Hybrid Nanoparticle Platform for Nanoscale Scintillation Proximity

Article Cite This: ACS Appl. Nano Mater. XXXX, XXX, XXX−XXX

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Hybrid Nanoparticle Platform for Nanoscale Scintillation Proximity Assay Colleen M. Janczak,† Isen A.C. Calderon,†,∥ Eka Noviana,†,⊥ Priyanka Hadvani,† Joo Ryung Lee,† and Craig A. Aspinwall*,†,‡,§ †

Department of Chemistry and Biochemistry, ‡BIO5 Institute, and §Department of Biomedical Engineering, University of Arizona, Tucson, Arizona 85721-00041, United States

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S Supporting Information *

ABSTRACT: β-Particle emitting radionuclides, such as 3H, 14C, 32P, 33P, and 35S, are important molecular labels due to their small size and the prevalence of these atoms in biomolecules but are challenging to selectively detect and quantify within aqueous biological samples and systems. Here, we present a core−shell nanoparticle-based scintillation proximity assay platform (nanoSPA) for the separation-free, selective detection of radiolabeled analytes. nanoSPA is prepared by incorporating scintillant fluorophores into polystyrene core particles and encapsulating the scintillant-doped cores within functionalized silica shells. The functionalized surface enables covalent attachment of specific binding moieties such as small molecules, proteins, or DNA that can be used for analytespecific detection. nanoSPA was demonstrated for detection of 3H-labeled analytes, the most difficult biologically relevant β-emitter to measure due to the low energy β-particle emission, using three model assays that represent covalent and noncovalent binding systems that necessitate selectivity over competing 3H-labeled species. In each model, nmol quantities of target were detected directly in aqueous solution without separation from unbound 3H-labeled analyte. The nanoSPA platform facilitated measurement of 3H-labeled analytes directly in bulk aqueous samples without surfactants or other agents used to aid particle dispersal. Selectivity for bound 3H-analytes over unbound 3H analytes was enhanced up to 30-fold when the labeled species was covalently bound to nanoSPA, and 4- and 8-fold for two noncovalent binding assays using nanoSPA. The small size and enhanced selectivity of nanoSPA should enable new applications compared to the commonly used microSPA platform, including the potential for separation-free, analyte-specific cellular or intracellular detection. KEYWORDS: scintillation proximity assay, core−shell nanoparticle, β-emission, radioisotope, composite nanomaterial

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INTRODUCTION Radionuclides are attractive molecular labels that do not significantly increase the size or mass of the labeled component compared to fluorescent or fluorogenic tags. Thus, radionuclides minimally affect binding, conformational changes, diffusion, and active transport. Lower background signals, and correspondingly higher sensitivity, are typically obtained for radioassays compared to fluorescence assays.1−3 Radioassays are broadly used to investigate biochemical pathways, enzyme activity,4,5 and ligand−receptor binding events. Radioassays provide high sensitivity, thus enabling quantification at sub-pM concentrations.6,7 However, 3H, 14C, 33P, and 35S are challenging to quantify directly in aqueous media due to their low decay energies (Emax from 18.6 to 1710 keV) and short β-particle penetration depths (from 4.3 μm to 7.9 mm at Emax).8−10 β-Particle emission from radiolabeled analytes is commonly quantified using liquid scintillation counting techniques where aqueous samples are dispersed in liquid scintillation cocktails (LSCs). LSCs are mixtures composed of aromatic, lightabsorbing hydrocarbons (e.g., toluene, diisopropylnaphthalene, alkylbenzenes), surfactants, and scintillant fluorophores that are used to disperse the analytes and shift the energy from the © XXXX American Chemical Society

decay event into the UV or visible range of the electromagnetic spectrum via energy transfer.8,11 While highly sensitive, LSC enables measurement only at a single time point for most biomolecular systems due to the high proportion of organic solvent. Solid scintillators, prepared from scintillant-doped polymers (e.g., polyvinyltoluene or polystyrene) or inorganic crystals (e.g., CdWO4, CeF3) and molded to form microwells or synthesized or ground into microparticle structures, can be used in place of LSC to enable aqueous measurement, though typically with lower sensitivity.8,12,13 Porous scintillating polymer bead resins have been developed for the detection of β-and α-particles, but these resins typically rely upon the extraction of radioactive metals using chelating ligands at the particle surface, limiting their use for biologically relevant βemitters such as 3H, 35S, and 33P.14−17 In both liquid and solid scintillation measurements, the lack of inherent analyte selectivity presents a major drawback. Both approaches detect the β-decay event resulting from the isotope via the energy transferred through the matrix. Isotopes yield Received: November 23, 2018 Accepted: February 7, 2019

