Inorganic Nanoparticles as Donors in Resonance Energy Transfer for

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Inorganic Nanoparticles as Donors in Resonance Energy Transfer for Solid-Phase Bioassays and Biosensors Yi Han, M. Omair Noor, Abootaleb Sedighi, Uvaraj Uddayasankar, Samer Doughan, and Ulrich J. Krull Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.7b01483 • Publication Date (Web): 31 Jul 2017 Downloaded from http://pubs.acs.org on August 1, 2017

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Inorganic Nanoparticles as Donors in Resonance Energy Transfer for Solid-Phase Bioassays and Biosensors

Yi Han, M. Omair Noor, Abootaleb Sedighi, Uvaraj Uddayasankar, Samer Doughan, Ulrich J. Krull*

Chemical Sensors Group, Department of Chemical and Physical Sciences, University of Toronto Mississauga, Mississauga, Ontario, Canada, L5l 1C6

[email protected]

ACS Paragon Plus Environment

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ABSTRACT

Bioassays for the rapid detection and quantification of specific nucleic acids, proteins and peptides are fundamental tools in many clinical settings. Traditional optical emission methods have focused on the use of molecular dyes as labels to track selective binding interactions, and as probes that are sensitive to environmental changes. Such dyes can offer good detection limits based on brightness, but typically have broad emission bands and suffer from timedependent photobleaching. Inorganic nanoparticles such as quantum dots and upconversion nanoparticles are photo-stable over prolonged exposure to excitation radiation and tend to offer narrow emission bands, providing greater opportunity for multi-wavelength multiplexing. Importantly, in contrast to molecular dyes, nanoparticles offer substantial surface area and can serve as platforms to carry a large number of conjugated molecules. The surface chemistry of inorganic nanoparticles offers both challenges and opportunities for control of solubility and functionality for selective molecular interactions by assembly of coatings through coordination chemistry. This report reviews advances in the compositional design and methods of conjugation of inorganic quantum dots and upconversion nanoparticles, and the assembly of combinations of nanoparticles to achieve energy exchange. The interest is exploration of configurations where the modified nanoparticles can be immobilized to solid substrates for the development of bioassays and biosensors that operate by resonance energy transfer (RET).


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Introduction

Resonance energy transfer (RET) is a process where energy transfer occurs between a donor and acceptor via a dipolar coupling and without photon emission when they are in sufficiently close proximity. The efficiency of this process is dependent on the distance, degree of spatial alignment of dipoles, and the overlap of the emission and excitation bands of the donor and acceptor.1-4 The efficiency of RET can be used for the detection of binding events that bring a donor and acceptor into close proximity, and has been implemented to determine ensemble and singlemolecule interactions of nucleic acids and proteins. The use of nanoparticle platforms that can serve as donors has provided opportunity for determination of a wide variety of molecular interactions with surface-immobilized biomolecules, with potential for the simultaneous detection of multiple targets by inclusion of multiple acceptors in the detection strategy.5

Quantum Dot – to - Dye energy transfer

Quantum Dots as donors in Fluorescence Resonance Energy Transfer (FRET) Several of the unique optical properties of quantum dots (QDs) make them excellent donors in FRET. These properties include: (i) broad absorption spectra with high molar extinction coefficient; (ii) narrow and size-tunable photoluminescence (PL) spectra with high quantum yield (QY), QY of aqueous QDs in the range of 0.2 to 0.7 depending on surface coating6; and (iii) large accessible surface area for conjugation. A discussion of the details of how these unique optical properties of


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QDs influence the efficiency of energy transfer begins with consideration of the Förster formalism:

aR06 E= 6 6 = aR0 + r

a 6  r  a +   R0 

(1)

In equation 1, E is FRET efficiency, a is the total number of acceptors that are placed equidistantly, r, from the same donor and R0 is the Förster distance, which is a characteristic of a given donor-acceptor FRET pair and is given by Equation 2:

(

Ro6 = 8.79 × 10 −28 mol × n −4κ 2 Φ D J

)

(2)

The Förster distance, Eq. (2), is characteristic of a specific donor-acceptor pair, and depends on factors including the refractive index of the surrounding medium, n, the donor quantum yield, ΦD, the relative orientation between donor emission and acceptor absorption dipoles, and the degree of spectral resonance between the two species. These latter two parameters are described by the orientation factor, κ2, and spectral overlap integral, J, respectively.

∫F J=

D

(λ )ε A (λ )λ4 dλ

∫F

D

(λ )dλ


(3)

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The spectral overlap integral, Eq. (3), is a function of the fluorescence intensity of the donor, FD, and molar absorptivity of acceptor, εA, as a function of wavelength, λ, normalized against the total donor emission. The broad absorption spectra of QDs as compared to molecular fluorophores allows for selection of an excitation wavelength where the direct excitation of an acceptor is minimized.3 This ensures that the majority of the dye acceptors are in the ground state at the excitation wavelength.1, 3 QDs also have high molar extinction coefficient, which becomes stronger as the excitation is moved to progressively shorter wavelengths away from the absorption/excitation spectrum of the acceptor.2 The large “effective” Stokes shift associated with QDs in combination with strong absorption spectra of QDs ensures efficient excitation of the QD donor. The broad absorption spectra of QDs are also useful for the development of multiplexed QD-FRET bioassays, where multiple colors of QDs can be concurrently and efficiently excited with a single excitation source.2 The narrow, symmetric and tunable (via size or composition) PL spectra of QDs can be adjusted for control of the spectral overlap integral in order to maximize the efficiency of energy transfer without significantly introducing crosstalk between the QD PL and acceptor emission.3 The near Gaussian PL profile of QDs greatly facilitates deconvolution of QD PL and acceptor emission from a composite PL spectrum.1 The relatively high QY of QDs is also useful and must be considered in the context of the linear dependency of the sixth power of the Förster distance on the donor QY.1 From the standpoint of multiplexed QD-FRET bioassays, the narrow and symmetric PL spectra of QDs allows for an integration of a greater number of


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color channels within a given spectral window as compared to molecular fluorophores.1 The large surface area afforded by QDs serves as a scaffold to allow multiple acceptors, a, to be concurrently arrayed around an exciton donor in a centrosymmetric configuration, which improves the efficiency of energy transfer.3 This enables acceptors to be FRET paired with a given QD donor despite exhibiting a weak spectral overlap, owing to the additive channels of energy transfer offered by multiple acceptors.3 Additionally, arraying of multiple binding sites each with an acceptor around a single QD donor broadens the dynamic range of QD-FRET bioassays by extending the quantity of binding chemistry that is available before saturation is reached.3 The aforementioned discussion assumes that the Förster formalism is applicable to QDs. Theoretical and experimental studies have confirmed that the Förster dipoledipole interaction mechanism applies to QD-dye (donor-acceptor) FRET pairs in case of direct band gap QDs (e.g. CdSe).7 The inverse 6th power dependence of energy transfer efficiency on the center-to-center donor-acceptor separation distance has been confirmed experimentally provided that the donor-acceptor separation distance is measured from the center of a QD, although there are also instances where separation distance could be considered from the QD surface.8-10 FRET efficiencies were found to scale with the value of spectral overlap integral, the number of acceptors interacting with central QD and the donor-acceptor separation distance that was imposed by the dimensions of a QD.8 The limitation of


