pubs.acs.org/Langmuir © 2009 American Chemical Society
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Automated Oligonucleotide Solid-Phase Synthesis on Nanosized Silica Particles Using Nano-on-Micro Assembled Particle Supports )
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Carole Farre,† Muriel Lansalot,‡ Rana Bazzi,§ Stephane Roux,§ Christophe A. Marquette, Ga€elle Catanante, Loı¨ c J. Blum, Nicolas Charvet,^ Cedric Louis,^ and Carole Chaix*,† †
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Universit e de Lyon, Univ. Lyon 1, UMR 5180, Laboratoire des Sciences Analytiques, B^ at. CPE - 43, Bd du 11 Novembre 1918, Villeurbanne, 69622 Cedex, France, ‡Universit e de Lyon, Univ. Lyon 1, CPE Lyon, CNRS UMR5265, Laboratoire de Chimie, Catalyse, Polym eres et Proc ed es (C2P2), LCPP Team, 43 Bd du 11 Novembre 1918, Villeurbanne, 69616 Cedex, France, §Universit e de Lyon, Univ. Lyon 1, UMR 5620, Laboratoire de Physico-Chimie des Mat eriaux Luminescents, 43, Bd du 11 Novembre 1918, Villeurbanne, 69622 Cedex, France, Universit e de Lyon, Univ. Lyon 1, UMR 5246, Institut de Chimie et Biochimie Mol eculaires et Supramol eculaires, B^ at. CPE - 43, Bd du 11 Novembre 1918, Villeurbanne, 69622 Cedex, France, and ^NANOH S.A.S., Parc GVIO, Place de l’Europe, 38070 Saint Quentin Fallavier, France Received September 22, 2009. Revised Manuscript Received October 27, 2009 This article describes an original strategy to enable solid-phase oligodeoxyribonucleotide (ODN) synthesis on nanosized silica particles. It consists of the reversible immobilization of silica nanoparticles (NPs) on micrometric silica beads. The resulting assemblies, called nano-on-micro (NOM) systems, are well adapted to ODN synthesis in an automated instrument. First, NPs are derivatized with OH functions. For NOM assembly preparation, these functions react with the silanols of the microbeads under specific experimental conditions. Furthermore, OH groups allow ODN synthesis on the nanoparticles via phosphoramidite chemistry. The stability of the NOM assemblies during ODN solidphase synthesis is confirmed by scanning and transmission electron microscopy (SEM and TEM, respectively), together with dynamic light scattering analyses. Then, the release of ODN-functionalized nanoparticles is performed under mild conditions (1% NH4OH in water, 1 h, 60 C). Our technique provides silica nanoparticles well functionalized with oligonucleotides, as demonstrated by hybridization experiments conducted with the cDNA target.
Introduction The intensive research carried out during the past decade in the field of nanotechnology now gives access to a broad range of nanoparticles in a monodisperse format with various shapes, sizes, and compositions. These well-controlled spherical supports are of increasing interest in both diagnostics and imaging.1-3 In the domain of in vitro diagnostics, trends are shifting toward the invention and utilization of miniaturized integrated analysis systems. Biomolecular probes, in the form of functionalized microbeads, are often used in such lab-on-a-chip devices either to extract the target biomolecules from samples selectively1 or to detect them.2 For imaging applications, new nanomaterials have been developed with optical-encoding abilities for selectively tagging a wide range of medically important targets, including bacteria, cancer cells, and individual molecules such as proteins and DNA.3 The main advantage of a spherical support over a planar support is the high specific surface area that it offers for biomolecule anchoring. Among the different types of nanomaterials, silica has emerged as an interesting support owing to its reproducible synthesis and versatile chemistry,4 its low in vivo toxicity,5 and its surface biocompatibility.6 In this field, Trau et al. have described an original method of “colloidal bar coding” that *Corresponding author. E-mail:
[email protected]. (1) Elaissari, A. Colloidal Biomolecules, Biomaterials, and Biomedical Applications; Marcel Dekker: New York, 2004. (2) Wang, L.; Lofton, C.; Popp, M.; Tan, W. Bioconjugate Chem. 2007, 18, 66. (3) Wang, L.; O’Donoghue, B.M.; Tan, W. Nanomedecine 2006, 1, 413. (4) Smith, J.; Wang, L.; Tan, W. Anal. Chem. 2006, 25, 848. (5) Jin, Y.; Kannan, S.; Wu, M.; Zhao, J. X. Chem. Res. Toxicol. 2007, 20, 1126. (6) Radin, S.; El-Bassyouni, G.; J. Vresilovic, E.; Schepers, E.; Ducheyne, P. Biomaterials 2005, 26, 1043.
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involves multifluorescent silica colloids adsorbed on microsized polymer beads.7 Each of the fluorescent beads can be functionalized with an oligonucleotidic sequence. The adhesion of nanosized silica particles is achieved by using polyelectrolytes for the electrostatic entrapment of the nanoparticles in a multilayer system.8 Specific and identifiable combinations of fluorescent dyes in silica colloids afford access to powerful encoding combinatorial libraries. Potentially, these strategies can result in extremely large libraries (>1010 compounds) and might be generically suitable for applications such as single nucleotide polymorphism (SNP) genotyping, gene expression, and drug discovery.9 Nevertheless, one drawback of nanoparticle strategies concerns the current biomolecule grafting methods that do not lead to highly functionalized beads or conjugates with well-oriented biomolecules on the surface. Such techniques often result in systems with a high signal background and low probe functionalization. Furthermore, purification methods of (biomolecule-nanoparticle) conjugates are tedious and timeconsuming because the centrifugation of nanosized objects often leads to their aggregation. In this context, we focused on the development of a new strategy to obtain ODN-functionalized nanosized particles that can be used to target DNA in biological samples. We have considered the possibility to perform direct and automated ODN solid-phase synthesis on nanosized silica supports. Silica is the standard material for this purpose because it can be easily (7) Lawrie, G. A.; Battersby, B. J.; Trau, M. Adv. Funct. Mater. 2003, 13, 887. (8) Grondahl, L.; Battersby, B. J.; Bryant, D.; Trau, M. Langmuir 2000, 16, 9709. (9) Battersby, B. J.; A. Lawrie, G.; Trau, M. Drug Discovery Today: HTS Suppl. 2001, 6, S19.