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DOI: 10.1021/acsanm.8b02136 ACS Appl. Nano Mater. XXXX, XXX, XXX−XXX

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ACS Applied Nano Materials

potential application of SPA for intracellular measurements under most circumstances. Thus, reduced SPA particle sizes may provide enhanced utility for SPA applications as well as enable new radioassay platforms to be developed and implemented for investigating biological systems. Here, we report a nanoparticle-based SPA format (referred to herein as nanoSPA, Figure 1a) based upon a recently reported core−shell, radioisotope-responsive nanomaterial.35 PS was selected as the core material due to its low cost and compatibility with scintillant fluorophores.36−38 The addition of silica shells to the scintillant fluorophore-doped PS particles makes the particles hydrophilic without using surfactants, which are often required for the dispersion of PVT and PS nano- and microparticles in aqueous samples, and facilitates the covalent attachment of binding receptors to the exterior nanoparticle surface. Furthermore, while the density of the core−shell nanoparticles allows the particles to remain dispersed in a sample, the core−shell nanoparticles can be easily recovered via centrifugation enabling sample enrichment and isolation as well as reuse of the particles if desired.

distinct decay energies, but the distribution of decay energies for a given isotope is indistinguishable when incorporated into different molecular species, limiting the selectivity. For example, substrates and products of an enzymatic reaction cannot be distinguished using LSC or solid scintillation approaches in a homogeneous manner. Radioassays, therefore, often require preseparation of the analyte mixture, which imparts significant limitations on the assay, particularly with respect to spatial and temporal resolution. Scintillation proximity assays (SPAs) were developed to overcome these limitations and prepare homogeneous radioassays.8,18 SPA is a derivative of solid scintillation counting, wherein specific binding of radiolabeled analyte to a solid scintillator surface serves to increase the probability of energy absorption by the scintillator (Figure 1) and thus detection of

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EXPERIMENTAL SECTION

Materials. Styrene, alumina, p-terphenyl (pTP), and 1,4-bis(4methyl-5-phenyl-2-oxazolyl)benzene (dimethyl POPOP) were purchased from Acros Organics. Tetraethylorthosilicate (TEOS), (3mercaptopropyl)trimethoxysilane (MPTS), (3-aminopropyl)triethoxysilane (APTES), 2,2′-azobis(2-methylpropionamidine) dihydrochloride (AIBA), N-(3-(dimethylamino)propyl)-N′-ethylcarbodiimide hydrochloride (EDAC HCl), N-hydroxysuccinimide (NHS), biotin-PEG-NHS (Mw = 3000), and Tween-20 were obtained from Sigma-Aldrich (St. Louis, MO). Dimethyl sulfoxide was purchased from Fisher Chemical (Hampton, NH). PEG6−9-dimethylchlorosilane (90%) was purchased from Gelest (Morrisville, PA). Isopropyl alcohol (IPA), chloroform, aqueous ammonium hydroxide (28%), and acetonitrile were obtained from EMD Millipore (Billerica, MA). Ethyl alcohol was purchased from Decon Laboratories (King of Prussia, PA). BioCount liquid scintillation cocktail was purchased from Research Products International (Mt. Prospect, IL). 3H-labeled sodium acetate (1.59 Ci/mmol) was purchased from PerkinElmer (Waltham, MA). NeutrAvidin was purchased from Pierce Thermo Scientific (Waltham, MA), and Nanosep centrifugal devices with 3 kDa MWCO membranes were purchased from Sartorius (Göttingen, Germany). All chemicals except styrene were used as received. Inhibitor was removed from styrene by passing the monomer through a 0.5 cm diameter by 3 cm long alumina column immediately prior to use. Acrydite-functionalized DNA oligomers 5′-/Acryd/AAA CCC TGT AAA CCC TGT AAA CCC TGT AAA-3′ and aminefunctionalized DNA oligomers 5′-/5AmMC6/TTT ACA GGG TTT ACA GGG TTT ACA GGG TTT-3′ (complementary oligomer), 5′-/5AmMC6/TTT ACA GAG TTT AGA GGG TTG ACA GGG ATT (4-base mismatched oligomer), and 5′-/5AmMC6/ AGC ACT GAG AGA AGA GTG TTG ACA CAT ATT-3′ (randomsequence 30-base oligomer) were obtained from Integrated DNA Technologies, Inc. (Coralville, IA). All scintillation measurements were performed using a Beckman LS 6000 IC scintillation counter. Precautions. All radioactive materials should be handled and disposed according to federal, state, and local guidelines. Styrene is flammable, is an irritant, and is a health hazard. Ethyl acetate, ethyl alcohol, isopropyl alcohol, TEOS, AIBA, and acetonitrile are flammable and are irritants. MPTS and pTP are irritants and hazardous to aquatic life. APTES is corrosive and an irritant. EDAC HCl and PEG6−9-dimethylchlorosilane are irritants. Chloroform exhibits acute toxicity and is a health hazard. Ammonium hydroxide solution is corrosive, an irritant, and hazardous to aquatic life.

