillants within the scintillating beads will allow for a higher
probability of more scintillants become excited by the β-emission and improve the
detection limit of SPA. The ratio of secondary scintillant to primary scintillant in both
liquid scintillation cocktail and scintillating particles is usually lower than 1:10 to minimize
signal quenching due to direct energy transfer from scintillation matrices to the secondary
scintillants.
Figure 3.1. Structures of pTP (A) and dimethylPOPOP (B). The excitation and emission maxima are listed below each structure. The overlapping emission-excitation spectra for pTP and dimethylPOPOP (C).
PS core NPs were prepared through a surfactant-free emulsion polymerization
method using the cationic azo initiator AAPH. This technique omits the difficult surfactant
removal steps that are necessary in conventional emulsion polymerization methods.
Cationic initiators also create partially positive charges on the surface of the formed PS
NPs that help prevent particles from aggregating and facilitate adsorption of negatively
charged silane precursors for the silica shell deposition process. Scheme 3.1 shows the
decomposition reaction of AAPH to form carbon radicals and N2. The rate of
decomposition of AAPH is dictated primarily by temperature and, to a minor extent, by pH
and solvent composition. The half-life of AAPH is approximately 175 h at 37°C in a
N
O O
N
λex = 278 nm; λem = 345 nm
λex = 370 nm; λem = 430 nm
A
B
0
0.2
0.4
0.6
0.8
1
1.2
270 320 370 420 470
Nor
mal
ized
inte
nsity
(AU
)
Wavelength (nm)
Excitation of dimethylPOPOP Emission of pTP
C
53
neutral pH and decreases to 10 h at 56°C.136 Polystyrene synthesis through AAPH radical-
initiated polymerization is typically performed at temperatures > 70°C to increase the rate
of radical formation.69,137 By increasing this rate, propagation of monomers into polymer
occurs more rapidly and given polymer sizes can be achieved in a shorter time period. In
addition, a higher concentration of initiators increases the number of active centers that can
grow into polymer chains, thus leading to higher quantity of particles produced during the
polymerization. In this work, PS nanoparticles with diameters less than 300 nm were
typically obtained by using AAPH at 0.1 % w/w to monomer weight (or 60 µg/mL final
concentration in the mixture) and holding the reaction temperature at 80°C for 6-12 h.
Smaller particles can be achieved by lowering the concentration of initiator. However, this
strategy is not practical for large-scale synthesis as it also decreases the conversion of
styrene to polystyrene and lowers the reaction yield. Another approach to fabricate smaller
NPs is by lowering the monomer concentration in the water/oil system. Fritz and coworkers
varied the concentration of styrene monomer from 0.48 mol/L to 0.29 mol/L and obtained
an approximately 15% size decrease in the fabricated PS NPs.69
Scheme 3.1. Thermal decomposition reaction of AAPH radical initiator
Primary and secondary scintillants were entrapped in the PS core NPs via swelling-
diffusion procedures using water-THF (43:7) mixture. This solvent composition allowed
for approximately 80% incorporation of hydrophobic dyes into PS NPs in a study
C C N
CH3
CH3NH2
HN N C
CH3
CH3
C NH
NH2
heatC C
CH3
CH3NH2
HN + N N
AAPH
C C N
CH3
CH3NH2
HN N C
CH3
CH3
C NH
NH2
heatC C
CH3
CH3NH2
HN + N N
AAPH
2
54
performed by Behnke et al,130 where a THF volume fraction of more than 14% led to
decrease in dye entrapment as the increase in THF volume might hamper diffusion of the
dyes into PS matrices. Additionally, in the absence of surfactants, THF concentrations
higher than 33% in the aqueous-organic system also caused PS particles to aggregate and
form larger clumps.138 Another solvent system using chloroform/isopropanol has also been
reported for incorporating water-insoluble dyes into PS NPs.130 As a general rule, although
the dyes must be soluble in the solvent mixture, the ratio of dye solubility in the solvent to
dye solubility in PS should be minimal to drive dye diffusion into PS NPs.
Modified Stöber synthesis using a mixture of TEOS-MPTS (9:1) was utilized in this
work to form thiol-functionalized silica shells necessary for immobilization of acrydite-
conjugated recognition elements. The silica shells further encapsulated the hydrophobic
dyes inside the nanostructures and provided hydrophilic surfaces that prevented particle
aggregation in the aqueous medium.
3.1.1. Morphology, Size and Size Distribution
Figure 3.2. TEM images of PS core NPs (A) and PS core-silica shell NPs using TEOS/MPTS (B) and PS core-silica shell NPs using TEOS/APTS (C). The bars on the bottom of the images indicate 500 nm lengths.