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approximating a QD as a point dipole is that it places a threshold on the minimum donor-acceptor separation distance, which is determined by the radius of a QD and its surface coatings.3 However, this limitation to some extent is mitigated by the ability to array multiple acceptors around a single QD donor.1 It is also important to note that the majority of studies that use QD-dye FRET pairs utilize a value of κ2 = 2/3 in the calculation of energy transfer, which is valid provided that the transition dipoles of donor and acceptor are dynamic and random in terms of orientation.3 For molecular fluorophores, free rotational motion around single bonds fulfills this condition despite having a fixed emission dipole orientation. In contrast, CdSe QDs have been reported to have a degenerate transition dipole that is oriented isotropically in two dimensions.1 This implies that the assumption of random orientation of transition dipole is not strictly valid for QDs. Nonetheless, a value of κ2 = 2/3 is a useful approximation for QD-dye FRET pairs given that multiple acceptors are arrayed around a central QD over a distribution of positions, where the acceptors typically have random and dynamic orientation of transition dipoles with respect to the QD transition dipole and the QD donor has partially random orientation of transition dipole.3 QDs can serve as FRET donors for development of solution-phase bioassays, and CdSe/ZnS QDs have been used to develop multiplexed assay strategies for the detection of nucleic acids. Typically, QDs were conjugated to multiple probe oligonucleotides and FRET-sensitized emission from molecular dyes that were associated with complementary target was determined as a quantitative measure of hybridization. Spectrally resolved simultaneous detection of multiple target


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sequences using ensemble measurements and a single excitation wavelength was possible. Such FRET methodology would have greater practical impact for assay development if the QDs could be physically immobilized on solid substrates to develop solid-phase bioassays, and potentially to develop reversible and reusable biosensors.

Solid substrates as platforms for localization of decorated QDs

Our group has investigated three different types of material for use as physical supports for decorated nanoparticles in the development of solid-phase QD-FRET nucleic acid hybridization bioassays. These physical supports include glass and fused silica substrates in the form of optical fibers11, 12, spherical beads13 and planar slides14, 15, and also plastic microtiter plates16 and paper substrates17-20. Each of these solid substrates offers unique advantages and capabilities for the development of QD-FRET nucleic acid hybridization bioassays. The popularity of glass and fused silica as substrates is due to the physical robustness, optical transparency and low autofluorescence of such materials, in combination with the simplicity and yield of surface derivatization using commercially available silane coupling agents.21 As examples, optodes that make use of total internal reflection (TIR) at the sensing region are advantageous for determination of target compounds at a distance.22 The exponential decay field associated with an evanescent wave during TIR confines the optical detection zone to hundreds of nanometers from the surface of an optical fiber, which allows


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surface-selective interrogation that can accommodate nanoparticles.21 Glass surfaces offer stable surface chemistry and good electrical insulating properties suitable for electrokinetically driven fluid flow within microfluidic channels, which offers advantages such as small sample volume, higher sensitivity, improved kinetics for binding interactions such as hybridization, and regeneration of selective chemistry for multiple cycles of use.23, 24

In contrast with glass and fused silica substrates, plastic microtiter plates are commonly used in clinical settings for high throughput bioassays.21 Instrumentation such as microtiter plate readers that are used to interrogate microtiter plates are widely available in research and clinical laboratories, and such optical readers can detect FRET emission. Microtiter plates modified with a variety of functional groups offer opportunity for nanoparticle immobilization, with one example being the introduction of functional groups on polystyrene surfaces.25 In addition, the automation provided by robotic sample handling makes microtiter plates appealing for routine analysis.

Paper-based platforms are attractive for point-of-care and point-of-need diagnostic applications. Several of the attributes of paper substrates are aligned well with the ASSURED (affordable, sensitive, specific, user-friendly, rapid and robust, equipmentfree and deliverable to end users) criteria that have been outlined by the World Health Organization for implementation of diagnostic technologies in the developing world and in resource-limited settings.26 Paper is a low-cost substrate based on


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polysaccharides, and is commercially available with a variety of different physical properties such as pore size and flow rates. Paper substrates can be easily patterned using wax printing to introduce hydrophobic barriers to guide fluid flow and to fabricate paper-based analytical devices (PADs)27, with capillary action that can drive fluid flow. Methods for chemical derivatization of cellulosic fibers of paper substrates are well established, and can be adapted to make use of methods that were initially intended to achieve immobilization on glass and fused silica. Additionally, the three dimensional nature of a paper substrate provides improved cross section of capture (cf. planar glass slide) to enable optical excitation and imaging using a handheld lamp and a smartphone camera19, respectively, and paper can be incinerated to eradicate biohazards27. Another significant advantage of a paper matrix for the development of solid-phase QD-FRET bioassays is the enhancement of FRET efficiency that has been reported with hydrated28 and dry19 paper formats.

Section 2 expands of chemistries used for immobilization of nanoparticles onto solid substrates. It is instructive to first consider modification of QD surfaces more generally, and Section 1 continues with a review of methods that are used to control aspects such as QD solubility, charge and conjugation.

Decorating Quantum Dots


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Quantum dot surface chemistry QDs with high crystallinity, relatively high QY, monodispersity and narrow PL spectra are typically synthesized via pyrolysis of organometallic precursors at high temperatures in organic solvents. As a result, the surface of a QD is capped with hydrophobic moieties, which is a configuration not suitable for the direct application of as-synthesized QDs in aqueous biological environments. It is necessary to modify the surface chemistry of native QDs to impart water solubility. This is typically done by the addition of a suitable coating to the surface of a QD. The two commonly reported strategies to make QDs water-soluble are: surface ligand exchange using thiol or imidazole functionalized small molecules as illustrated in Figure 1a (i), and polymer encapsulation of QDs as shown in Figure 1a (ii).29 Important considerations for the choice of a method to achieve aqueous solubility of QDs include: (1) a high affinity by the new ligand for the surface of a QD while maintaining long-term colloidal stability across a broad range of pH and ionic strength conditions, (2) subsequent capacity for bioconjugation, (3) maintaining a compact and small hydrodynamic size of the decorated QD to facilitate the distance requirements FRET, (4) preservation of optical properties of QDs and functionality of the attached biomolecule(s), (5) minimization of non-specific adsorption, (6) lowcost and commercial availability of the QD surface ligand or amenability to a large scale synthesis of the QD ligand at a low cost, and (7) low toxicity for cellular and in vivo studies.30, 31


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Figure 1. (a) Illustration of two general methods to confer aqueous solubility to QDs. (i) Ligand exchange of native hydrophobic ligands of QDs by hydrophilic ligands that coordinate to the surface of a QD, and (ii) polymer encapsulation of QDs where hydrophobic moieties of an amphiphilic polymer intercalate with the native


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hydrophobic ligands of QDs. (b) Chemical structures of selected QD surface ligands. (i) Schematic of various modules associated with dihydrolipoic acid-polyethylene glycol (DHLA-PEG) ligand derivatives with various distal functional groups; (ii) monodentate thioalkyl acid ligands; (iii) bidentate DHLA ligand; (iv) bidentate DHLA-PEG ligand with different functional groups (R) at the distal end and (v) tetradentate DHLA-PEG ligand with a methoxy group at the distal end. Panel (a) adapted with permission from reference30, Copyright 2011 American Chemical Society. Panel (b) adapted with permission from reference32, Copyright 2014 Elsevier.

In the case of the ligand exchange method, the native hydrophobic surface capping ligands (e.g. trioctylphosphine (TOP), trioctylphosphine oxide (TOPO) and longchain alkylamines) of QDs are replaced with hydrophilic organic molecules that are typically heterobifunctional.33 The ligand exchange reaction is driven by the high affinity of hydrophilic ligands that self-assemble via coordination as a monolayer on the surface of QDs. This process of exchange is also driven by mass action, where hydrophobic QDs are incubated with a large molar excess of the desired ligand molecule to thermodynamically and kinetically facilitate the cap exchange.31 QD surface ligands are comprised of at least two components: proximal anchoring group(s), and hydrophilic group(s). The anchoring groups are responsible for interacting with the surface of QDs, while the hydrophilic groups impart solubility in aqueous media. For many ligands, a distal functional group is also introduced to provide a site for bioconjugation.