Published on Web 12/11/2009
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derivatized with the functional arm that serves to initiate solidphase synthesis. However, because of technical considerations associated with ODN automated synthesis (notably the high pressure used in the instrument), the solid supports must be micrometer sized to be held in the synthesizer during the ODN synthesis. In a previous paper, we described the elaboration of silica colloid aggregates suitable for automated DNA synthesis.10 These aggregates offer a high specific surface area on which to carry out oligonucleotide solid-phase synthesis. However, the amount of ODN material per gram of support was lower than expected, revealing that oligonucleotide synthesis was mainly initiated on the external surfaces of the aggregates. Nevertheless, these ODN-functionalized silica aggregates efficiently increased ELOSA (enzyme-linked oligosorbent assay) test sensitivity when spotted on a microarray. Targeting the same goal, another approach was reported by Seliger et al.11 ODNs were synthesized on the surfaces of composite beads constituted from a micrometric polystyrene core coated with either silica or gelatin, onto which were deposited dextran-coated magnetite nanoparticles. Herein, we report the synthesis of a two-particle assembly, called a nano-on-micro (NOM) system, composed of nanosized silica particles of about 50 nm immobilized onto 4 μm silica microbeads. The term NOM has already been used in the literature for similar assemblies elaborated with quantum dots and latex microspheres.12 In our case, a silica/silica NOM system was investigated. Nanoparticles were first derivatized with alcohol functions to be further immobilized on the microbead surface through reversible anchoring. The original chemical approach used herein was previously described by our group for diol immobilization on CPG (controlled pore glass) in order to achieve oligonucleotide solid-phase synthesis.13,14 We demonstrated that the linkage formed between silanol and alcohol functions was stable in reagents used for DNA solid-phase synthesis. Furthermore, this linkage was cleaved under mild conditions (water/ methanol at 60 C). We applied this original chemical approach to the immobilization of silica NPs on silica microbeads in a one-step reaction. The resulting colloidal assemblies have been characterized by scanning and transmission electron microscopies (SEM and TEM, respectively) and X-ray photoelectron (XPS) and infrared (IR) spectroscopies. Then, the NOM were introduced into a DNA synthesizer to achieve automated ODN synthesis. Compared to the coupling methods of biomolecules on nanosized particles, our strategy afforded a very simple purification process for the rapidly sedimented assemblies, allowing easy elimination of supernatants. Finally, mild conditions for NP release from microbeads have been determined to obtain NPs functionalized with ODN probes that are well anchored onto the surface.
Experimental Section Material. Aminohexanol, 3-(triethoxysilyl)propyl isocyanate, diethylene glycol dimethyl ether (diglyme), 28% ammonium hydroxide in water (NH4OH), and 1,3-propanediol were purchased from Aldrich (Saint Quentin Fallavier, France). Isopropanol was purchased from SDS (Peypin, France). All reagents were used without further purification. The following ultramild (10) Pacard, E.; Brook, M.; Ragheb, A.; Pichot, C.; Chaix, C. Colloids Surf., B 2006, 47, 176. (11) Seliger, H.; Hinz, M.; Ditz, R.; Koch, M.; Lapido, P.; Margel, S., The support-on support concept for in-situ oligonucleotide synthesis on nanoparticles. Nucleosides, Nucleotides Nucleic Acids 2007, 26, 1167. (12) Lucas, L. J.; Chesler, J. N.; Yonn, J.-Y. Biosens. Bioelectron. 2007, 23, 675. (13) Laurent, A.; de Lambert, B.; Charreyre, M.-T.; Mandrand, B.; Chaix, C. Tetrahedron Lett. 2004, 45, 8883. (14) Laurent, A.; Chaix, C. Org. Process Res. Dev. 2006, 10, 403.