A B C
55
Spherical and smooth PS NPs approximately 200 nm in diameter (Figure 3.2A)
were obtained from the surfactant-free emulsion polymerization using AAPH as confirmed
by transmission electron microscopy (TEM). The diameters measured in TEM images were
smaller than the hydrodynamic diameters (Z-average diameter) of the NPs measured by
dynamic light scattering (DLS), which was 312 ± 10 nm (Figure 3.3A). Z-average diameter
(Dz) is an intensity-weighted harmonic mean calculated from scattering intensity of the
particles (Si) and diameter the particles (Di) (Eq 3.1). In DLS measurement, an intensity
weighted average diffusion coefficient (Dt,avg) is calculated by cumulants analysis based on
the scattering signals and Dz is calculated using Stoke-Einstein equation (Eq. 3.2).139 kB is
Boltzman’s constant, T is temperature, and η is viscosity of the dispersant.
D! = !!
(!! !!)
D! = !! !
!"#!!,!"#
Although water theoretically does not wet hydrophobic surfaces such as those of PS
core NPs, the presence of positive charges on the surface of NPs from AAPH might cause
hydration of the particles, resulting in an apparent increase in particle diameter during DLS
measurement. The discrepancy might also occur due to particle shrinkage as the NPs were
dried prior to TEM measurements. Furthermore, the Z-average size generated by DLS
measurement is only comparable to other techniques when the sample is mono-modal (i.e.
only one peak), has spherical shape and very narrow size distribution and does not form
aggregates. As shown in Figure 3.3A, a secondary peak with a maximum at about 5 µm,
which was probably due to particle aggregation or other particulates, was also present and
affected the particle size determination by DLS.
3.1 3.2
56
Figure 3.3. Size and size distribution of PS core NPs (A) and PS core-silica shell NPs (B) measured by Dynamic Light Scattering. Z-average diameters were 312 ± 10 nm and 442 ± 14 nm for PS core and PS core-silica shell NPs, respectively. Colored lines represent multiple measurements on the same particle sample.
PS NPs coated using a combination TEOS and MPTS showed coarse shell
structures that looked like ‘berries’ (Figure 3.2B), unlike core shell NPs fabricated using
TEOS and (3-aminopropyl)triethoxysilane (APTS) (Figure 3.2C) and TEOS only79 whose
surfaces were relatively smooth and homogenous. This suggests that some portions of silica
had undergone nucleation and growth in the medium prior to adsorption onto the PS NPs.
Small and large silica particles with diameters up to about 700 nm were also observed by
TEM (Figure 3.2B, right). These particles might be resulted from a continuous growing of
silica nuclei in the solution and a combination of multiple growing silica chains. The
thicknesses of silica shells were in the range of 10-30 nm for most of the particles, bringing
the average diameter of thiol-functionalized core-shell NPs to about 250 nm. The average
A
B
57
hydrodynamic diameter of the core-shell NPs was 442 ± 14 nm as determined by DLS
(Figure 3.3B).
Since there were at least three types of NPs formed after silica coating procedures
(i.e. PS core-silica shell NPs, bare PS core NPs and silica NPs), the size distribution of the
NPs became slightly broader as seen from Figure 3.3. Polydispersity indexes (PdI) for PS
core NPs and PS core-silica core-shell NPs obtained from Zetasizer® instrument were
0.203 ± 0.02 and 0.288 ± 0.03, respectively. PdI is a dimensionless parameter that is
calculated based on cumulant analysis in a DLS experiment and reflects the heterogeneity
of sizes of particles in a sample. PdI values smaller than 0.05 are rarely seen, except with
highly monodisperse standards and PdI values greater than 0.7 indicate that the sample has
a very broad size distribution.140 Typically, particles whose PdI values are less than 0.2 are
considered monodisperse.141 However, the “monodisperse” term itself is self-contradictory
and the use of this term to express the uniformity of the particle size is not suggested.142
3.1.2. Surface Charge Characterization
ζ-potential is a measure of the effective electric charge on the NPs surface. When a
particle has a net surface charge, ions of opposite charge will be adsorbed on the surface to
form the Stern layer (Figure 3.4).143 The Stern layer is then surrounded by a diffuse region
in which ions are less strongly associated. Within the diffuse layer, is a hypothetical
boundary, the surface of hydrodynamic shear or slipping plane, inside of which the ions
and particles form a stable entity. ζ-potential is measured at this boundary and its
magnitude provides information about particle stability within the medium. NPs with ζ-
potentials close to 0 mV tend to agglomerate or aggregate whereas NPs with higher
magnitude of ζ-potentials (either + or -) will be more stable as they tend to repel each other.
58
Surface charges of PS core and PS core-silica shell NPs were characterized by
measuring their ζ-potentials at pH 3-10 in 10 mM NaCl. In zeta potential measurements, an
electrical field is applied across the sample and the electrophoretic mobility is measured
from the light scattering of the particles. The Henry equation (Eq. 3.3) is then used to
calculate the ζ-potential:
U! = ! ! ! !(!!)