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Monodentate

thioalkyl

acids,

such

as

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mercaptoacetic

acid

(MAA),

mercaptopropionic acid (MPA), and mercaptoundecanoic acid (MUA) shown in Figure1b (ii), are among the commonly used commercial reagents that can confer aqueous solubility to QDs. The proximal thiolate group coordinates strongly with ions such as Cd2+ and Zn2+ present on the surface of QDs (dependent on selected composition), while the distal carboxylic group when subjected to ionization under sufficiently basic conditions imparts water solubility and provides colloidal stability to QDs by electrostatic repulsion.34 While monothiol based coatings tend to be compact in nature, they suffer from a lack of long-term stability due to high lability of monothiol ligands at the QD surface.35 A significant shortcoming of QDs capped with thioalkyl acids is their propensity to undergo aggregation in high ionic strength or low pH solutions owing to the charge neutralization of carboxylate groups. In comparison to monothiol ligands, aqueous QDs capped with bidentate thiol ligands (e.g. dihydrolipoic acid (DHLA) shown in Figure 1b (iii)) offer prolonged shelf life that ranges from several months to a year.36 This is due to a cooperative chelate effect of the dithiol functionality as an anchoring group on the same DHLA molecule. The colloidal stability in case of DHLA-capped QDs is also governed by electrostatic repulsion between negatively charged carboxylate groups, hence DHLA-capped QDs are also prone to aggregation depending on pH and ionic strength conditions.36 To circumvent this limitation, various modular designs of DHLA-ligand derivatives that include DHLA appended with polyethylene glycol (PEG) or zwitterionic functionalities have been used (Figure 1b (i) and 1b (iv)), where colloidal stability in aqueous conditions is imparted by the interaction of hydrophilic moieties (e.g. PEG


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or zwitterionic functional groups) with the solvent instead of reliance on the deprotonation of distal carboxyl group.35, 36 As a result, these coatings offer colloidal stability across a wider range of pH and ionic strength conditions, minimal nonspecific adsorption and improved biocompatibility when compared with using only the DHLA as an anchor.35, 36 Derivatives of DHLA-PEG ligands containing a distal functional group (e.g. primary amine, carboxyl, methoxy and biotin) have also been reported, and can be used for bioconjugation and covalent modifications (Figure 1b (iv).37 In addition to the bidentate DHLA ligands, multidentate (tetradentate) thiol ligands comprised of two DHLA anchoring groups appended to either a PEG chain or a zwitterionic functionality can be used (Figure 1b (v)), which further augments colloidal stability of aqueous dispersions of QDs in a variety of extreme conditions (pH range 1.1 to 13.9 and 2 M NaCl).38 In contrast with zwitterionic ligands, the PEGbased ligands exhibit larger hydrodynamic size and can potentially provide a barrier against metal-affinity driven self-assembly of biomolecules via a histidine (His) moiety (vide infra).39 An inherent shortcoming of thiols as anchoring groups for QD functionalization is that thiols are known to serve as traps for holes (electron-hole pair caused by optical excitation)40, which can greatly reduce the QY of QDs upon ligand exchange. The holes trapped by thiol ligands on the surface of QDs can also promote oxidation of thiol ligands to disulfides, which augment the photochemical instability of aqueous QDs capped with thiol ligands.41

Numerous improvements of

ligand exchange methods have been reported to ameliorate this loss in the QY of QDs and to augment the colloidal stability of QDs capped with thiol-based ligand. These improvements include: (1) the use of an organic base to increase the reactivity of the thiol


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anchoring group of MPA ligand with the ZnS shell of a QD34, (2) metalation of DHLA ligand with zinc to produce tetrahedrally coordinated (DHLA)2Zn2 complex which preserves QD shell structure, i.e., from etching during the cap exchange process42, (3) using UV irradiation for photochemical transformation of lipoic acid (LA)-modified ligands to produce heterogenerous population of LA derivatives, where higher order oligomers (dimers, trimers and tetramers) exhibit faster and stronger coordination for ZnS-overcoated QDs to promote cap exchange for aqueous solubility43,

44

, (4)

ultraefficient cap exchange method requiring ligand-to-QD molar ratio (LQMR) of as low as 500 (20-200 fold less than most methods), involving the use of tris(2carboxylethyl)phosphine for reduction of LA to DHLA and NaOH for deprotonation of the thiol groups. The low LQMR was beneficial in retaining the original fluorescence of hydrophobic QDs (>90%) by preventing the QD shell from etching during the cap exchange process.45 In addition, ligand exchange of hydrophobic ligands with hydrophilic ligands can also potentially result in unpassivated sites, which promote quenching of QD PL.30 In addition to thiols, dithiocarbamate ligands derived from amino acids have also been reported to achieve aqueous solubility of QDs.46 Interestingly, in contrast with bidentate DHLA ligands, QDs capped with dithiocarbamates exhibit high QY that are similar to values determined for organic QDs.46 Polymeric encapsulation of QDs has been reported using two different approaches. Traditionally, this has been accomplished by means of amphiphilic polymers where pendant alkyl chains intercalate via hydrophobic interactions with native hydrophobic ligands of QDs (e.g. TOP or TOPO).47 The hydrophilic component of


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amphiphilic polymer usually incorporates functional groups (e.g. carboxylic groups or amines groups) and/or PEG chains to support subsequent chemical modification and to promote water solubility, respectively. Recently, the hydrophobic portion of the amphiphilic polymer has also been used to incorporate functional groups for subsequent immobilization of molecules.48 Polymer encapsulated QDs yield robust structures with improved optical properties in addition to long-term stability under a variety of conditions as compared to ligand coated QDs. However, polymer encapsulation of QDs typically results in a significant increase in the hydrodynamic size of QDs (> 20 nm)47, which is detrimental for FRET applications owing to the strong dependency of energy transfer efficiency on the donor–acceptor center-tocenter separation distance. More recently, polymers have been developed which exhibit coordinating groups such as dithiol49 or pyridine50. These groups interact directly with the surface of a QD by multidentate ligand exchange interactions, which serves to reduce the hydrodynamic size of QDs. Additional functional groups, such as primary amines and carboxylic groups, are also incorporated for further bioconjugation. In addition to polymer encapsulation of QDs, silica shell encapsulation of QDs has also been reported where QDs are first ligand exchanged with a silane coupling agent, such as 3-mercaptopropyltrimethoxysilane, followed by shell growth that involves hydrolysis and condensation reactions.51 A more detailed discussion on surface coatings of QDs for aqueous solubility can be found elsewhere.52


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Conjugation of biomolecules to QDs

QDs are made “functional” for use in bioassays and biosensing by conjugation of biomolecules to the QD surface. The strategies for the preparation of QDbioconjugates can be broadly classified into three categories: (1) physisorption, (2) covalent interaction and (3) coordination linkage.31,

32

Physisorption of

biomolecules on the surface of QDs relies on electrostatic, polar or hydrophobic interaction, and involves a spontaneous association of biomolecules with the surface coating of a QD.1 In the case of covalent interaction, a new bond is formed between the functional group of a biomolecule and a functional group that is associated with the surface coating of a QD. Coordination linkage is based on dative interaction and involves spontaneous self-assembly of a functional group of a biomolecule on the surface of inorganic shell of a QD or the surface coating of a QD.32 Important considerations for bioconjugation chemistry for the preparation of QDbioconjugates include: retention of optical properties of QDs while maintaining colloidal stability; preservation of the activity of a biomolecule; control of biomolecule orientation; the stoichiometry of biomolecule conjugation; stability of the QD-biomolecule conjugate; mild conditions for the preparation of QDbioconjugates such that the reaction conditions do not adversely affect the biomolecule activity, and use of a low concentrations of reactant(s) while maintaining high yield of biomolecule coupling. Covalent methods for the preparation of QD-bioconjugates are primarily derived from chemistries used for protein labeling and make use of functional groups such


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as primary amines, carboxyls and thiols.30 These chemistries require additional reagents for activation such as carbodiimide, succinimidyl ester, maleimide or pyridyl disulfide.