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deprotection nucleoside CE-phosphoramidite synthons were used for ODN synthesis: 50 -dimethoxytrityl-N-phenoxyacetyl-20 -deoxyadenosine,30 -[(2-cyanoethyl)-(N,N-diisopropyl)]-phosphoramidite (Pac-dA-CE phosphoramidite), 50 -dimethoxytrityl-N-acetyl-20 deoxycytidine, 30 -[(2-cyanoethyl)-(N,N-diisopropyl)]-phosphoramidite (Ac-dC-CE phosphoramidite), 50 -dimethoxytrityl-N-pisopropyl-phenoxyacetyl-deoxyguanosine, 30 -[(2-cyanoethyl)-(N, N-diisopropyl)]-phosphoramidite (iPr-Pac-dG-CE phosphoramidite), and 50 -dimethoxytrityl-20 -deoxythymidine,30 -[(2-cyanoethyl)-(N,N-diisopropyl)]-phosphoramidite (dT-CE phosphoramidite). The 2-[2-(4,40 -dimethoxytrityloxy)ethylsulfonyl]ethyl(2-cyanoethyl)-(N,N-diisopropyl)-phosphoramidite (chemical phosphorylation reagent), the [(30 ,60 -dipivaloylfluoresceinyl)-6-carboxamidohexyl]-1-O-(2-cyanoethyl)-(N,N-diisopropyl) phosphoramidite (fluorescein phosphoramidite) and all oligonucleotide synthesis reagents (i.e., activator solution (0.45 M tetrazole in acetonitrile), cap mix A (5% phenoxyacetic anhydride (Pac2O) in tetrahydrofuran (THF)/pyridine), cap mix B (16% methylimidazole/THF), oxidizing solution (iodine (I2) 0.02 M in water/ pyridine/THF), deblocking mix (3% trichloroacetic acid (TCA) in dichloromethane), deprotection solution (0.05 M potassium carbonate (K2CO3) in methanol (MeOH)) and dimethylformamide (DMF, synthesis grade), acetonitrile (CH3CN, synthesis grade) and dichloromethane (DNA synthesis grade) were purchased from Glen Research (Sterling, Virginia)). Silica bead powders (700 nm, 4 μm) and nanoparticles (NPs, 50 nm, 5.17 1013 part mL-1 in DMF suspension) were provided by Nano-H (Lyon, France). VBSTA buffer (sodium diethylbarbiturate (veronal) 30 mM, KCl 30 mM, NaCl 0.2 M, tween 0.1%, BSA 1%, pH 8.5) and VBS buffer (veronal 30 mM, KCl 30 mM, NaCl 0.2 M, pH 8.5) were prepared with products obtained from Sigma-Aldrich (Saint Quentin Fallavier, France). Measurements. 1H and 13C nuclear magnetic resonance (NMR) spectra were recorded on a Bruker 200 spectrometer in CDCl3 at room temperature. The chemical-shift scale was calibrated relative to the tetramethylsilane peak used as a reference. 13 C solid-state NMR spectra were recorded on a Bruker Avance 500 spectrometer with a 4 mm CPMAS probe using either the standard CPMAS experiment at 10 kHz spinning speed with a 2 ms contact time and 2 s relaxation delay, which allows complete relaxation of the protons, or the single-pulse 13C/1H experiment using a 30 pulse angle and a 5 s recycle time. Mass spectrometry analysis was carried out on an Applied Biosystems electrospray (ESI). MALDI-TOF mass spectrometry analyses were performed on a Voyager DE-PRO Applied Biosystems instrument using 3-hydroxy picolinic acid as the matrix (IBCP, Lyon, France). Spectrophotometric titrations were performed with a Uvikon XL spectrophotometer at 498 nm. The Fourier transform infrared (FTIR) spectra were recorded on powder-pressed KBr pellets using a Nicolet 460 spectrometer with a resolution of 4 cm-1. X-ray photoelectron spectroscopy (XPS) measurements were performed on PHI Quantera SXM instrument using a monochromatic Al KR source on an analysis area diameter of ∼200 μm. Dual-beam charge neutralization was used on all specimens. High-resolution spectra of all elements were recorded using 26 or 140 eV pass energy. Under these operating conditions, the measured FWHM values of the Ag 3d5/2 line were 0.55 and 0.92 eV, respectively. The binding energies (BEs) were referenced to the Fermi level of the electron analyzer, and the BE scale was calibrated against Au 4f7/2 and Cu 2p3/2 at 83.96 and 932.62 eV, respectively. Dynamic light scattering (DLS) measurements were carried out with a Zetasizer NanoZS (Malvern Instruments) to determine the hydrodynamic diameter (Dh) and the particle size distribution (PSD) of the NPs, the silica beads, and the NOM assemblies. Scanning electron microscopy (SEM) was performed at an acceleration voltage of 15 kV (Hitachi S800, Centre Technologique des Microstructures - Lyon 1, Villeurbanne, France). Transmission electron microscopy (TEM) pictures were obtained at an acceleration voltage of 80 kV (Philips CM 120, Centre Technologique Langmuir 2010, 26(7), 4941–4950
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Table 1. Different Ratios (R) of Nanoparticles per Silica Bead Investigated in This Study NS
NL
Ra
2.48 1013 1.41 109 17 600 2.40 109 34 500 8.27 1013 7.81 109 274 000 7.70 1014 a R = NS/NL, where NS and NL are the numbers of small silica particles and large silica beads in the suspension, respectively.
des Microstructures - Lyon 1, Villeurbanne, France). Diluted solutions of silica NPs, silica beads, and NOM assemblies were examined after deposition on a Formvar-coated copper grid and evaporation to dryness. Conventional microscopy fluorescence images were carried out on a Zeiss Axioplan 2 imaging apparatus equipped with a 100 1.3 oil-immersion objective and a camera. Samples were observed by both fluorescence and transmitted light to locate the particles. Fluorescein was excited by a laser with a 450-490 nm band-pass filter, and fluorescence from the sample was collected with a 515-565 nm band-pass filter. Chemiluminescence was measured with a CCD camera (Intelligent Dark box II, Fuji Film; Image Gauge 3.12, Fuji Film).