! ! (3.3)
where Ue is the electrophoretic mobility, ε is the dielectric constant, η is the viscosity of the
medium, and f(κa) is the Henry function for the ratio of the particle radius, the Debye
length. The Henry function can be approximated as 1.5 (Smoluchowski model) for sample
measurements made in aqueous medium with salt concentrations ≥ 10 mM.144
The ζ-potential of PS core NPs was 49.2 ± 0.6 mV at pH 3.0 and decreased in an
approximately linear fashion from pH 3 to 7 (Figure 3.5). The ζ-potential values did not
significantly change from pH 7-9 (i.e. steady at about 14-16 mV) and then decreased to -
11.7 ± 2.5 mV at pH 10.0. The positive ζ-potentials at the pH lower than 10.0 might be due
to the presence of cationic initiator (AAPH) which gave an overall positive charge on the
Figure 3.4. Schematic representation of zeta potential: ion concentrations and potential differences as a function of distance from the surface of a charged particle suspended in a medium. Reproduced from Liese and Hilterhaus, Chem. Soc. Rev., 2013, 42, 6236-6249.143
59
Figure 3.5. ζ-potentials of PS core and PS core-silica shell NPs measured in 10 mM NaCl at pH 3-10. The error bars represent the standard deviation of three technical replicates.
surface of the NPs. The amidine moiety of AAPH is a strong base with a pKa value equal
to 27.1.145 Thus, most of the amidine groups will be in the protonated form even in a very
basic pH and give overall positive charge on PS NPs over a wide range of pH. However,
since the amount of initiator was considerably small compared to the whole PS matrix and
PS itself has negative charges due to electron density of the aromatic rings,146 the ζ-
potential approaches negative values with increasing pH.
PS core-silica shell NPs had overall negative surface charge at pH 3-10 as
confirmed by ζ-potential measurements. ζ-potential of core-shell NPs was -6.61 ± 1.0 mV
at pH 3.0 and became increasingly negative as the pH increased. This can be attributed
primarily to deprotonation of silanol groups and, to a minor extent, deprotonation of thiol
groups with increasing pH. At neutral pH, ζ-potentials of PS core-silica shell NPs and PS
core NPs were -35.4 ± 3.4 mV and 16.1 ± 2.0 mV, respectively, supporting the idea that
silica coating on PS NPs increased the stability of NPs in the aqueous medium, in the sense
of preventing NPs aggregation.
-60
-40
-20
0
20
40
60
3 4 5 6 7 8 9 10
ζ-po
tent
ials
(mV
)
silica-PS core-shell NPs scintillant-doped PS NPs
pH
PS core NPs PS core-silica shell NPs
60
3.1.3. Emission Spectra of Scintillating NPs
Figure 3.6. Emission spectra of PS core, scintillant-doped PS and PS core-silica shell NPs (λex = 270 nm) collected using 0.05 mg/mL suspension of NPs in water.
Emission spectra of PS core, scintillant-doped PS and PS core-silica shell
nanoparticles upon excitation at 270 nm (i.e. wavelength for maximum absorbance of PS)
are shown in Figure 3.6. PS shows maxima at 295 nm and 307 nm whereas emission
maxima the scintillants can be seen at 329 nm and 343 nm for pTP and at 405 nm and 425
nm for dimethyl POPOP. Resonance energy transfer from PS matrices to the scintillants
can be observed from the decrease in emission intensity of PS after the scintillants were
doped into the NPs. Thus, the emission intensity of pTP and dimethyl POPOP might be a
summation of fluorescence from direct excitation by the light source and fluorescence from
resonance energy transfer. When measured at a similar suspension concentrations (w/v), PS
core-silica shell NPs gave approximately five times lower emission intensity than the
scintillant-doped PS NPs. This could be attributed to the presence of silica shells and silica
NPs that dominated the sample weight since there was no separation done after silica
coating procedures, and/or the leaching of the dyes during the coating process.
0
50000
100000
150000
200000
250000
300000
350000
400000
450000
500000
290 310 330 350 370 390 410 430 450 470 490
Em
issi
on in
tens
ity (A
U)
Wavelength (nm)
PS core NPs scintillant-doped PS NPs silica-PS core-shell NPs
PS core NPs Scintillant-doped PS NPs PS core-silica shell NPs
61
3.1.4. Scintillation Response to Free 3H
Figure 3.7. Scintillation responses of PS core, scintillant-doped PS and PS core-silica shell NPs to free 3H-sodium acetate as measured using 0.5 mg NPs in 1 mL sample (A) and after normalization to the number scintillating particles (B). The error bars represent the standard deviation of three sample replicates.