53, 54

The popularity of these coupling methods arises from

ubiquitous display of carboxyl and amine groups by proteins and that these functional groups can be easily incorporated into surface coatings of QDs. The commercial availability of aqueous QDs includes nanoparticles with these functional groups. However, given the concurrent presence of carboxylic groups and amine groups on proteins, cross-linking between proteins and formation of QD-protein-QD constructs are not uncommon from such reactions. This not only leads to mixed avidity but also results in an uncontrolled variation in the valence of resultant QDprotein bioconjugates, including limitations in the control of biomolecule orientation.32 It should be noted that cross-linking is a less severe issue for oligonucleotide bioconjugation to QDs as oligonucleotides can be easily monofunctionalized with a reactive group during solid-phase synthesis. In addition to these chemistries, chemoselective and bioorthogonal methods for the preparation of QD-bioconjugates

that

include

strain-promoted

azide-alkyne

cycloaddition,

hydrazone ligation, oxime ligation, alkene-tetrazine ligation and Staudinger ligation have also been reported.30 The implementation of these chemistries for the preparation of QD-bioconjugates is described in detail elsewhere.53 Affinity binding based on the avidin-biotin interaction is another coupling chemistry that has been used for the preparation of QD-bioconjugates. The popularity of avidin-biotin chemistry originates from its high association constant (Ka) of ca. 1015 M-1, which is one of the strongest known non-covalent interactions.32 The binding is


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stable across a wide range of pH and ionic strength conditions. Additionally, commercial availability of streptavidin (SAv)-coated QDs (SAv is a tetrameric homologue of avidin that is isolated from Streptomyces avidinii) in combination with a wide range of commercial kits and reagents for biotinylation makes this bioconjugation chemistry readily accessible. The biotin functionality can be added to peptides or oligonucleotides during solid-phase synthesis. Some of the shortcomings of this coupling chemistry include inability to control the orientation of biomolecule attachment. In the case of biomolecules that carry multiple biotin sites, cross-linking resulting in a formation of heterogeneous population of QDbiomolecule-QD assemblies can be a disadvantage.32 Such cross-linking is not a major concern for an oligonucleotide strand that is mono-functionalized with a biotin functionality. Biomolecules displaying a thiol functional group or a polyhistidine tag can be selfassembled on the surface of CdSe/ZnS QDs by dative interaction. In the case of a thiol functionality, the interaction is with the Zn2+ ions or sulfur moieties present on the QD surface.34, 53, 54 The nature of the QD surface capping ligand responsible for aqueous dispersion of QDs is a crucial factor in governing this type of interaction, as the

inorganic

surface

of

QD

must

be

accessible.

Given

the

dynamic

association/dissociation of monothiol interaction with QD surfaces, the longevity of the resulting QD-bioconjugates under non-equilibrium conditions is a significant concern. The stability of QD-bioconjugates prepared via the dative interaction of a thiol functionality can be improved by using biomolecules that exhibit multiple thiols (e.g. dithiol), which improves the stability of linkage by a cooperative


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chelation effect.54 However, spontaneous oxidation of a dithiol group to a disulfide group can impede the stability of resulting bioconjugates. It is also important to ensure that the thiols that coordinate to the QD surface are not integral to protein structure and function. One of the most robust strategies for the preparation of QD-bioconjugates is based on polyhistidine-metal-affinity interaction, which refers to the ability of histidine residues to coordinate with transition metals (e.g. Co2+, Cu2+, Ni2+ and Zn2+) via the imidazole side group that serves as a Lewis base.32 The strong affinity of binding between a polyhistidine tag and the metal ions can be used for the preparation of self-assembled QD-bioconjugates. Polyhistidine-metal-affinity driven self-assembly of biomolecules to the QD surface has been reported using three different methods: (1) direct coordination of polyhistidine tag to the inorganic surface of QDs; (2) mutual chelation of Ni2+ ions by carboxylic groups of polymer encapsulated QDs and a polyhistidine tag associated with a biomolecule; and (3) modification of the QD surface with nickel-nitrilotriacetic acid (Ni-NTA) groups.32 Each of these approaches extends the applicability of polyhistidine-metal-affinity interaction for self-assembly of biomolecules to different QD surface chemistries. Preparation of QDbioconjugates using a polyhistidine motif offers a number of advantages. It is a bioorthogonal means of bioconjugation, as a polyhistidine moiety does not exist naturally in proteins.32 The single attachment point provides some control of biomolecule orientation and avoids undesired cross-linking reactions.32 Owing to the high affinity of binding, there is improved control of the average stoichiometry of the QD-bioconjugate, ameliorating the need for additional purification steps.32


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This method does not compete with hydrolysis, hence facilitates rapid bioconjugation32 with self-assembly reaching equilibrium within ca. 100-200 s, with dissociation constants (Kd) in the range of 10-10-10-7 M.39 The relatively small size of a polyhistidine motif is also beneficial for the retention of native function of a protein.32 For the case of amphiphilic polymer encapsulated QDs, the inorganic surface of the QD is inaccessible.55 As a result, the direct coordination of polyhistidine motif to the inorganic shell of a QD is not possible for the preparation of QD-bioconjugates. However, when the amphiphilic polymer exhibits carboxylic groups, the mutual chelation of Ni2+ ions by surface carboxylic groups and a polyhistidine motif tag can be used for self-assembly of biomolecules.56 In this bioconjugation strategy, the role of surface carboxylic groups is analogous to NTA, which when associated with Ni2+ ions is known to coordinate strongly with the polyhistidine motif.32 Alternatively, in the absence of surface carboxylic groups for polymer encapsulated QDs, the surface of QDs can be modified with NTA, which when supplemented with Ni2+ ions, can be used for polyhistidine mediated self-assembly of biomolecules.57 For a greater indepth discussion on bioconjugation using the polyhistidine motif, the reader is referred to a review article by Blanco-Canosa and coworkers.32 In addition, bioconjugation of QDs has been extensively addressed in recent reviews and interested readers are referred to these review articles.53, 54