Nanoparticle Silanization. Synthesis of 3-(Triethoxysilyl)propyl-hydroxyhexyl Urea (TESPHU). 6-Amino-1-hexanol (1.25 mmol, 146 mg) was first dissolved in 2.5 mL of isopropanol. After the addition of 3-(triethoxysilyl)propyl isocyanate (1.25 mmol, 310 μL), the mixture was stirred at room temperature for 30 min. An aliquot of this solution was evaporated to dryness under reduced pressure and characterized by 1H NMR and ESI mass spectrometry (Figure S1, Supporting Information). 1 H NMR (200 MHz, CDCl3, δ): 0.59 (t, 2H, Si-CH2), 1.19 (t, 9H, 3 CH3), 1.35-1.60 (m, 10H, 5CH2), 3.10-3.20 (m, 4H, 2CH2N), 3.64 (t, 2H, CH2-OH), 3.76-3.87 (q, 6H, CH2OSi), 4.82-4.96 (2t, 2H, 2NH). MS (ESI, m/z): [M þ H]þ calcd for C16H36N2O5Si, 364.6; found, 365.0; [M þ Na]þ 387.2. Silanization of the 50 nm Silica Nanoparticles. A crude solution of TESPHU (563 μL, 0.5 mol L-1) was added to a suspension of NPs (200 mg) in 20 mL of DMF, and the mixture was stirred slowly at 120 C for 16 h. The suspension was centrifuged (22 400g, 1 h), and the supernatant was eliminated. Then, the particles were washed via three centrifugation/redispersion cycles in DMF. For each wash, the sonicator was used to redisperse the NPs completely. In addition, the samples analyzed by 13C solid-state NMR were redispersed in a DMF/diglyme mixture and heated overnight at 90 C to mimic the conditions of NOM elaboration. Hydroxyl-Function Titration on Nanoparticles. Nanoparticles (50 mg) were dried for 1 h at 120 C and then coevaporated twice to dryness with dry acetonitrile. Dry acetonitrile (0.5 mL) was added to the flask under argon. dT-CE phosphoramidite (5 mg) was dissolved in dry acetonitrile (67 μL), and then 14 μL of activator solution was added. This mixture was added to the suspension of NPs and stirred at room temperature for 1 h. An oxidizing solution (306 μL) was introduced, and the mixture was stirred for 2 additional hours. The NPs were centrifuged (15 000g, 30 min), washed via three centrifugation/redispersion cycles in acetonitrile, and evaporated to dryness under reduce pressure. Then, 5 mL of a deblocking mix was added to release the dimethoxytrityl (DMT) group from the nucleotide synthon, and the solution became orange. Absorbance at 498 nm was measured for DMT group quantification (ε = 70 mL μmol-1 cm-1). Elaboration of the NOM Assemblies. The surface of the silica beads (700 nm or 4 μm) was hydrated by sonication in water (1 h, 20 C) prior to the grafting reaction. Water was then eliminated by centrifugation and replaced with diglyme. Then, the suspension of silanol-activated micrometric beads in diglyme was added to the suspension of alcohol-functionalized NPs in DMF at room temperature. The best grafting was obtained with a global concentration of silica of 6 mg mL-1 in a solution of DMF/ diglyme (1/4 v/v). The mixture was slowly stirred and held at 90 C Langmuir 2010, 26(7), 4941–4950
overnight. The obtained NOM assemblies were separated from the nonadsorbed NPs by sedimentation. The nanoparticle grafting on the beads was then stabilized by a dehydration process achieved by heating the NOM for 3 h in an oven at 120 C. As depicted in Table 1, different ratios R of small silica nanoparticles (NS) per large silica bead (NL) were investigated. A slightly different procedure was used for the preparation of the model system used for the solid-state NMR study. OH-functionalized NPs were replaced by 1,3-propanediol in the mixture. A solution of 6 M 1,3-propanediol in diglyme was prepared by adding 1,3-propanediol (6 mmol, 437 μL) in 563 μL of diglyme. The surfaces of the silica beads (300 mg) were hydrated by sonication 1 h prior to the grafting reaction. Water was then eliminated by centrifugation and replaced by 1 mL of diglyme. Then, the 1,3-propanediol solution was added to the suspension of silanol-activated microbeads. The mixture was slowly stirred and held at 90 C overnight. Then, the particles were washed by three centrifugation/redispersion cycles in diglyme. Finally, the microbeads were dried at 120 C for 16 h. Functionalized microbeads were analyzed by 13C solid-state NMR. Purification was achieved by alternately sedimenting and resuspending the obtained NOM assemblies in diglyme. One can note that the purification of the reaction medium can be quite tricky when nanoparticle aggregates and expected NOM assemblies are both present in the solution because centrifugation or sedimentation does not allow one to discriminate between the two species. Consequently, the experimental conditions must be optimized to avoid nanoparticle aggregate formation. Then, after a few minutes, NOM assemblies sediment in diglyme when nanoparticles stay in suspension. Stability Studies of the NOM Assemblies. The stability of the NOM assemblies (50 nm NPs/700 nm beads) was first assessed in various solvents (water, ammonium hydroxide, and deprotection solution) at 20 and 60 C using dynamic light scattering (DLS). Nanoparticle release from the assemblies was followed by the appearance of a second population that corresponds to free NPs. NOMs were considered to be stable until the second population was observed. The NOM stability was then evaluated in the reagents used in ODN synthesis. Scanning electron microscopy (SEM) analyses were carried out on the 50 nm/4 μm NOM assembly. The material was stirred in 500 μL of each ODN synthesis reagent (activator, cap mix A, cap mix B, oxidizing solution, and deblocking mix) in separate vials for 1 h. Then, the samples were washed with dry acetonitrile. After treatment, the resulting materials were dried under reduced pressure and characterized by SEM. Finally, the stability of the NOM assembly during ODN automated synthesis was checked. With the aim of reproducing the conditions of a 23-mer synthesis, NOM assemblies (50 nm NPs/4 μm beads) were introduced into the synthesis column and all of the reagents (except for the nucleoside CEphosphoramidite solutions in acetonitrile that were replaced by pure acetonitrile) were injected into the column by running the instrument’s 1 μM phosphoramidite program for a 23-mer ODN elongation. ODN Synthesis. Oligonucleotides were synthesized using an Applied Biosystems 394 RNA/DNA synthesizer (Applied Biosystems, Foster City, CA) using the standard protocol (1 μmol scale coupling program). PTFE filters with 0.2 mm thickness and 1 μm porosity were used to close the synthesis columns (RothSochiel, Lauterbourg, France). NOM material (50 mg) was introduced into each column. A 23-mer HBV (hepatitis B 0 virus) probe (50 ATC TCG GGA ATC TCA ATG TTA GT3 ) was synthesized from alcohol functions available at the surface of the NOM assemblies. This sequence corresponds to the capture probe of the HBV DNA ELOSA (enzyme-linked oligosorbent assay) test developed by bioMerieux.15 After the first and last nucleotide incorporations, DMT groups were released from the (15) Minard-Basquin, C.; Chaix, C.; Pichot, C.; Mandrand, B. Bioconjugate Chem. 2000, 11, 795.