Scintillation responses of PS core, scintillant-doped PS and PS core-silica shell NPs
to free 3H were measured at fixed suspension concentration (i.e. 0.5 mg NPs suspended in 1
mL water with increasing activities of 3H-sodium acetate. The results shown in Figure 3.7
indicates that the highest scintillation response was given by scintillant-doped PS NPs,
followed by PS core-silica shell and PS core NPs. As also supported by the fluorescence
spectra, incorporating scintillants into the PS NPs significantly increased the emission of
NPs and hence, elevated the scintillation signals. However, the scintillation response of PS
core-silica shell NPs was considerably low and only about one tenth of that from NPs with
no shells (Figure 3.7A). This lower scintillation signal might be attributed to fewer
scintillation particles per weight unit and leaching of scintillants in isopropanol-water
mixture during core-shell formation. Figure 3.7B shows scintillation responses normalized
to the number of scintillating particles (i.e. assuming dye leakage from NPs is not
0
200
400
600
800
1000
1200
1400
1600
1800
0 1 2 3 4 5 6
Cou
nts p
er m
inut
e (C
PM)
Activity (μCi)
PS core NPsscintillant-doped PS NPssilica-PS core-shell NPs
0
200
400
600
800
1000
1200
1400
1600
1800
0 2 4 6
Nor
mal
ized
CPM
Activity (µCi )
n PS core NPs Δ scintillant-doped PS NPs u PS core-silica shell NPs
n PS core NPs Δ scintillant-doped PS NPs u PS core-silica shell NPs
A B
62
significant and therefore the emission intensities in Figure 3.6 are directly proportional to
the number of scintillating NPs).
Lower scintillation responses of the PS core-silica shell NPs might be due to a
faster sedimentation of the heavier hybrid particles. In addition, the overall positive charge
on the surface of non-shell NPs might also cause nonspecific adsorption of 3H acetate ions
which led to an increase in scintillation signals. As this adsorption was not likely to happen
on the core-shell NPs due to the repulsion of acetate ions by negatively charged silica
shells, the difference in scintillation responses between scintillant-doped PS and PS core-
silica shell NPs became even more notable.
3.2. Scintillation Proximity Assay using DNA Hybridization
DNA hybridization is a process in which two complementary single-stranded DNA
or RNA molecules bind to each other to form a double stranded molecule through base
pairing. Hybridization is central to many important biochemical techniques, e.g.
polymerase chain reaction and Southern blotting. The thermodynamics of DNA
hybridization is also quite well understood, making this process easily driven by altering
the temperature. The temperature at which 50% of the oligonucleotides are in the duplex
form with a perfect complementary strand is the melting temperature (Tm), which can be
calculated using thermodynamic basis sets for neighbor interactions as shown in Eq. 3.4.147
T! = !! ⋅!"""
!!! ! !" (!!! )− 237.15+ 16.6 log[K!] (3.4)
ΔH and ΔS are the enthalpy and entropy for helix formation, respectively, R is the molar
gas constant (1.987 cal/K.mol), Ct is total concentration of the DNA strands and [K+] is the
63
concentration of K+ or Na+ in the annealing buffer. The affinity of single-stranded DNA to
its complementary strand increases with increasing sequence length and decreases with
increasing temperature. Stevens and coworkers reported dissociation constant (Kd) values
ranged from 10-11 M at 20°C to 10-7 M at 70°C for a 20-mer DNA immobilized on
microparticles.148 A Kd value of 2.6 x 10-11 M was reported by Okahata et al. for a 30-mer
oligonucleotide immobilized on a quartz crystal microbalance at 20°C.149
DNA hybridization-based SPA was performed to determine whether the
scintillation NPs could be utilized for nanoSPA applications. A 30-mer oligonucleotide was
immobilized on the PS core-silica shell NPs and 3H-labeled complementary strand was
annealed to the immobilized strand such that the radionuclide is brought into proximity to
the NPs and more energy from β emission can excite the scintillants (Figure 3.8).
Figure 3.8. Schematic illustration of SPA using DNA hybridization. When the tritiated complementary strand is annealed to the immobilized oligonucleotide, the energy from β emission excites the scintillants in the NP and light is then produced.
3.2.1. Immobilization of Oligonucleotide on PS Core-Silica Shell NPs
Acrydite-conjugated oligonucleotide was immobilized through direct coupling to
thiol-functionalized silica shells. This reaction proceeds by Michael addition where a
nucleophile reacts with an α,β-unsaturated carbonyl compound. Michael additions are
64
typically carried out in organic solvents in the presence of strong bases such as
triethylamine, N,N- diisopropylethylamine (DIPEA), 1,8-diazabicycloundec-7-ene (DBU),
n-alkylamine and trialkylphosphine.150 Although the reaction is rarely done in an aqueous
solution, Nuhiji and coworkers found it useful for immobilizong acrydite-DNA onto thiol-
functionalized microspheres using 2-(N-morpholino)ethanesulfonic acid (MES) buffer pH
5.4.131
Scheme 3.2. Mechanism of conjugation reaction between thiol-functionalized NPs and acrydite-oligonucleotide.
Since the Michael reaction favors basic conditions such that nucleophiles are
deprotonated and can attack the Cβ carbocation of the acrydite group (Scheme 3.2),
immobilization of ssDNA on the NPs using MES buffer pH 5.4 was limited. As seen from
Figure 3.9A, there was only small increase in sample emission intensity as compared to the
control experiment. Extending the reaction from 6 h to 24 h increased the yield by roughly
two fold (Figure 3.9B). Yet, considering that the number of reactive nucleophiles is limited
at pH 5.4 ( only 0.0006% of total thiol groups are deprotonated since the pKa of
O
NH
P OHOH
O
S-
SO-
NH
P OHOH
O
H OH
S
O
NH
P OH
OH
O
H2O
OH
acrydite
thiolate group on NPs
α
β
65
Figure 3.9. Immobilization of acrydite-ssDNA onto thiol-functionalized NPs probed using FITC-labeled complementary strand using (A) MES 0.1 M pH 5.4 for 6 h, (B) MES 0.1 M pH 5.4 for 24 h, (C) phosphate 0.1 M pH 7.4 for 24 h, and (D) phosphate 0.1 M pH 7.4 for 60 h. Results for control and sample were shown in left and right, respectively. Spectra colored in blue and red were collected prior to and after hybridization, respectively. Red spectra were corrected to the background spectra (i.e. blue spectra) to compensate for particle loss during wash steps.