Figure 1. nanoSPA platform. (a) Schematic illustrating the function of nanoSPA. When radioisotope decay occurs near the nanoSPA surface (right), enhanced photoemission is observed, compared to radioisotope decay in solution beyond the penetration depth of the βparticle (left). TEM images of nanoSPA particles with (b) hydroxylfunctionalized silica shells, (c) amine-functionalized silica shells, and (d) thiol-functionalized silica shells.

the decay event. When the decay event occurs near the scintillator surface, the detection probability is enhanced resulting in increased photon emission from the solid scintillator upon analyte binding.8,18 SPA is particularly useful for low energy β-particles that exhibit low penetration depths, e.g., 3H (dp ≈ 0.5 μm in H2O).8 Consequently, SPA lends itself to monitoring binding kinetics under steady state conditions,18−22 as well as separation-free, selective quantification of radiolabeled analytes in complex mixtures. Modern SPA particle formats utilize scintillant-doped polyvinyltoluene (PVT) or polystyrene (PS) particles, or yttrium silicate (YSi) or yttrium oxide (YOx) microparticles, to which receptors have been covalently attached.20,23−26 SPA has been used to measure quantitative binding and/or binding kinetics of enkephalins, thyroxin, morphine, inositol phosphates, and many other analytes.19,21,27−34 These encouraging applications demonstrate the key advantages of SPA, including the ability to perform time-resolved analyte measurements, high selectivity, and separation-free assays. Current SPA platforms are limited by the large size of SPA particles (5− 10 μm) which results in low surface area to volume ratios, as well as particle settling. Additionally, these sizes preclude the B