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Figure 1. Stepwise process developed in this study. Step 1: nanoparticle functionalization by TESPHU. Step 2: grafting of hydroxylated nanoparticles onto native silica beads. Step 3: oligonucleotide (ODN) solid-phase synthesis. Step 4: release of the ODN-functionalized nanoparticles. support with a deblocking mix solution and titrated at 498 nm (ε = 70 mL μmol-1 cm-1). Nucleobase deprotection was achieved in deprotection solution for 4 h at 20 C under gentle stirring. The stability of the assemblies after ODN synthesis was controlled by SEM. With the aim of confirming the presence of ODN on the NOM 0 assemblies, we synthesized fluorescein-ODN0 (5 fluorescein-ATC TCG GGA ATC TCA ATG TTA GT 3 ). After 4 h in the deprotection solution, fluorescence was observed by optical fluorescence microscopy. Then, nanoparticle release was achieved in 1% NH4OH in water for 1 h at 60 C and characterized by transmission electron microscopy (TEM). The quality of the ODN sequence synthesized from the NOM assemblies was controlled by MALDI-TOF mass spectrometry. To do so, it was necessary to synthesize an ODN that could be released from the nanoparticles. A specific baso-cleavable derivative (chemical phosphorylation reagent) was coupled to the NPs prior to initiating the HBV ODN synthesis. This derivative was cleaved during the deprotection step (16 h in NH4OH 28%/ water) to liberate an ODN-30 -phosphate in solution (ATC TCG GGA ATC TCA ATG TTA GT-PO32-). After filtration to eliminate the microbeads and concentration to dryness, the crude solution was analyzed by MALDI-TOF MS ([M þ H]þ calcd, 7134.6; found, 7134.8 and [M þ H]2þ calcd, 3567.8; found, 3567.8) (Figure S8, Supporting Information).
Control of the DNA Capture Efficiency by the ODNs Grafted onto the NOM Assemblies. The accessibility and selectivity of the ODNs grafted onto the silica NOM assemblies were evaluated using a chemiluminescent (CL) assay. The ODNgrafted assemblies were first saturated for 20 min at 37 C in VBSTA buffer. After washing, 100 μL of either a HBV biotinylated complementary or noncomplementary target (100 nM) was added, and the suspension of NOM assemblies was incubated for 1 h at 37 C under gentle stirring. After being washed in VBS buffer, the hybridized assemblies were saturated with VBSTA buffer and incubated at 37 C with a solution of HRP-labeled streptavidin. Finally, a CL measurement solution composed of luminol, hydrogen peroxide, and p-iodophenol was added to the NOM assemblies, and the emitted light was integrated for 10 s using a CCD camera (Intelligent Dark box II, Fuji Film; Image Gauge 3.12, Fuji Film).