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66
mercaptopropyl group is 10.65),151 the immobilization yield should be further improved by
performing the reaction at a higher pH. The caveat is, however, that the reaction may not be
carried out above pH= 9 since silica will be quickly dissolved.152 Liu and coworkers reported
that percentages of dissolved silica from the porous hollow silica NPs at pH 7.0 and 8.0
were 7.32% and 23.4%, respectively. Nonporous silica NPs should dissolve in a slower rate
as the surface area that is in contact with water is less than that in porous NPs. Nonetheless,
the study done using those porous silica NPs suggests that there could also be a significant
increase in silica dissolution nonporous silica NPs and silica shells at pH above 8.0. Thus, to
improve the reaction rate while also maintaining the integrity of the silica shell, phosphate
buffer pH 7.4 was utilized for the immobilization. As expected, the immobilization was
more efficient at pH 7.4 since the percentage of deprotonated thiols increases to
approximately 0.06% (Figure 3.9C). Also, by extending the reaction time to 60 h, more
ssDNA became immobilized (Figure 3.9D).
To estimate the amount of oligonucleotides that were successfully immobilized on
the PS core-silica shell NPs, the emission intensity of the ssDNA-functionalized NPs after
hybridization with the FITC-labeled complementary strand was subtracted from the
background intensity (i.e. scattering spectra of NPs before the addition of FITC-labeled
probe) and the concentration of hybridized probes was estimated using FITC-ssDNA
calibration curve (Figure 3.10). Assuming all hybridized probes annealed to the
immobilized ssDNA by 1 to 1 ratio, approximately 0.42 nmol ssDNA became attached on
0.5 mg NPs, thus giving slightly greater than 50% immobilization efficiency as calculated
from the total amount of oligonucleotide added into the reaction.
67
Figure 3.10. Estimating amount of immobilized oligonucleotide using FITC-labeled complementary strand. Fluorescence spectra of different concentrations of FITC-ssDNA were collected at λex 480 nm (A) and a calibration curve was established using the emission intensities at 520 nm (B). Emission spectra of ssDNA-immobilized NPs were obtained before and after DNA hybridization with FITC-ssDNA (C). Intensity of the after-hybridization spectrum was subtracted from the background intensity and concentration of the hybridized probes was estimated using equation from the calibration curve (D).
Considering PS core-silica shell NPs with 200 nm core diameters and 25 nm shell
thicknesses, 1 mg of NPs will be equivalent to about 6.7 x 1010 particles and 131 cm2
surface area. The surface coverage of thiol groups on the NPs was 2.2 x 105 moieties per NP
or 1.1 x 1014 moieties/cm2. The oligonucleotide was immobilized with a surface coverage of
7.6 x 103 molecules per NP or 3.9 x 1012 molecules/cm2 particle surface. Thus,
approximately 3.5% of the total thiol groups on the NPs were converted into oligonucleotide
conjugates.
0 10000 20000 30000 40000 50000 60000 70000 80000
500 520 540 560 580 600
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Wavelength (nm)
0.42 μM
A B
C D
Emission intensity = 72000 [FITC-ssDNA] + 1000 R2 = 0.99
0.05 µM 0.1 µM 0.2 µM 0.3 µM 0.5 µM 1.0 µM
68
3.2.2. Oligonucleotide Radiolabeling
To radiolabel the complementary strand, amine-terminated oligonucleotide was
conjugated to 3H-sodium acetate through EDC/NHS coupling. Carbodiimides such as EDC,
N,N'-dicyclohexylcarbodiimide (DCC) and N,N'-diisopropylcarbodiimide (DIC) are zero-
length crosslinking agents that are widely used to mediate the formation of amide or
phosphoramidate linkage between a carboxylate group and an amine or a phosphate and an
amine, respectively.153–156 DCC and DIC are used to perform coupling chemistry in organic
solvents such as DMF whereas EDC can be used for bioconjugation in the aqueous media.