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ACS Applied Nano Materials Preparation of nanoSPA Particles. Scintillant fluorophoredoped PS core nanoparticles were prepared following the protocol described previously.35 Briefly, styrene (3 g, inhibitor-free) was added to 100 mL of degassed H2O in an Ar-flushed 500 mL round-bottomed flask heated to 70 °C in an oil bath. Polymerization was initiated by adding 10 mg of AIBA dissolved in approximately 200 μL of H2O to the reaction flask. The H2O/styrene mixture was stirred rapidly for at least 6 h. Excess styrene and some H2O were removed from the nanoparticle solution under reduced pressure using a rotary evaporator to reduce the total volume to approximately 100 mL. A 1−2 mL aliquot of the solution was removed and lyophilized to determine the mass-based concentration (mg/mL) of polystyrene core nanoparticles. PS nanoparticles were doped with scintillant fluorophores by dissolving 53 mg (135 μmol) of dimethyl POPOP and 262 mg (1.14 mmol) of pTP in 20 mL of 1:9 isopropyl alcohol/chloroform (v:v). Scintillant fluorophores dissolved in solvent were added directly to the aqueous PS nanoparticle solution in a 500 mL round-bottomed flask. The nanoparticle solution was agitated using a bath sonicator for several minutes to disperse organic solvent droplets throughout the H2O, and the solution was stirred rapidly for at least 1 h. Organic solvents were then removed under reduced pressure using a rotary evaporator. Silica shells were added to scintillant fluorophore-doped PS nanoparticles by dispersing 2 mL of PS nanoparticle stock solution (approximately 56 mg of nanoparticles) in 200 mL of isopropyl alcohol with 38 mL of H2O. For hydroxyl- or thiol-functionalized silica shells, 7.5 mL of NH4OH was added to the flask. For aminefunctionalized shells, 5.0 mL of NH4OH was added to the flask. The dispersion was stirred briskly for several minutes while either 2.0 mL of TEOS or a mixture containing 1.8 mL of TEOS and 0.2 mL of either APTES or MPTS was added dropwise. Stirring was continued for 1 h before particles were collected by centrifugation and rinsed several times with H2O. Transmission Electron Microscopy. Transmission electron micrographs of nanoparticles were obtained by drying a 10 μL aliquot of nanoparticle solutions on carbon films deposited on copper grids (Electron Microscopy Sciences, Hatfield, PA). Samples were observed using a Technai G2 Spirit transmission electron microscope at 100 kV accelerating voltage (FEI Company, Hillsboro, OR). Direct Attachment of 3H-Sodium Acetate to nanoSPA. 3Hlabeled succinimidyl acetate was synthesized by reacting 3H-labeled sodium acetate (300 nmol, 45 μL of 1 μCi/mL, 150 mCi/mmol) with EDAC HCl (770 μmol) and NHS (800 μmol) in 100 μL of H2O. After 1.5 h reaction time, increasing volumes of the mixture (1−10 μL) were added to 0.1 mg of amine-functionalized nanoSPA in 150 μL of 25 mM borate buffer, pH 8.0. As a control, equivalent activities of 3H-labeled sodium acetate (not treated with EDAC HCl and NHS) were added to amine-functionalized nanoSPA. After 30 min reaction time, 145 μL of the reaction mixture was diluted with 10 mL of H2O immediately prior to measurement with the scintillation counter. nanoSPA Functionalization with Biotin and PEG. Aminefunctionalized nanoSPA particles (200 mg) were suspended in 10 mL of phosphate buffered saline (PBS) (0.1 M sodium phosphate, 0.15 M NaCl, pH 7.40). Biotin-PEG-NHS (23 mg, Mw = 3000) was added into the particle solution and allowed to react overnight (>12 h) with mixing. The particles were washed with PBS three times, and then resuspended in PBS to make a 20 mg/mL particle solution. A 100 mg sample of biotinylated nanoSPA was collected by centrifugation at 1600g for 10 min and then washed with acetonitrile three times. The particles were resuspended in a 5 mL solution of 2% PEG6−9dimethylchlorosilane in dry acetonitrile (v/v) and allowed to react, with mixing overnight. The particles were then washed successively three times with acetonitrile, three times with ethanol, and three times with water. The particles were finally suspended in water (20 mg/ mL). Another sample of nonpassivated particles was prepared using biotin-PEG-NHS but were not treated with PEG6−9-dimethylchlorosilane. A third set of particles was not biotinylated but was passivated with treatment of PEG6−9-dimethylchlorosilane for comparison.