Results and Discussion As mentioned in the Introduction, the main aim of this work is the automated synthesis of ODN probes at the surface of OH-functionalized silica NPs (close to 50 ( 5 nm). However, ODN automated synthesis relies on the use of solid supports that have to be entrapped in synthesis columns, and to date, there are no commercially available filters displaying such a low porosity 4944 DOI: 10.1021/la903572q
(50 nm) and ability to resist the reagents and pressure used in a DNA synthesizer. Our strategy to solve this problem was to immobilize the nanoparticles onto a support that is able to retain them within the columns of the DNA synthesis instrument. The bond between the silica nanoparticles and the support should both resist the ODN synthesis reagents and be reversible, allowing ultimate nanoparticle release. To achieve this, silica beads were used as the solid support and hydroxyl-functionalized 50 nm silica colloids were used as the nanosized particles. The strategy developed in this work is depicted in Figure 1. First, the surface of the NPs was functionalized with alcohol groups through silanization with 3-(triethoxysilyl)propyl-hydroxyhexyl urea (TESPHU). Second, the functionalized NPs were anchored to the surfaces of the silica microbeads. The assemblies obtained were then used as a solid support for the automated ODN synthesis. Finally, ODN-functionalized NPs were released from the microbeads. Each step in the process is detailed below. Functionalization of Silica Nanoparticles with TESPHU. The functionalization of the NPs with hydroxyl arms was achieved by silanization using 3-(triethoxysilyl)propyl-hydroxyhexyl urea (TESPHU). This reagent was prepared from commercial 3-(triethoxysilyl)propyl isocyanate and 6-amino-1-hexanol, as detailed in the Experimental Section. The product obtained was characterized by 1H NMR and ESI MS (Figure S1, Supporting Information) and used without further purification. TESPHU was then added to the DMF suspension of NPs (50 nm, 10 mg mL-1), and the mixture was held at 120 C for 16 h under gentle stirring. The obtained NPs were exhaustively characterized to show the grafting of TESPHU onto the surface of the silica NPs. First, XPS analyses were performed for TESPHU, bare silica NPs, and OHfunctionalized NPs. In the case of TESPHU (Figure S2, Supporting Information), the C 1s spectrum showed one broad peak that could be curve fit with three peaks. The main peak at 285 eV was assigned to the carbons in the aliphatic chain (C-H, C-C), the second peak at 286.4 eV was assigned to the carbon-nitrogen (C-N) and/or carbon-oxygen (C-O) bond, and the third one at 288.8 eV was assigned to the carboxylate carbon (OdC-N2) of the urea function. The same analysis was then carried out on bare and OH-functionalized NPs. The subtraction of the C 1s spectrum of the bare NPs from that of the OH-functionalized NPs provided a signal that could be curve fit with the three distinctive peaks of TESPHU at 284.6, 286.7, and 288.8 eV, clearly evidencing the successful silanization of the NPs. In addition, the peak corresponding to the urea function was not present for the bare NPs. Because XPS is a surface-analysis technique (to a depth of ca. 10 nm), these results attest to the presence of the OH functions at the surfaces of the NPs. These results are supported by infrared spectroscopic analyses carried out for TESPHU and functionalized nanoparticles. The characteristic bands of the TESPHU alkyl arm were clearly observed on the OH-functionalized NPs. Indeed, the bands at 2950-2900, 1650, 1400, and 680-520 cm-1 correspond to the aliphatic ν C-H, ν CdO, δ C-H, and δ CH2, respectively (Figure S3, Supporting Information). Further insight into the silanization of the silica NPs by TESPHU was obtained via 13C solid-state NMR spectroscopy (Figure 2). To confirm the success of the silanization and the stability of the hydroxyl arm during NOM preparation, 13C solidstate NMR spectroscopy was carried out on OH-functionalized NPs after treatment under conditions used for NOM preparation (stirring at 90 C overnight in DMF/diglyme 1/4 v/v). The comparison of the spectra of TESPHU in CDCl3 and OHfunctionalized NPs in the solid state clearly indicates (i) the Langmuir 2010, 26(7), 4941–4950
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13
C NMR spectra of (A) TESPHU (CDCl3) and (B) OH-functionalized NPs (solid state, CPMAS).
success of the silanization and (ii) the stability of OH functionalization under the experimental conditions of NOM preparation. Finally, the number of accessible OH groups was estimated by dimethoxytrityl (DMT) titration (Experimental Section). Up to 1100 OH groups were quantified on the surface of each functionalized 50 nm particle (about 0.14 OH function nm-2). The incorporation of primary alcohols by silanization with TESPHU led to the good reactivity of NPs toward the dT-CE phosphoramidite synthon used during ODN solid-phase synthesis. We assumed that the main part of these OHs can initiate ODN synthesis after nanoparticle loading on the microbeads. Because the silanization of the NPs was successful, the next step consisted of the assembly of NPs onto silica microbeads. Elaboration of the NOM Assemblies. The goal of the assembly process was to graft a nanoparticle monolayer onto the beads. To do so, micrometric silica beads needed to exhibit a hydrated surface to (i) increase the number of silanols and (ii) facilitate anchoring of the hydroxylated nanoparticles, probably through hydrogen bonding interactions (Figure 3A). Indeed, considering the second point, we assumed that NP anchoring was favored when H bonds existed between the hydroxylated compound and the silica support before further evolving to covalent Si-O-C linkages once the mixture was heated (90 C, overnight). The preliminary activation of the microbeads was achieved by sonication in an aqueous solution (Experimental Section). These optimized conditions resulted from previous studies reported by our group on oligoethylene glycol grafting onto a CPG support.13 In this work, we observed that the water adsorbed on the silica Langmuir 2010, 26(7), 4941–4950
surface has a predominant influence on the grafting reaction.14 Temperature was also a key parameter in the reaction. The optimized temperature was determined to be between 80 and 120 C. In this work, the NP anchoring temperature was established at 90 C. This temperature allows partial dehydration of the microbeads during the reaction, favoring alcohol/silanol coupling. The number of NPs that could adsorb on the microbeads was estimated by dividing the microbead surface area by the crosssectional area of a plane bisecting a nanoparticle.16 In our case, one 4 μm silica bead could theoretically be covered with 25 600 NPs. This estimated value is not far from that given by the theory,17 which assumes hexagonal close-packing of NPs onto a planar surface (eq 1) pffiffiffi N sat ¼ 2π= 3ð1 þ rL =rs Þ2 ¼ 23 800
ð1Þ
where rL is the radius of the microbeads, rS is the radius of the nanoparticles, and Nsat is the maximum number of NPs per microbead required to form a monolayer. Our experiments confirmed experimentally that a low R value (R = 17 600, Table 1) leads to incomplete microbead coverage. In contrast, many nanoparticle aggregates are observed together with low coverage of the NOM assemblies for a high R value (R g 274 000). (16) Fleming, M. S.; Mandal, T. K.; Walt, D. R. Chem. Mater. 2001, 13, 2210. (17) Ottewill, R. H.; Schofield, A. B.; Waters, J. A.; Williams, N. S. J. Colloid Polym. Sci. 1997, 275, 274.