In EDC coupling, carboxylic acid reacts with EDC to form O-acylisourea active ester
which then reacts with a primary amine to form a stable amide bond. Formation of O-
acylisourea is optimal at pH 4-5. However, this intermediate is severely susceptible to
hydrolysis, making the coupling reactions less efficient. The use NHS and its more soluble
form, sulfo-NHS, in addition to EDC had been explored to improve the stability of the
reaction intermediate and improve the reaction efficiency.157,158 The reaction between
carboxylic acid and EDC/NHS produces NHS ester that is more stable than O-acylisourea
and also amine-reactive. The NHS ester is hydrolyzed within hours or minutes, depending
on water-content and pH of the reaction solution.159,160
Oligonucleotide radiolabeling was initially performed using a standard two step
EDC/NHS coupling procedure (Scheme 3.3)161. Approximately 4 nmol 3H sodium acetate
was mixed with 40 nmol EDC and 100 nmol NHS in 20 µL MES buffer 0.1 M pH 6 and
allowed to react for 15 min. Amine-terminated oligonucleotide (2 nmol) was then dissolved
in 50 µL phosphate buffer 0.1 M pH 7.4 and reacted with the acetate solution for 2 h.
Unfortunately, no detectable yield was observed. Since a tritiated compound was used in the
experiment, troubleshooting the coupling issue was difficult due to the limited availability
69
Scheme 3.3. EDC/NHS coupling reaction for oligonucleotide radiolabeling
of analytical tools that are designated for use with radioactive materials. Thus, a model
reaction using non-tritiated sodium acetate and hexylamine was used to optimize the
reaction conditions. Several different conditions were investigated and the yields (i.e.
reaction intermediate (NHS-acetate) and/or final product (N-hexylacetamide)) were
monitored using 1H-NMR Spectroscopy using DMSO as an internal standard. Figure 3.11
shows examples of NMR spectra collected upon completions of step I and step II of the
EDC/NHS coupling. Table 3.1, 3.2 and 3.3 summarize the reaction conditions used in the
trials and their corresponding yield percentages.
The formation of NHS-ester (i.e. step I) in aqueous system was optimum at pH 5.3,
giving approximately 41.2% yield or 0.082 mmol NHS-acetate to react with hexylamine in
step II (Table 3.1). The final product was formed at a much lower efficiency. Even with 20
to 1 ratio of NHS ester to hexylamine (assuming that the amount of NHS-ester formed from
step I is roughly equivalent for all reactions done using the same parameters), the yield was
3H
3H-CH2
3H-
3H-
3H-CH2
CH2
-3H
70
Figure 3.11. H-NMR spectra of NHS-acetate (A) and N-hexylacetamide in CDCl3 obtained after completion of first and second steps of EDC/NHS coupling reaction, respectively. Both of the compounds were extracted from the reaction mixture using DMF. A peak of 6 equivalent protons of DMSO is seen at δ 2.62 ppm.
N
O
O
O
O
N-succinimidyl acetate
2.84
2.842.34
A
B
71
Table 3.1. Optimization of the step I of EDC/NHS coupling reaction condition. Concentrations of sodium acetate, EDC and NHS were 20 µM, 120 µM and 120 µM, respectively. The reaction length was 2 h.
No Solvent % Yield
1 Water 13.1
2 10% ethanol 4.7
3 MES buffer pH 6.0 5.2
4 MES buffer pH 5.3 41.2
Table 3.2. Optimization of the step II of EDC/NHS coupling reaction condition in buffered solution. The first step of reaction was performed in MES buffer pH 5.3 (i.e. the optimized condition as shown in Table 3.1)
No Sodium acetate-hexylamine mol ratio pH Length (h) % Yield
1 10 : 1 5.3 12 5.1
2 10 : 1 7.2 6 10.6
3 50 : 1 7.2 6 12.3
4 10 : 1 8.0 4 10.4
Table 3.3. Optimization of EDC/NHS coupling reaction condition in DMF-water combination. Step 1 and II were performed in DMF and phosphate buffer 0.1 M pH 8.0, respectively
No
Step I
Intermediate purification
Step II
% Yield Sodium acetate (mM)
EDC (mM)
NHS (mM)
Length (h)
Sodium acetate/ ssDNA
mol ratio
Length (h)
1 100 100 100 2 Yes 10:1 4 81.0
2 100 100 100 2 No 10:1 4 59.0
only 12.3% (Table 3.2). Performing the first step using DMF, followed by extraction of the
reaction intermediate and continuation of the second step in phosphate buffer pH 8.0, on
the other hand, gave a considerably higher yield (i.e about 81%, Table 3.3). This suggests
72
that there may be competing reactions that hinder the formation of N-
hexylacetamide when the excess reactants from step 1 were not removed. Nonetheless,
as this intermediate purification cannot be employed when radioactive materials are used
due to radiation safety compliance, another trial using DMF as solvent for the first step,
followed by addition of buffered solution containing hexylamine was also performed. By
omitting the purification step, the reaction yield was slightly lower (ca. 59%), supporting
the previous argument that the presence of reactants (and probably by-products as well)
from the first step perturbed the NHS-ester/amine reaction. Since this DMF/water
solventsystem gave a relatively high yield compared to the reactions performed in water,
this protocol was then applied for the oligonucleotide radiolabeling.