Radiolabeling of NeutrAvidin. NeutrAvidin was tritiated by conjugating to 3H-labeled sodium acetate using EDAC/NHS coupling chemistry. 3H-labeled sodium acetate (390 nmol, corresponding to 600 μCi activity) was reacted with EDAC HCl (200 μmol), and NHS (200 μmol) in 100 μL of DMSO at 25 °C for 1 h. Protein (6.0 mg of NeutrAvidin) was dissolved in 1 mL of PBS and added to the 3Hlabeled sodium acetate/EDAC HCl/NHS mixture and allowed to react at room temperature overnight. 3H-labeled NeutrAvidin was then purified from other tritiated species using a molecular weight cut off centrifugal filter (3 kDa for NeutrAvidin) and resuspended in PBS. Biotin/NeutrAvidin nanoSPA. Biotin-functionalized nanoSPA (1.5 mg) was dispersed in 1 mL of PBS with increasing activities of 3 H-labeled NeutrAvidin (0.25 nmol/μL protein solution with 334 nCi/μL activity and 1.33 Ci/mmol specific activity). As a control for nonproximity effects, equal activities of 3H-labeled sodium acetate were added in place of labeled protein. All samples were prepared in triplicate. nanoSPA Functionalization with ssDNA. Thiol-functionalized nanoSPA (50 mg) was dispersed in 1 mL of 0.1 M phosphate buffer pH 7.4 and was reacted with 80 nmol of acrydite-functionalized 30mer ssDNA for 72 h at room temperature with brisk stirring. Particles were rinsed several times with PBS and resuspended in the same buffer. Control nanoSPA was prepared using the same procedure by omitting the ssDNA from the reaction mixture. Radiolabeling of ssDNA. Complementary and random-sequence ssDNA were tritiated via conjugation to 3H-labeled sodium acetate using EDAC/NHS coupling chemistry. 3H-labeled sodium acetate (100 nmol corresponding to 154 μCi activity) was reacted with EDAC HCl (10 μmol) and NHS (10 μmol) in 30 μL of dimethylformamide (DMF) by stirring the solution for 3 h at 25 °C. Amine-functionalized oligonucleotide (100 nmol) was dissolved in 90 μL of 0.1 M phosphate buffer pH 9.5 and mixed with 3H-labeled sodium acetate/EDAC HCl/NHS solution. The mixture was stirred briskly for 3 h at room temperature and then separated using a Sephadex G-50 column (packed bed volume approximately 3 mL). Column fractions were subjected to scintillation counting, and specific activity of radiolabeled ssDNA was estimated by dividing the area of the labeled ssDNA peak by total peak area and multiplying by 154 μCi/100 nmol. ssDNA Hybridization nanoSPA. ssDNA-functionalized nanoSPA (5 mg) were dispersed in 5 mL of annealing buffer (100 mM phosphate, 50 mM NaCl, and 10 mM EDTA; pH 7.4) containing 0.1% Tween-20 in a 7 mL scintillation vial. nanoSPA samples 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 for 5 min and allowed to cool to room temperature for 40 min while shaking at 300 rpm prior to each scintillation measurement. As a control, nonfunctionalized core−shell nanoparticles 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 (complementary ssDNA oligomer, 4-base mismatch oligomer, and random-sequence oligomer) was also tested by binding inhibition. A 5 mg portion of ssDNA-functionalized nanoSPA was saturated with 3H-labeled complementary ssDNA (approximately 6 nmol) and titrated with increasing amounts of unlabeled complementary, 4-base mismatch, or random-sequence 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 being shaken at 300 rpm. Determination of Detection Limits and Selectivity. The limits of detection (LOD) for the direct attachment of 3H-labeled sodium acetate, and for the binding of 3H-labeled NeutrAvidin to biotin-functionalized nanoSPA, were calculated using the equation LOD = ksblk/m, where k = 3, sblk is the standard deviation of the measurements of the blank samples, and m is slope of the leastsquares fit line for data points at increasing analyte concentrations. For the direct attachment of 3H-labeled sodium acetate, the first 6 data points were used for least-squares fitting, omitting the data point at 6 nmol. For the binding of 3H-labeled NeutrAvidin, the data points C


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ACS Applied Nano Materials from 0 to 0.5 nmol were used in the calculation. For these model assays, selectivity at the highest activity was calculated as the ratio of the magnitude of the SPA response divided by the magnitude of the nonspecific binding response.

Initially, the surface charge characteristics of nanoSPA particles were examined by measuring ζ potential in 100 mM NaCl solutions of varying pH, from 3 to 10 (Figure S1a, Supporting Information) to determine the stability of aqueous nanoSPA dispersions.43−46 The ζ potential of nanoSPA particles with amine-functionalized shells decreased significantly with increasing pH, approaching −8.0 mV at pH 10. This trend can be attributed to protonation of the amine at low pH, and a combination of deprotonation and hydrolysis at high pH.47,48 The ζ potential of nanoSPA particles with hydroxylor thiol-functionalized shells was ≤−5 mV at pH 4 or lower, and decreased slightly to −11 and −7 mV, respectively, at pH 5. These suggest that nanoSPA particles with hydroxyl- or thiol-functionalized shells may aggregate in very acidic samples, and nanoSPA particles with amine-functionalized shells may aggregate at very alkaline pH. However, biological samples are unlikely to have such extreme pH conditions. Furthermore, no change in particle size was indicated by light scattering or turbidimetry measurements acquired over a period of 10 days (Figure S1b−d, Supporting Information), demonstrating that nanoSPA particles with hydroxyl-, amine-, and thiol-functionalized shells are readily dispersed in aqueous media. The amine and thiol functional groups on the silica shells were subsequently used for covalent attachment of binding elements to nanoSPA to demonstrate proof-of-concept nanoSPA-based radioassays. Three model nanoSPAs were utilized: (1) 3H-labeled sodium acetate was covalently bound to the nanoSPA particle surface using N-(3-(dimethylamino)propyl)N′-ethylcarbodiimide hydrochloride (EDAC HCl) and Nhydroxysuccinimide (NHS) coupling. (2) Biotin-functionalized nanoSPA particles were used to measure 3H-labeled NeutrAvidin. (3) ssDNA-functionalized nanoSPA was used to bind the complementary ssDNA oligomer. These assays were chosen to demonstrate selectivity, sensitivity, and maximum enhancement efficiency. To validate the function of nanoSPA, 3H-sodium acetate was covalently attached to the nanoSPA particle surface to determine if signal due to β-particles emitted by bound 3H (proximity) could be distinguished from signal due to βparticles emitted by unbound 3H which has a longer mean distance from the nanoSPA core (nonproximity). Nonproximity-derived signal, which represents the normal background signal for all SPA applications, must be significantly lower than proximity-derived signals for useful SPA measurements. Figure 2 shows the nonproximity and proximity responses for nanoSPA upon addition of increasing 3H activity. The proximity signal was ca. 30× greater than the nonproximity signal at 6.2 nmol (930 nCi) of 3H. At lower activity, an enhancement greater than 4× was observed for 0.1 nmol (15.5 nCi) 3H activity (42 CPM vs 9.4 CPM). The limit of detection for this assay was 6.0 nCi 3H, corresponding to approximately 0.04 nmol, or 280 nM. While the ratio of proximity to nonproximity signals is dependent on several factors including nanoSPA concentration, analyte concentration, Emax, and specific activity, this experiment clearly demonstrates the feasibility of nanoSPA measurements that are analogous to those performed using slab-style SPA scintillators, such as microwell plates, with similar sensitivity. nanoSPA was then compared to SPA particles with diameters >5 μm, referred to as microSPA, which are commonly used for SPA measurements (Figure S2, Supporting Information). A comparison of the scintillation response of nanoSPA and microSPA particles (Supporting Information)