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Figure 3. (A) Grafting reaction of OH-functionalized nanoparticles onto silica beads. (B) Schematic representation of the obtained NOM assembly.
Figure 4. SEM pictures of NOM assemblies (A) after grafting and (B) after 1 h in each ODN synthesis reagent successively.
The best compromise was finally achieved with R = 34 500, and the grafting efficiency was first assessed by examining the morphology of the assemblies obtained by scanning electron microscopy (Figure 4A). The SEM pictures showed an efficient coverage of microbeads with no nanoparticle aggregates detected. However, the packing appeared to be incomplete and less uniform than the packing that can be obtained with other kinds of 4946 DOI: 10.1021/la903572q
interaction, such as electrostatic interaction18,19 or specific chemical (amine-aldehyde)16 or biochemical (streptavidin-biotin).16,20 In these systems, the surface functionality (charges, reactive groups, and biomolecules) prevents any side interaction between particles of the same kind, which is not the present case. The incomplete, less uniform surface coverage observed in our system may be ascribed to some aggregation of the NPs prior to their assembly with silanol-activated microbeads. Aggregation may occur via some hydrogen-bond-type interaction between the NPs carrying both alcohol and (to a lesser extent) silanol functions. Despite the lesser selectivity of H-bonding-driven assembly, the silica-based assemblies obtained displayed the desired morphology (i.e., a micrometric bead carrying hydroxylated NPs for later use in ODN solid-phase synthesis). Besides, the residual silanol groups on the surfaces of the microbeads should not interfere with ODN synthesis, which is more selective toward primary alcohol groups. This has been confirmed by DMT titration after the dT-CE phosphoramidite grafting reaction on both bare 4 μm silica beads and NOM. Ten milligrams of each material was incorporated into the instrument column, and one coupling cycle was run with the dT synthon. DMT quantification was directly performed on the synthesizer. OH groups (2.1 μmol/g) were quantified by this method for the NOMs. The control achieved on the bare 4 μm silica beads indicated 0.34 μmol/g. This value corresponds to the highest amount of DNA that could be synthesized from the bare 4 μm silica support. In contrast, nanoparticles efficiently cover a large part of the microbead surface in the case of NOMs; consequently, the extent of these undesirable DNA syntheses is certainly much lower. The formation of NOM assemblies was also proven by spectroscopic techniques. First, XPS analyses of the NOM assemblies confirmed the presence of the organic arm coming from TESPHU on the NOM surface (Figure S2, Supporting Information). Indeed, the broad C 1s peak could be curve fit with the same peaks as those detectable for TESPHU and OH-functionalized NPs (284.9, 286.4, and 288.5 eV). These results indicate the successful grafting of the NPs onto the beads. (18) Lawrie, G.; Grondahl, L.; Battersby, B.; Keen, I.; Lorentzen, M.; Surawski, P.; Trau, M. Langmuir 2006, 22, 497. (19) Lansalot, M.; Sabor, M.; Elaissari, A.; Pichot, C. Colloid Polym. Sci. 2005, 283, 1267. (20) Wang, L.; Yang, C.; Tan, W. Nano Lett. 2005, 5, 37.
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Figure 5. Principle of automated oligonucleotide synthesis. Summary of the different steps, reagents, and reaction times of one coupling cycle and NOM stability in these reagents.
According to the literature,21 the link between the NPs and the microbeads was expected to be covalent. To show the formation of a Si-O-C bond resulting from the reaction of the silanol with the OH-functionalized NPs, 13C solid-state NMR experiments were carried out. Indeed, the resonance of the R carbon in the OH function was expected to be significantly different once covalent grafting was achieved (Figure 3B, see the two methylene in red). Unfortunately, the measurements did not bring to light any conclusive information because the amount of the organic compound was too low in the NOM assemblies. Nevertheless, we had the beginnings of an answer by grafting model compound 1,3-propanediol onto the silica beads using the same conditions as for the NOM elaboration (DMF/diglyme, 90 C overnight). The obtained 13C NMR spectrum (Figure S4, Supporting Information) showed broad peaks (FWHM > 100 Hz), indicating covalent grafting of the diol (physically adsorbed species would exhibit narrower peaks). The curve fitting of this spectrum22 and notably of the peak at 37 ppm that can be unambiguously attributed to the midchain methylene of 1,3-propanediol (HO-CH2-CH2-CH2-OH) showed two broad peaks (112 and 254 Hz, respectively), indicating that 1,3-propanediol was likely grafted through either one (monodentate) or two (bidentate) OH groups. This preliminary study is in favor of a covalent linkage between the NPs and the (21) Zaborski, M.; Vidal, A.; Ligner, G.; Balard, H.; Papirer, E.; Burneau, A. Langmuir 1989, 5, 447. (22) Massiot, D.; Fayon, F.; M., C.; King, I.; Le Calve, S.; Alonso, B.; Durand, J.-O.; Bujoli, B.; Gan, Z.; Hoatson, G., Modelling one and two-dimensional solidstate NMR spectra. Magn. Reson. Chem. 2002, 40, 70.