Control of adsorption of oligonucleotides on QDs


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For the development of QD-FRET bioassays, QD surface coatings based on thioalkyl acids (e.g. MAA, MPA and DHLA) are attractive given their compact ligand size, ease of preparation of aqueous QDs and subsequent capacity for bioconjugation.21 As thioalkyl acid ligands were among the first reported QD surface coatings to impart aqueous solubility to QDs33, initial efforts by our group to develop QD-FRET nucleic acid hybridization bioassays also investigated QDs coated with MAA and MPA ligands.58, 59 It was found that the analytical performance of QD-FRET nucleic acid hybridization bioassays was greatly affected by the adsorption of oligonucleotides on MAA/MPA-capped QDs.58, 59 Given that non-specific adsorption is undesired in bioassay development, Algar and Krull investigated the origin of adsorption of oligonucleotides on MPA- and MAA-QDs using FRET as a transduction method.60, 61 The adsorption was investigated using Cy3-labeled oligonucleotide sequences and green-emitting CdSe/ZnS QDs as donors. FRET is a useful tool to study adsorption of oligonucleotides on QDs. This method only requires oligonucleotides to be labeled with a suitable fluorescent acceptor dye that is paired with a QD donor, and the strong dependency of FRET efficiency on the donor-acceptor separation distance (Equation 1) allows differentiation between adsorbed and freely diffusing oligonucleotides with high sensitivity and minimal perturbation of the system.61 The caveats to using FRET to interrogate oligonucleotide adsorption include: that this method is unable to differentiate between changes in FRET efficiency that originate from changes in the stoichiometry of adsorbed oligonucleotides on the QD surface and dynamic changes in the ‘tightness’ of adsorbed oligonucleotides on the QD surface; and that the FRET efficiency response can potentially saturate prior to the


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saturation of a QD surface with adsorbed oligonucleotides.61 Nonetheless, FRET is a useful tool to study adsorption of oligonucleotides on the QD surface with the expectation that conditions that suppress the number of oligonucleotides adsorbed on the QD surface will also reduce the ‘tightness’ of oligonucleotide bound on the QD surface.61 Thus, both of these effects will work in concert to decrease FRET efficiency.

Figure 2. (a) Adsorption behavior of oligonucleotides on MPA-QDs. (i) Dependence of the pH of buffer on the adsorption of Cy3-labeled mixed base sequence on MPAQDs. (ii) Changes in the Cy3/QD PL ratio (FRET ratio) for the adsorption of increasing molar ratio of Cy3-dA20 sequence on MPA-QDs at pH 7.4. The inset shows changes in the FRET ratio for the adsorption of Cy3-dC20, Cy3-dA20, Cy3-d(TGGG)5 and Cy3-dT20 on MPA-QDs at pH 7.4. (iii) Changes in the adsorption of Cy3-labeled


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mixed base sequence on MPA-QDs as a function of increasing NaCl concentration at pH 8.5 and pH 9.3. (b) (i) Chemical structure of a glutathione (GSH) ligand. Changes in the FRET ratio for solution-phase hybridization of oligonucleotide probe modified GSH-QDs with increasing number of (ii) 3' Cy3-labeled fully-complementary (FC) proximal target and (iii) 5' Cy3-labeled FC distal target at pH 9.2. Note the difference in the sensitivity response of (ii) and (iii). In (a), panels (i), (ii) and (iii) adapted with permission from reference61. Copyright 2011 Elsevier. In (b), panels (ii) and (iii) adapted with permission from reference19. Copyright 2014 American Chemical Society.

As presented in Figure 2a (i), experiments involving titration of MPA-QDs (as donors) with increasing numbers of acceptor dye-labeled oligonucleotides showed that adsorption was strongest at acidic pH and decreased with increase in solution pH.61 In the case of CdSe/ZnS QDs coated with MAA, adsorption was found to be more than 10-fold greater at pH 4.8 than at pH 9.5.60 The pH dependent adsorption experiments exhibited a profile analogous to an acid-base titration curve, showing that the pKa of QD-bound MPA ligands was ca. 7.8.61 In contrast, the pKa of MPA in bulk solution has been reported to be 4.3.62 This elevation of pKa of MPA at the QD interface is consistent with a previously published study that has reported an elevation of pKa of carboxylic group at the nanoparticle interface.63 Adsorption experiments and competitive binding experiments showed that different nucleobases exhibit varying degrees of tendency to adsorb on MPA-QDs. As shown in Figure 2a (ii), the order for adsorption affinity of different nucleobases on MPA-


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QDs was found to be dC > dA ≥ dG >> dT.61 The adsorption of oligonucleotides on MAA- or MPA-QDs was found to be driven by hydrogen-bonding, where neutral carboxylic groups of thioalkyl acid ligands interacted favorably with nucleobases.60, 61

Support for the hydrogen-bonding mechanism in mediating adsorption of

oligonucleotides on MAA- and MPA-QDs was provided by experiments that showed that the addition of a hydrogen bond disrupter (i.e., formamide) suppressed adsorption of oligonucleotides on the QD surface.60 In addition, the extent of adsorption of double-stranded oligonucleotides was found to be significantly less than single-stranded oligonucleotides.60 The extent of oligonucleotide adsorption on MPA-QDs was also found to be dependent on the ionic strength of solution as shown in Figure 2a (iii). A log-linear relationship between adsorption and ionic strength was observed at pH 8.5, while at pH 9.3, where ionization of MPA ligands on QDs is expected to be complete, adsorption was found to be negligible up to 100 mM NaCl concentration.61

Adsorption also impacted the conformation of oligonucleotide probes conjugated to the QD surface, and influenced hybridization kinetics and stability of duplex formation at the QD interface.60 Hybridization of QD-probe conjugates with a fullycomplementary (FC) target that was labeled with an acceptor dye at either the distal (5') or proximal (3') terminus offered similar FRET efficiencies, which were also found to be independent of the linker length used for QD bioconjugation.60 This suggested that at low density, oligonucleotides laid along the surface of QDs instead of orienting upright from the QD surface. Increasing the number of conjugated


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probes on the QD surface caused immobilized probes to orient upright from the QD surface when subjected to hybridization with complementary targets.60 The rates of hybridization of complementary oligonucleotides to the QD-probe conjugates were found to scale proportionally to the rate of adsorption of non-complementary oligonucleotide.60 In addition, adsorption impacted the thermodynamic stability of DNA duplex at the QD interface.60 For a FC target, the hybrids at the QD interface exhibited sharper melt curve transition and a decrease in melt temperature (Tm) by 2 °C as compared to the bulk solution hybridization.60 In contrast, hybrids containing mismatches exhibited an increase in Tm and broadening of melt curve transition as compared to bulk solution counterparts.60 Melt curves obtained under the conditions in which adsorptive interactions became less favorable showed transitions which closely resembled bulk solution hybridization.60 These effects were attributed to the competition between probe-target interaction and adsorption of oligonucleotides on the QD surface. The impact of adsorption interactions on the stability of hybrid was found to be temperature dependent.60 Adsorptive interactions served to stabilize duplex formation below the Tm, while they facilitated duplex denaturation above the Tm, resulting in sharper melt curve transitions.60

The insights into the mechanism of oligonucleotide adsorption on thioalkyl acid capped QDs were seminal in shaping further development and improving the analytical performance of QD-FRET nucleic acid hybridization bioassays. In recent studies, we have used glutathione capped QDs (GSH-QDs) for the preparation of water soluble QDs and subsequently used these QDs for the assembly of QD-FRET


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nucleic acid hybridization bioassays.17-20 As shown in Figure 2b (i), GSH is a tripeptide exhibiting a thiol group, a primary amine group and two carboxylic groups. The proximal thiol and primary amine groups coordinate to the ZnS shell of CdSeS/ZnS (core/shell) QDs while the distal carboxylic groups under sufficiently basic conditions provide colloidal stability in aqueous media. Given that the distal carboxylate groups are responsible for aqueous solubility of GSH-QDs (pH and ionic strength dependent), it was anticipated that the interaction of nucleobases with neutral carboxylic groups of GSH via hydrogen bonding mechanism could potentially contribute to the non-specific adsorption of oligonucleotides. The adsorption of oligonucleotides on MAA- and MPA-QDs was suppressed under basic conditions (pH > 9).61 By conducting the hybridization assays at pH 9.2 using GSHQDs that were modified with single-stranded oligonucleotides as probes, the nonspecific adsorption of oligonucleotides on GSH-QDs was sufficiently minimized that no surface passivation of the QD surface was required.18, 20 It is likely that the partial zwitterionic character of GSH also contributed to the suppression of oligonucleotide adsorption on GSH-QDs.64 Hybridization bioassays conducted with a FC target that was labeled with an acceptor dye at the proximal (3') or distal (5') terminus showed significantly different assay sensitivities as can be seen in Figure 2b (ii) and (iii), respectively, which was in contrast with MPA-QDs. The FC target labeled at the proximal end showed ca. 100-fold higher assay sensitivity as compared to the FC target that was labeled at the distal end.19 This provided further confirmation that the oligonucleotide probes were not adsorbed on the surface of GSH-QDs. It is also interesting to note that under the conditions where adsorption was favored on