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RESULTS AND DISCUSSION nanoSPA particles were prepared using a core−shell architecture to maximize dispersion and aqueous compatibility yet retain an environment suitable for β-decay stimulated scintillation. The silica shell increases compatibility with aqueous solutions, reduces aggregation typically seen with PS particles, and serves as a platform for further surface modification whereas the PS core enables dispersion of scintillant fluorophores in a hydrophobic environment that serves as an energy transfer matrix. SPA efficiency is partly dependent upon the efficiency with which energy from radioactive decay is absorbed, which in turn is related to the path length of β-particles through the absorbing medium;8 thus smaller PS core nanoparticles might lead to severely reduced absorption and insufficient signal during measurement in commercially available liquid scintillation counters. Conversely, enhanced energy absorption efficiency leads to greater signal, but may also result in larger nonproximity signal, making signal differentiation difficult or impossible. The proximity signal to nonproximity signal ratio is also dependent upon radionuclide activity, Emax, and particle concentration. Thus, nanoSPA particles were designed with PS cores large enough to absorb adequate energy and provide sufficient signal at nCi activities, but small enough to reduce nonproximity signals, remain easily dispersible in solution, and maximize the surface area to volume ratio. The PS cores of nanoSPA particles were prepared in a surfactant-free emulsion polymerization, into which scintillant fluorophores were incorporated after polymerization via solvent-induced swelling.39−41 For initial nanoSPA experiments p-terphenyl (pTP) was used as a primary scintillant fluorophore, and 4-bis(4-methyl-5-phenyl-2-oxyzolyl)benzene (dimethyl POPOP) was used as a wavelength-shifting, secondary scintillant fluorophore. Previously, we reported silanol terminated silica shell deposition.35 Here, we have expanded the range of surface functionalities by combining 10% functional silane [3-(aminopropyl)triethoxysilane (APTES) or 3-mercaptopropyl trimethoxysilane (MPTS)] with tetraethylorthosilicate (TEOS) during shell synthesis to make amine- or thiol-functionalized silica shells on the scintillant-doped cores. Functionalization in this manner is simple and significantly expands the surface functionalization capabilities by providing the necessary surface functionality for widely used linkers, including N-hydroxy succinimidyl esters and maleimides. Transmission electron microscopy (TEM) images (Figure 1) show 150−200 nm scintillant-loaded PS core nanoparticles (PS core nanoparticles) surrounded by denser silica shells 25−50 nm thick. Whereas the scintillantloaded PS cores appear smooth, irrespective of silane modification, the morphology of the silica shell varies dependent upon the silane functionality. For both TEOS and APTES modified silica shells, a smooth surface was observed (Figure 1b−c) compared to MPTS, where a rough surface was observed (Figure 1d). Thus, the surface morphology and roughness of silica shells on core−shell nanoparticles depend on reaction pH and the rates of silica sol nucleation and growth, similar to the observations of Lu et al.42 D