To approximate the radiolabeling efficiency, labeled oligonucleotide was separated
from unreacted label using a Sephadex G-50 column and the radioactivity of the fractions
was determined using liquid scintillantion counting. The results were then plotted to
establish a chromatogram as shown in Figure 3.12 and the amount of radiolabeled
oligonucleotide was quantified based on the peak area.
Figure 3.12. Separation of radiolabeled oligonucleotide using a Sephadex G-50 column. Peak 1 and 2 correspond to radiolabeled oligonucleotide and unreacted label, respectively.
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2
73
Although a similar EDC/NHS protocol was used to radiolabel the oligonucleotide,
only about 1.2% of oligonucleotide became conjugated to the tritiated acetate. This might
be due to the lower concentrations of the reactants (i.e. 1 mM 3H-sodium acetate and 25
µM amine-terminated oligonucleotide) which limited the reaction rate, and an inherently
slower kinetic attributed to the oligonucleotide. Several conditions, summarized in Table
3.4, were explored to maximizing the labeling efficiency, though the maximum yield
achieved was 7.1%. As seen in Table 3.4, using DMF only resulted in no product
formation, suggesting that the solubility of oligonucleotide in DMF may be very low. This
solubility problem may also contribute to the low yields for reactions of oligonucleotide
radiolabeling in DMF/water mixture compared to that of model reaction using hexylamine.
Table 3.4. Results of oligonucleotide radiolabeling using EDC/NHS coupling chemistry
No
Step I Step II
% Yield Solvent
3H-Na acetate (mM)
EDC (mM)
NHS (mM)
Length (h) Solvent
3H-Na acetate/ ssDNA
mol ratio
Length (h)
1 DMF 1 100 100 2 phosphate 0.1 M pH 8.0 10:1 4 1.2
2 DMF 1 100 100 2 phosphate 0.1 M pH 8.0 10:1 1 1.8
3 DMF 3.33 333 333 3 phosphate 0.1 M pH 9.5 5:1 1 7.1
4 DMF 3.33 333 333 4 phosphate 0.1 M pH 9.5 1:1 1 3.7
5 DMF 1 100 100 2 - 5:1 24 -
3.2.3. DNA Hybridization on PS Core-silica Shell NPs
Despite low radiolabeling yields, DNA hybridization was explored to evaluate SPA
on oligonucleotide-functionalized PS core-silica shell NPs. The results of the DNA SPA
experiment are shown in Figure 3.13. As expected, the scintillation signals for binding
74
Figure 3.13. DNA hybridization on the developed SPA NPs. Specific activity of the 3H-labeled ssDNA was 57 nCi/nmol. Sample volumes were 5 mL. The error bars represent the standard deviations of three sample replicates.
experiment were higher than the control experiment (i.e. non-proximity). When
radiolabeled oligonucleotide hybridizes to its immobilized complementary strand, the 3H
label is brought into proximity to the scintillation NPs, allowing for higher probability of
the energy from β emission to excite the scintillants. Instead of yielding a linear response
with increasing concentrations of radiolabeled analyte, the curve shape was rather
sigmoidal. Sigmoidal binding curves typically occur in receptor-ligand interactions when
the affinity of the ligand increases after some initial binding of ligands to the receptor.162 In
that case, the bound ligand might change the allosteric structure of the protein and promote
the binding of next ligand. However, as the binding of 3H-oligonucleotide to its
immobilized recognition element occurs in 1 to 1 ratio, this change of affinity was not
expected. The non-linear response was more likely due the aggregation of particles during
the heating step in denaturation/annealing process. Each data point in Figure 3.13 was
obtained by titrating a set of oligonucleotide-modified particles with increasing amounts of
0
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150
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250
300
350
0 1 2 3 4 5 6 7
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Complementary
Control
Proximity (SPA)
Non-proximity
75
3H-labeled complementary strand, followed by heating to 70°C and cooling down to room
temperature prior to measurements, not by adding a series of different analyte
concentrations to separate samples. Although the aggregates are usually resolvable
macroscopically by sonicating/vortexing the sample, multiple heating steps might
accumulate the smaller aggregates. Due to aggregation that occurred over the time course,
scintillation particles might become packed close to one another and the β emission could
excite more scintillants than when the particles were not associated.
Figure 3.14. Affinity binding studies for nonlabeled complementary, mismatch and random strands. Approximately 6 nmol of 3H-labeled complementary strand was added to saturate the binding sites on SPA NPs and establish 100% response. Sample volumes were 5 mL. The error bars represent the standard deviations of three technical replicates.
To test the binding affinity of the immobilized strand, three oligonucleotides with
different sequences (complementary, mismatch by 4 bases and completely random) were
allowed to bind to the SPA NPs under inhibition mode. Addition of non-labeled
oligonucleotides to SPA NPs that are already saturated with 3H-complementary strand will
cause competitive binding between 3H-labeled and nonlabeled strands, thus reducing the
0
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120
0 1 2 3 4 5 6 7
% 3 H
-labe
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com
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enta
ry st
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76
number of bound 3H-labeled strands and its corresponding scintillation signal. As
predicted, the complementary strand had the greatest affinity to the SPA NPs, followed by
the mismatch and the random sequences (Figure 3.14). Interestingly, the IC50 of the non-
labeled complementary strand was less than 0.6 µM while in fact the concentration of 3H-
labeled complementary strand in the mixture was 1.2 µM. This slight discrepancy may also
be attributed to the multiple time-rehybridization processes.