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MPA-QDs (pH 7.4), the ratio of the FRET-sensitized acceptor dye PL to QD donor PL (FRET ratio) saturated at an acceptor to QD ratio of 4 to 1, as can be seen in Figure 2a (ii). In contrast, hybridization of GSH-QDs modified with oligonucleotide probes with a dye-labeled FC target offered a saturation of FRET ratio response at an acceptor to QD ratio of 40 to 1 for both the proximal (Figure 2b (ii)) and distal (Figure 2b (iii)) labeled targets. This suggests that the suppression of oligonucleotide adsorption on a QD surface also impacts the loading capacity of oligonucleotides probes on the QD surface, where adsorption excludes some of the surface area of a QD from oligonucleotide conjugation. The use of thiol ligands for the development of homogenous QD-FRET nucleic acid hybridization assays has also been reported by other groups65-67. In these studies, various thiol ligands were appended to a PEG moiety to suppress non-specific adsorption of oligonucleotides and to promote colloidal stability of QDs in aqueous medium.

QD-FRET assay using intrinsically labeled probes

Transduction of nucleic acid hybridization by a QD-FRET method relies on a change in the positioning of an acceptor dye with respect to the donor QD surface upon target hybridization. This results in a modulation of FRET efficiency response, which serves as an analytical signal.68 From the standpoint of QD-FRET transduction of nucleic acid hybridization, the proximity between the donor QD surface and the acceptor dye has primarily been accomplished either by directly labeling a target strand with the acceptor dye18, 20, 58, 59 or by introducing a sandwich hybridization


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format that makes use of a labeled reporter strand11,

12, 17, 19.

Although these

approaches are functional, they introduce additional processing steps, potentially increasing the complexity of assays. Additionally, these approaches are strictly limited to in vitro assay configurations (cf. ex vivo or in vivo QD-FRET transduction of nucleic acid hybridization). Given that the detection of unlabeled targets is desirable, our group has recently reported a homogenous assay format for QD-FRET transduction of nucleic acid hybridization that made use of intrinsically labeled oligonucleotide probes for the detection of unlabeled targets.69 The oligonucleotide probe strands were labeled with two adjacent molecules of a derivative of thiazole orange (TO) intercalating dye. The modified probes were then bioconjugated to the surface of green-emitting QDs (QD525) using SAv-biotin for coupling.69 The QDs served as donors for the excitation of TO fluorescent dyes by FRET. In the absence of probe-target duplex formation at the QD interface, the two TO molecules formed an H-aggregate dimer, resulting in quenching of the fluorescence emission of the dye molecules due to excitonic interaction between the two dye molecules.69 Upon hybridization, the H-aggregate dissociated as the dye molecules preferentially intercalated

with

the

double

stranded

DNA

duplex,

resulting

in

restoration/enhancement of the fluorescence emission of the dye molecules. The relative positioning of the dye molecules from the donor QD surface, the distance between the two dye molecules and the attachment location (DNA phosphate backbone or thymine nucleobases) greatly impacted the analytical performance of the assay.69 The hybridization bioassays provided a limit of detection (LOD) of 10 nM (2 pmol) and a dynamic range spanning one order of magnitude, and this


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performance was identical to targets of 34 and 90 nucleobase length.69 The selectivity of the assay was shown by single nucleotide polymorphism (SNP) discrimination. Albeit in its infancy at the current stage, with further development and optimization, the use of intrinsically labeled probes in conjunction with the QDFRET transduction method may potentially allow direct probing of the dynamics of intracellular gene expression levels.

Solution versus solid-phase RET bioassays Advantages of solid-phase bioassays

Nanoparticle (NP)-based RET bioassays may be done with NPs dispersed in solution (solution-phase RET) or immobilized on a solid substrate (solid-phase RET). Solidphase bioassays offer some opportunities beyond the solution-phase methods. Nanoparticle immobilization in solid-phase bioassays eliminates the need to limit the reaction conditions to only those that allow for colloidal stability of nanoparticles. For instance, a high ionic strength condition may be applied to a solid-phase assay to accelerate the reaction between similarly charged nanoparticles and DNA, but the same high ionic strength condition may compromise the stability of solution-phase nanoparticles. Bioconjugation onto a surfaceimmobilized nanoparticle is greatly enhanced by applying a large excess of biomolecules or biorecognition elements, and solid-phase immobilization allows for washing to remove unbound molecules. Washing of the surface may also be advantageous to remove interferences prior to the detection step. While washing of


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NPs that are in solution-phase can be accomplished, the process requires tedious and less efficient purification steps. Moreover, solid-phase bioassays offer multiplexing potential by spatial arraying of probes on the solid-substrate, and potentiate the use of near-field optical techniques such as surface plasmon resonance spectroscopy and photonic crystal enhanced fluorescence.11 In addition to the general advantages of the solid phase bioassays, RET bioassays may particularly benefit from the close proximity of immobilized NPs at an interface. As discussed in section 1.3, the RET efficiency increases with the number of acceptors assembled onto the nanoparticle donor. In a similar fashion, when donors and acceptors are immobilized at a high density on a solid substrate, a single acceptor may accept energy from multiple donors resulting in increased FRET efficiency.20

Immobilization of nanoparticles on solid surfaces

A key step in the assembly of solid-phase RET bioassays is the immobilization of nanoparticle donors on a solid substrate. Our group has investigated the immobilization of nanoparticles on a variety of solid substrates including glass, paper, and fused silica optical fibers.18 The goal was to develop surface chemistries that are facile, that provide robust surface immobilization, and may potentially be applied to a variety of substrates. For instance, thiol, imidazole and amine groups are known to coordinate with the surface of CdSe/ZnS QDs,70 and carboxylate,


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phosphate and amine groups are the most widely used anchoring groups for immobilization of lanthanide-based UCNPs.71 Another strategy was to use selective and high-affinity interactions of biomolecules for nanoparticle immobilization. Early attempts to immobilize QDs made use of monothiol-functionalized fused silica optical fibers as the substrate.18 The lability of binding between thiol and ZnS surface of QDs contributed to instability when using monodentate thiolfunctionalized surface. Efforts then took inspiration from previous work that addressed functionalization of QDs in bulk solution, and multidentate surface ligand exchange (MSLE) was explored to enhance the stability of immobilization on surfaces. The first MSLE embodiment included the functionalization of bidentate DHLA groups (dithiol) on silica optical fibers.18 Next, the bidentate ligand was replaced with a tetradentate thiol ligand to further enhance the robustness and surface density of QD film on glass and silica substrates. 13 Despite the increased coordination by multidentate thiol groups with the ZnS surfaces of QDs, the immobilized QDs tended to dissociate from surfaces as the thiol groups oxidized over time. The search for more stable chemistries then investigated multidentate imidazole-functionalized surfaces for QD immobilization using MSLE. A variety of substrates, including glass, polystyrene microtiter plates and paper substrates were functionalized with multidentate imidazole groups and provided for stable QD immobilization.18 Another strategy for QD immobilization was via high-affinity biomolecular interactions such as DNA hybridization and Streptavidin (SAv)-biotin interactions. As one example, QD immobilization was achieved via DNA hybridization between