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Figure 3. Response of biotin-functionalized nanoSPA as a function of 3 H-labeled NeutrAvidin (blue ○) compared to nonspecific adsorption of 3H-labeled NeutrAvidin to nanoSPA lacking biotin (green △) and nonproximity effects of 3H-labeled sodium acetate added to biotinylated nanoSPA (red ◇).

proximity signal observed for biotin null particles. Aminefunctionalized silica is susceptible to nonspecific protein adsorption;52−54 thus addition of PEG to the nanoSPA surface served to minimize the signal due to nonspecific adsorption. Compared to the nonproximity signal, an 8-fold increase in signal response was observed for nonoptimized biotinfunctionalized particles at 1.5 nmol of NeutrAvidin, which is comparable to commercial microparticle SPA assays, that exhibit 3× to 15× enhancement.19,20,23,24,27 Based on these data, the limit of detection for this assay is 9 pmol of 3Hlabeled NeutrAvidin which is more than 10 times lower than nonspecific adsorption of 3H-labeled NeutrAvidin. These results demonstrate that functionalized nanoSPA can be used to selectively detect 3H-labeled protein. The NeutrAvidin used in these experiments was labeled using EDAC/NHS coupling of 3H-labeled sodium acetate, demonstrating that nanoSPA could be used for protein analytes (e.g., project-specific mutants) for which there is no commercially available 3H-labeled material. In addition, the data shown in Figure 3 demonstrate that nanoSPA can be used to obtain sub-nmol detection limits, though this may be improved by increasing the 3H-labeling efficiency, which for these experiments was 20%. While many protein-labeling protocols for reactive fluorescein and rhodamine tags recommend limiting reagent concentrations to yield 2−6 fluorophores per protein to prevent quenching,55−57 3H can be incorporated into proteins using chemical or metabolic labeling techniques without concern of quenching.58−60 In addition, small molecules can be synthesized with several 3H per molecule, making greater sensitivity possible for radioassays. Finally, nanoSPA was evaluated for specificity using a DNAbinding interaction. nanoSPA particles were functionalized with a 30-base ssDNA oligomer to which the complementary, 3 H-labeled strand (57 nCi/nmol) was hybridized. As expected, the measured signal increased with increasing 3H-labeled complementary strand due to hybridization with the ssDNA oligomer immobilized on the nanoSPA surface (Figure 4a). The sigmoidal shape of the curve observed in Figure 4a may be due to both a decreased probability of binding during the set annealing time at low ssDNA (less than 1 nmol) concentration and surface saturation at high ssDNA concentrations (5 nmol

Figure 2. Proximity-dependent signal enhancement. (a) Scintillation response of 3H-labeled sodium acetate (blue ◇) covalently linked to nanoSPA particle surface compared to unlinked 3H-labeled sodium acetate in free solution (red □). (b) The same data expanded to show the scintillation response at low activities.

shows that while the large SPA particles demonstrate a ca. 3× greater absolute scintillation response (Figure S3, Supporting Information), the net enhancement, determined from the ratio of proximity-derived signal to nonproximity-derived signal, is equivalent, if not better, for nanoSPA compared to microSPA at 0.5 μCi/mL (Figure S3, Supporting Information). Though, on a per decay event basis, larger particles are expected to give a larger absolute signal due to the closer match to the βparticle path length and greater absorption of energy, nanoSPA provides similar enhancements, likely due to higher surface area available for binding and better dispersion of nanoSPA.49−51 Following the assessment of sensitivity, we prepared biotinfunctionalized nanoSPA to demonstrate the specificity of the nanoSPA radioassay sensor platform. Briefly, nanoSPA particles with amine-functionalized silica shells were further modified with PEG6−9-dimethylchlorosilane with either 0% or 4% biotin-PEG-NHS ester (Mw = 3000). nanoSPA particles were then used to measure 3H-labeled NeutrAvidin directly in pH 7.4 phosphate buffer. Figure 3 shows the response of nanoSPA particles as a function of increasing NeutrAvidin. When 3H-labeled sodium acetate was used as a nonproximity marker, the observed background was low, measuring ≤130 CPM at the highest 3H activity. The specificity was evaluated using PEG6−9-coated nanoSPA particles that contained or lacked PEG-biotin. Upon addition of NeutrAvidin, the scintillation response for the biotinylated nanoSPA increased while the signal for nanoSPA lacking biotin remained equivalent to the nonE


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detection limit for the competitive assay is estimated to be 1.4 nmol of unlabeled complementary ssDNA, with ca. 2× and 4× selectivity for the complementary versus 4-base mismatch and the complementary versus random sequences, respectively. Despite the similarity between the complementary and the 4base mismatch oligomers, ssDNA-functionalized nanoSPA can be used to differentiate between the two sequences. Overall, data from these three model assays support the selectivity of the nanoSPA platform and demonstrate a wide potential applicability for homogeneous radioassays, with signal enhancements equivalent to existing microparticle SPA platforms but with substantially improved dispersibility.