Compared to other SPA experiments done by Gal and coworkers utilizing the
binding of 3H-labeled oligonucleotide,163 the scintillation responses obtained in this study
were lower by roughly two orders in magnitude. This is mainly due to low specific activity
of 3H-oligonucleotide made in-house using EDC/NHS coupling. The specific activity of
3H-oligonucleotide used here was 57 mCi/mmol, which was about 7000 times lower than
that used in their study (i.e. 420 Ci/mmol), indicating a lower number of 3H label per mole
unit of oligonucleotide molecules. Low specific activity also indicates a large number of
molecules without 3H label in the oligonucleotide mixture. As these non-labeled
oligonucleotides also compete for the binding sites, they could limit the amount of bound
3H-labeled strands and the corresponding scintillation signals. Nonetheless, despite the low
specific activity of the radiolabeled analyte, low nanomole amount of analyte was still
detected, demonstrating that the developed SPA NPs would be even more promising when
used with commercially available tritiated analytes whose specific activities are typically in
the range of 0.1-60 Ci/mmol.
77
CHAPTER 4: SUMMARY AND FUTURE DIRECTIONS
4.1. Summary
Core-shell silica-polystyrene NPs (nominal diameter ca. 200 nm) were synthesized
and doped with pTP and dimethyl POPOP to produce scintillation NPs that emit photons in
the blue region of visible light. The core-shell scintillation NPs were responsive to free 3H
and gave linear scintillation response with increasing activity of 3H. The developed
scintillation NPs was also tested for scintillation proximity assay (SPA) using DNA
hybridization. Oligonucleotide-functionalized NPs (ca. 7.6 x 103 molecules per NP) were
titrated with increasing concentrations of 3H-labeled complementary strand (specific
activity = 57 mCi/mmol). Increase in scintillation signal due to oligonucleotide binding to
the NPs were detected compared to the control experiments in which no binding occurred,
demonstrating that the scintillation NPs can be utilized for SPA. Some attributes of the
fabricated scintillation particles including their small sizes (< 300 nm in diameters) and
sufficient stability in the aqueous media make them potentially useful as biosensors in
cellular studies.
4.2. Future Directions
The ultimate goal of this research is to apply SPA to cellular sensing of small
molecules, particularly carbohydrate messengers such as glucose and inositol phosphate.
Future works will involve construction of scintillation particles that are responsive to these
analytes. 3H-labeled glucose and inositol phosphate with high specific activity (ca. 25-50
Ci/mmol) are commercially available, thus reducing the complications that may come from
low efficiency in-house analyte radiolabeling. Macromolecular receptors including proteins
78
and aptamers can be explored for construction of the SPA NPs. Covalent linkage between
receptors and NPs is definitely preferred over direct adsorption as this will ensure receptor
association with scintillation NPs in the biological environment. Zero-length conjugation
using carbodimides such as EDC and 1-cyclohexyl-3-(2-morpholinoethyl) carbodiimide
(CMC) can be utilized for this purpose due to the presence of amine and carboxylate
moieties in proteins and phosphate groups in aptamers.164 However, as these moieties
spread throughout the macromolecular structures, caution must be taken as the linkage may
also hinder access to the binding sites of these receptors (Figure 4.1A). Additional tethers
such as oligo(ethylene glycol) (OEG) may be used to provide additional degrees of
freedom of the conjugated receptors and prevent protein denaturation on the surface of NPs
(Figure 4.1B).165 Antibody assisted receptor immobilization (Figure 4.1C) could also be
employed.163 Nevertheless, this strategy is only useful if the binding of receptor proteins to
the antibody does not affect their ligand binding pockets.
Figure 4.1. Conjugating bioreceptors onto NPs through: (A) zero-length crosslinking, (B) OEG tethered linkage, and (C) antibody-assisted immobilization.
Multicolor scintillation particles will potentially enable simultaneous detection of
different analytes in the cellular setting. Thus, fabrication of such particles will also be a
79
part of future works. Organic dyes with large Stoke shift such as 3-hydroxyflavone (3HF),
2-(2-hydroxyphenyl)-benzothiazole (HBT), and 1-phenyl-3-mesithyl-2-pyrazoline (PMP)
can be doped into PS NPs as primary scintillants with or without additional dye as a
wavelength shifter as their emissions are already in the 400-600 nm region of the visible
light.166–168 Utilization of tertiary scintillants in addition to pTP and dimethyl POPOP can
also be explored to red-shift the emission of currently developed NPs. Moreover, the use of
quantum dots would also be advantageous for preparing these multicolor NPs since their
emission peaks are narrower than typical organic dyes, therefore preventing or minimizing
emission overlap between one type scintillation NP and another type NP (e.g. blue and
green emitting NPs).
80
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