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two complementary oligonucleotides, one tethered to glass and the other functionalized on QD surfaces.24 In another example, high-affinity binding between SAv and biotin was used for immobilization of QDs on glass substrates for FRET bioassays of DNA targets. 11, 24 Immobilization of UCNPs on solid substrates can also be achieved. This has usually been reported in the literature as being driven via physical adsorption. We have used streptavidin-coated UCNPs on paper substrates for Luminescence Resonance Energy Transfer (LRET ) DNA hybridization bioassays.72 First, the oleate surface ligands on UCPs were oxidized to render the nanoparticles ligand free and then sodium citrate and streptavidin were sequentially coated on the surface. Although physical adsorption provides a facile strategy for immobilization of UCNPs, weak binding results in instability and desorption of immobilized nanoparticles. Moreover, the low-density coverage of adsorbed nanoparticles obviates any RET signal enhancement due to the interaction between neighboring RET pairs in adjacent UCNPs. Therefore, we developed an strategy that allows for immobilization of UCNPs onto modified paper substrates via covalent chemistry.73 Amine modified UCNPs were prepared by ligand exchange using o-phosphorylethanolamine (PEA) and subsequently immobilized on the aldehyde functionalized glass coverslips. The closely-packed solid-phase UCNPs showed an improvement in assay sensitivity in comparison with both solution-phase bioassays and also less densely packed solidphase bioassays. Improvement in sensitivity would result from greater availability of surface area for selective reaction and potentially also from optical “cross-talk” from nearest-neighbour interactions.73


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QD immobilization in microfluidic channels

Microfluidics is a powerful tool in various applications in biology including bioassays, bioconjugation, drug development and delivery, and tissue engineering. Microfluidic chips present a promising platform for solid-phase RET bioassays with significant advantages over the bioassays done in bulk solution. Such advantages include a reduction of the quantity of sample and reagents required for bioassays and improved speed, sensitivity, resolution and throughput of the bioassays. However, immobilization of NPs by adsorption on the surface of microfluidic chips does not necessarily provide sufficient stability. For instance, the applied electric field in chips that operate using capillary electrophoresis and electro-osmotic flow imposes significant force on the QD-oligonucleotide conjugates immobilized on thiol-functionalized chip surface, and has been seen to cause desorption and migration of nanoparticles.21 One successful strategy to create stable NP films on glass microfluidic channels utilized DNA hybridization.24 The surfaces of the microfluidic channels were covalently coated with single-stranded oligonucleotide that served as a tether. Two different oligonucleotides were conjugated on QDs, one of which was used as an anchor to hybridize to the complementary oligonucleotide tether attached to the glass, and the other remained available to serve as a probe for hybridization with target DNA that could be transported in the microfluidic channel. This strategy allowed for analysis of target DNA using FRET, subsequent NP removal by introduction of a denaturing conditions to dehybridize the tether-anchor system,


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and re-use of the microfluidic chip by recoating with fresh QDs for subsequent cycles of analysis of target oligonucleotides.

Another approach made use of SAv-biotin interaction for QD immobilization on the surface of a glass-PDMS microfluidic chip for FRET bioassays of DNA targets.21 Electroosmotic flow (EOF) was used to deliver SAv-QDs through the microfluidic channel to dynamically immobilize the NPs onto the biotin-functionalized glass substrate. Subsequently, application of voltage was used to deliver biotinfunctionalized DNA probes to decorate the immobilized QDs. The FRET system was then ready for determination of aliquots of Cy3-labeled oligonucleotide targets that were transported by application of voltage. Two types of commercial SAv-QD conjugates were used; SAv molecules were either conjugated directly or through a PEG spacer to the QD surface. The FRET bioassays demonstrated that the QDs which were directly conjugated with SAv provided a significantly higher FRET efficiency, and this is consistent with provision of a closer proximity between QD surfaces and the Cy3 molecule labels that served as acceptors.15 This strategy was extended to explore a two-plex FRET hybridization assay using PL spatial profile of QDs of two different color (QD525 and QD605) as donors and Cy3 and Alexa-647 as acceptors (Figure 3).14


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Figure 3. Schematic representation of multiplexed nucleic acid hybridization bioassays in a hybrid glass/PDMS based microfluidic channel using immobilized QDs as FRET donors. Adapted with permission from reference14. Copyright 2013 Elsevier. Extending microfluidic methods for application to NP decoration

Microfluidics flow has been used for on-chip immobilization of NPs and subsequent probe conjugation on NP surfaces.14, 15 The immobilization of NPs allows treatment with a sequence of reaction and washing solutions, and laminar flow in microfluidic


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environments offers excellent control of reaction conditions at surfaces that achieve reproducibility with high speed. This suggests potential for use of microfluidic chips as a manufacturing platform for NP decoration. Moreover, a particular advantage of using NP decoration in microfluidic channels is the potential to achieve coatings on defined areas of NP so that different molecules can be conjugated to one NP. When NPs are strongly immobilized on a surface, one side of NPs is sterically blocked by the surface. Thus, ligand immobilization may only occur on the solution-facing side of the NPs. Subsequent removal of such NPs from the surface into solution exposes the unmodified sides of the NPs that can be further derivatized. Asymmetrically decorated (Janus) NPs may be produced using solid-phase decoration. Single-phase microfluidics (SPM) based on continuous flow is limited as a platform for NP decoration, as SPM is plagued by cross-contamination and slow mixing due to the laminar nature of the flow. Another challenge for NP decoration using continuous flow microfluidics is the scale-up, as the throughput only linearly increases with the footprint of the device. Thus, reaching manufacturing scales requires substantial increases in the numbers of channels on the chips.

Droplet-based microfluidics One form of droplet-based microfluidics is based on injection technology that produces sequential multiple discrete droplet volumes that are supported in a flowing immiscible phase.,74. Various droplet manipulations, including merging, splitting, sorting, trapping and pairing may be used to fulfill different functionalities


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required for complex NP decoration processes. Compartmentalization of reactions in individual droplets of small size enables more precise control over reaction conditions, and the convective flow inside the droplets helps to speed reactions. Moreover, the production in a manufacturing scale may be achieved by increasing the rate of micro-reactor droplets without the need for increasing the device footprint.

NP decoration requires several steps including addition of NPs and ligand solutions, removal of excess reagent, washing and recovery of decorated NPs. We have recently developed a solid-phase method for decoration of QDs and gold NPs with DNA oligonucleotides.75 Negatively charged NPs are first electrostatically associated onto the surfaces of positively charged magnetic beads (MBs), to create MB-NP conjugates (Figure 4a). Negatively charged oligonucleotides are electrostatically adsorbed onto the MB surfaces when added to a suspension of MB-NP conjugates. This creates a high local oligonucleotide concentration at the surface of the MBs that promotes the conjugation reaction. This oligonucleotide preconcentration effect has been observed to result in conjugation of oligonucleotides onto NP at kinetic rates increased by over 1000 fold in comparison to bulk solution reactions, and an oligonucleotide surface density ~5 fold higher than the best achieved for QDs when using solution-phase methods (Figure 4(b) and 1(c)). Decorated NPs were subsequently released from MBs by changing pH and ionic strength. Every step of the process, including MB-NP conjugation, DNA decoration and NP recovery was complete in less than one minute. The process was entirely governed by


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electrostatic forces, thus switching from one step to another only required a change in ionic strength and/or pH. This method offers simplicity and speed, two important criteria required for integration into the droplet microfluidic manufacturing platform for decoration of NPs.