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CONCLUSIONS nanoSPA particles provide selectivity and enhanced sensitivity for radiolabeled analytes in an easily dispersible and readily functionalized nanoparticle geometry. Selective and sensitive detection can be performed directly in aqueous samples without the need for LSC which may lead to denaturation and other deleterious processes and thus hinder the analysis. As such, nanoSPA can potentially be used for temporally resolved measurements in biological samples where sample volume is limited and/or where larger particles are more likely to settle during the time course of the measurement. While the absolute signal for nanoSPA particle is lower than that observed for similarly prepared microparticles due to the smaller signal generation path length in nanoSPA, the signal enhancements for proximity versus nonproximity signals were comparable between the two particle size scales. Furthermore, the silica surface provides a more easily modified substrate for attachment of recognition elements compared to common polymer microSPA beads enabling versatile, separation-free analysis using SPA. Due to the small diameter of nanoSPA and the previously demonstrated bioinert nature of silica nanoparticles,61−64 future applications of nanoSPA may prove useful for application to complex biological systems that cannot be assessed using current radioisotope detection technologies.

Figure 4. nanoSPA response to DNA hybridization. (a) ssDNAfunctionalized (blue ○) and unfunctionalized nanoSPA particles (red □) were incubated with increasing concentrations of 3H-labeled complementary ssDNA oligomers. (b) Competitive radioassay for 3Hlabeled DNA. ssDNA-functionalized nanoSPA were incubated in the presence of 6 nmol of 3H-labeled complementary ssDNA with addition of the indicated concentration of 30-base unlabeled complementary (orange ○), 4-base mismatch (gray △), and random (violet ◇) DNA sequences. (c) Percent signal saturation for the competitive radioassay at 6 nmol of ssDNA where * and ** represent p < 0.005 and p < 0.001, respectively, in a two-tailed Student’s t test.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsanm.8b02136.

and greater), as only 8 nmol of ssDNA was used for nanoSPA functionalization. The utility and selectivity of nanoSPA were further evaluated by performing a competitive binding assay in which unlabeled ssDNA was added to nanoSPA previously saturated with 6 nmol of 3H-labeled complementary ssDNA. The nanoSPA samples were then subjected to the same heating and cooling cycles used for hybridization of the 3H-labeled complementary ssDNA strand. Under these conditions, the unlabeled complementary ssDNA was expected to replace a proportional amount of 3H-labeled complementary ssDNA at the nanoSPA surface. As predicted, the measured signal at nanoSPA saturation with 3H-labeled ssDNA complementary strand decreases with increasing unlabeled complementary ssDNA by up to 64% over the range from 0 to 6 nmol, but only by 36% when treated with an unlabeled ssDNA with 4 mismatched bases. Furthermore, the saturated nanoSPA signal decreased by only 16% when treated with an unlabeled random-sequence 30-base ssDNA oligomer. The minimum 3 H-labeled complementary strand detected was 1 nmol. The

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Materials, Methods, and data for the comparison of nanoSPA to microSPA particles (PDF)

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Craig A. Aspinwall: 0000-0002-4359-5812 Present Addresses ∥

I.A.C.C.: Department of Bioengineering, Northeastern University, Boston, MA 02115, USA ⊥ E.N.: Department of Chemistry, Colorado State University, Fort Collins, CO 80523, USA Author Contributions

The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. F


Article

ACS Applied Nano Materials Notes

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The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported in part by the National Science Foundation under Grant 1807343, the National Institutes of Health via the National Institute of Biomedical Imaging and Bioengineering under Grant R21EB019133, and the National Institute of General Medical Sciences under Grant 1R01GM116946. The transmission electron microscope used in this study was purchased by the Life Science North Imaging Facility at the University of Arizona with funding from the National Institutes of Health, under Grant 1S10OD011981-01. The content is solely the responsibility of the authors and does not necessarily represent the official views of the sponsors.

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Hybrid Nanoparticle Platform for Nanoscale Scintillation Proximity

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