(a)

(b)

(c)


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Figure 4. (a)Schematic representation of solid-phase QD decoration, and ligand density quantification using a FRET assay. QDs were conjugated to the surfaces of MBs to form MB-QDs, then DTPA-modified DNA was immobilized on QD surfaces. Following the release of QD-probe conjugates from MBs, the probes were hybridized with Cy3-labeled complementary targets. The density of surface-immobilized oligonucleotides was monitored using gQD-Cy3 FRET assay. (b) and (c) show the kinetics of DNA immobilization on QDs using solution-phase and solid-phase methods , respectively. Adapted with permission from reference75. Copyright 2016 American Chemical Society.

QD-to-AuNP energy transfer

Significant efforts have been directed towards improving the energy transfer efficiency in RET bioassays that use QDs. QDs offer numerous advantages as donors, but one commonly cited drawback is the physical size of the nanocrystals as this influences RET distance.76 Energy transfer originates from the center of the QD, and the physical size of the QD and its coating limits the distance between an acceptor and the donor. Given that distances may be relatively fixed, efficiency of energy transfer might be improved by consideration of spectral overlap and by dipole alignment. The use of metallic NPs as acceptors has been explored to improve RET efficiency. Metallic NPs, with gold nanoparticles being the most popular, attribute their unique optical properties to the phenomenon of localized surface plasmon


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resonance (LSPR).77 These unique optical properties include aspects such as high molar extinction coefficients and broad absorption spectra, making gold NPs efficient acceptors in non-radiative energy transfer based bioassays. Furthermore, the LSPR spectra of metal NPs can also be tuned by controlling the size, shape and chemical composition.78

Numerous studies have demonstrated the utility of gold NPs as acceptors in energy transfer bioassays that involve QDs as donors, and these have recently been reviewed.79 Such bioassays typically involve immobilization of biorecognition elements (e.g. DNA, antibodies, peptides) on the surface of both the donor and acceptor NPs. Selective interactions between the biorecognition elements and the target molecules are designed to bring the two nanoparticles together, facilitating energy transfer. Many of the early efforts to develop such bioassays involved the use of very small gold nanoparticles (diameter – 1.4 nm), which are commercially available under the brand name of Nanogold.80 The popularity of this material was facilitated by its commercial availability as a monofunctional nanoparticle. Monofunctionality ensured that each nanoparticle was conjugated to a single biomolecule/ligand. This permitted its precise arrangement around a single QD, without the complication of forming cross-linked aggregates. Such structural control becomes particularly important when assembling responsive/functional multinanoparticle constructs that respond to the presence of a target molecule. In large aggregates, the nanoparticles/recognition elements near the center of the complex may experience a different environment than those that exist at the periphery. This


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can result in non-uniform response to target molecules. The use of small AuNPs enabled controlled conjugation onto QDs, and also permitted greater energy transfer efficiencies than traditional fluorophores or quenchers.81

The use of larger AuNPs (≥ 3nm diameter) should increase energy transfer efficiencies due to the presence of stronger plasmon resonance bands. However, the increased surface area of larger AuNPs inhibits facile monofunctionalization, and provides for a greater number of permutations in which the nanoparticles may assemble. In fact, most bioassays that have used AuNPs larger than 3 nm in diameter were not designed to form responsive multi-nanoparticle complexes, but rather relied on the aggregation of nanoparticles in the presence of a target molecule to bring together the QDs and AuNPs in order to quench the QD photoluminescence.82, 83

For these bioassays, the functionality or responsiveness of the multi-nanoparticle

complex was not of concern, and thus the requirements for design were not stringent. Only a few studies have used larger AuNPs as part of a target responsive multi-nanoparticle complex84. Another challenge encountered with use large gold nanoparticles is an inner filter effect. The strong extinction coefficients of these AuNPs tends to block the excitation and emitted radiation, decreasing the signal intensities that are usually observed.84 A decrease in analytical performance is typically observed when large AuNPs are used. Our group recently demonstrated that difficulties encountered when implementing AuNPs with sizes greater than 3 nm, namely the inner filter effect and the large surface area, may be overcome by the use of monofunctionalized QDs.85 By


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monofunctionalizing the QDs, the large surface area of the AuNPs no longer presents a challenge as each QD may only bind to one AuNP. Furthermore, this configuration enables the assembly of multiple QDs around a single AuNP, decreasing the concentrations of AuNPs that need to be used, which consequently reduces the impact of the inner filter effect. In the study, two different configurations were tested as shown in Figure 5. Configuration 1 involved monovalent QD-DNA conjugates placed around a single AuNP that was functionalized with multiple copies of a complementary DNA sequence. Configuration 2 involved placing monovalently functionalized AuNP-DNA conjugates around a QD functionalized with a complementary DNA sequence. A competitive DNA displacement reaction was used to manipulate the separation between the QDs and AuNPs, enabling the investigation of energy transfer interactions for the two configurations. Furthermore, the influence of the inner filter effect was also evaluated for the two different configurations by monitoring fluorescence intensities of identical QD concentrations in the two different configurations. For configuration 1, three different sizes of AuNPs (6 nm, 13 nm and 30 nm; diameters) were investigated, while for Configuration 2, a 6 nm AuNP was used as the acceptor. The performances of the bioassays were measured on the basis of increase in fluorescence intensity as the QDs were separated from the AuNPs using the DNA strand displacement reaction. Based on the results of the study, placing fifteen QDs around a 13 nm AuNP (Configuration 1) provided optimal performance. To obtain a similar response using Configuration 2, three 6 nm AuNPs had to be placed around a single QD. While energy transfer efficiencies were similar, the absolute fluorescence intensities of


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Configuration 1 was five times greater than that of Configuration 2, allowing for better overall analytical performance as gauged by emission intensity. The difference in absolute fluorescence intensities is attributed to the reduced innerfilter effect when smaller concentrations of AuNPs are used in Configuration 1 as compared to Configuration 2. This becomes especially important with the growing trend of using low-cost detectors, such as cell phone cameras, to collect optical signals for sample analysis.86

Figure 5. Evaluating assay configurations to optimize the design of QD-AuNP based energy transfer bioassays. The schematic diagram serves to depict the multinanoparticle structures formed in solution. The graph demonstrates the increase in fluorescence intensity as a function of increasing amount of target DNA, which is added to dissociate the QDs from the AuNPs. Adapted with permission from reference86. Copyright 2015 American Chemical Society.


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Over the course of the study, two main deficiencies with handling the nanomaterials were identified. Firstly, to optimally design multi-nanoparticle complexes, the concentration of the nanomaterials must be accurately identified so that the complexes generated are reproducible. Nanoparticles do not have a defined molecular weight and this is a consequence of the variability of composition, dispersity in size and the distributions of arrangements of stabilizing ligands that cover the nanoparticles. This results in an uncertainty in their molar concentrations. Significant effort is being directed towards developing methods for the quantification of nanoparticles, and these were recently reviewed.87 Most current methods are only applicable to large nanoparticles (>10 nm), with small nanoparticles such as QDs being challenging to quantify. A novel technique represented in Figure 6 was developed by our group to determine the molar concentration of small nanoparticles such as QDs.88

Figure 6. Schematic diagram summarizing the method used to determine nanoparticle concentration. Nanoparticles are first functionalized with a suitable ligand at low equivalences (

Inorganic Nanoparticles as Donors in Resonance Energy Transfer for

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