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beads, probably driven by H bond interactions and catalyzed by temperature. Finally, FTIR analyses of the NOM showed bands at 2950-2900, 1650, and 680-520 cm-1 characteristic of the organic arm of TESPHU, confirming the presence of OH-functionalized NPs in the assembly (Figure S3B, Supporting Information). Solid-Phase Oligonucleotide Synthesis from Silica NOM Assemblies. The principle of solid-phase oligonucleotide synthesis is based on the successive incorporation of nucleotides in a growing chain that is anchored to a solid support. Each nucleotide addition follows the reaction cycle detailed in Figure 5. Before going any further, the stability of the NOM assemblies in the solutions used for ODN synthesis was evaluated by DLS analysis. Because of sedimentation issues that prevent this analysis on micrometric systems, the study was carried out using 700 nm silica beads. For a constant number of silica beads, the specific surface area available for grafting is notably different between 700 nm and 4 μm beads and R was thus optimized at 1700 for this new system (Nsat = 800). DLS analysis of the NOM assemblies in DMF showed only one size distribution centered at 1 μm (Figure 6A). Incubation of the same assemblies in either the activator solution (90 min) or the oxidizing solution (60 min) did not induce any destabilization (Figure S5, Supporting Information). This was confirmed by SEM, which showed no noticeable difference in the assemblies’ morphology before and after treatment (data not shown). The stability of the NOM assemblies elaborated with the 4 μm beads after successive 1 h treatments in each synthesis reagent (activator solution, cap mix A, cap mix B, oxidizing solution, and DOI: 10.1021/la903572q
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Figure 7. Fluorescence imaging of the NOM assemblies grafted with HBV probes bearing a fluorescein tag: (A, C) fluorescence and (B) transmitted light signal. Figure 6. Dynamic light scattering analyses (number distribution). (A) NOM assemblies (using 700 nm silica beads) and nanoparticles in DMF (superimposed distributions). After incubation of NOM in (B) 28% NH4OH/H2O at 20 C, (C) 28% NH4OH/H2O at 60 C, (D) deprotection solution (0.05 M K2CO3/MeOH), (E) H2O at 20 C, and (F) H2O at 60 C.
deblocking mix) was then evaluated using SEM (Figure 4B). A majority of the nanoparticles were still grafted onto the microbeads after the various treatments, confirming that NOM assemblies were stable under ODN synthesis conditions. The stability of the 4-μm-based NOM assemblies was further tested inside the automated ODN synthesizer. Indeed, mechanical stirring and pressure can potentially lead to the release of nanoparticles. The synthesis of a 23-mer ODN was simulated by introducing NOM and all of the reagents into the synthesis column, except for the nucleoside CE-phosphoramidite solutions in acetonitrile that were replaced by pure acetonitrile, and running the appropriate synthesis program. The SEM pictures of the assemblies after this treatment confirmed their stability against the pressure and temperature used in the standard ODN synthesis protocol (Figure S6, Supporting Information). The last step of the ODN synthesis procedure usually consists of the deprotection of the nucleic bases and phosphates by basic treatment. With classical CE-phosphoramidites, assemblies must be stable under the strongly alkaline conditions required to remove the nucleic base protecting groups (28% NH4OH in water, 60 C, 16 h). However, the cohesion of our NOM assemblies relies on Si-O-C bonds that can be easily broken under such conditions. The ODN-functionalized NPs must not be released at this stage of the process. The stability of the assemblies in these solvents was followed by DLS measurements (Figure 6). It should be noted that the peak intensities recorded by this technique and indicated by number percentage are related to the number of particles present in the solution. Consequently, the peak areas of both 50 nm NP and 700 nm bead populations on DLS profiles could not be compared directly because the number of small particles was much higher than the number of microbeads. Nevertheless, we confirmed 4948 DOI: 10.1021/la903572q
that the assemblies were not stable in 28% NH4OH/H2O at 60 C. Serious degradation was observed, even after only 20 min in this medium (Figure 6C). At 20 C, the stability was not sufficient to perform complete oligonucleotide deprotection after synthesis (Figure 6B). Consequently, we chose a set of phosphoramidite monomers that met the desired criteria for ultramild deprotection (i.e., phenoxyacetyl (Pac)-protected dA and 4-isopropyl-phenoxyacetyl (iPr-Pac)-protected dG, along with acetyl-protected dC). The protecting groups were eliminated in a solution of 0.05 M K2CO3 in MeOH for 4 h at 20 C. DLS analyses (Figure 6D) revealed that the assemblies were stable under these ultramild deprotection conditions. The base protecting groups can be completely removed without any destabilization of the assemblies. Having demonstrated the stability of the NOM assemblies under ODN synthesis conditions, the materials were then evaluated as potential solid supports for automated ODN synthesis. Current procedures require the entrapment of the solid support in the synthesis column. In our case, the assemblies were kept inside the columns using a PTFE filter of 1 μm porosity. Indeed, preliminary tests showed that the NOM support based on the 4 μm silica beads was efficiently trapped without obstructing the filters (which was not the case with the assemblies formed with the 700 nm beads). Hydroxyl groups available on the surface of the grafted NPs were used to initiate oligonucleotide synthesis by attaching the first0 nucleoside of the ODN 30 -end (Figure 5). The 0 synthesis of the (5 ATC TCG GGA ATC TCA ATG TTA GT3 ) sequence, which corresponds to the capture probe of the HBV (hepatitis B virus) diagnostic assay developed by bioMerieux, was successfully performed.15 DMT titration after the first and the last nucleotide incorporations clearly showed that no NP product was lost during ODN synthesis. In this experiment, 2.5 μmol of DMT/g of NOM was quantified after the first dT phophoramidite incorporation and 2.05 μmol of DMT/g of NOM was measured after the last nucleotide incorporation of the ODN synthesis (dA phosphoramidite coupling at the 50 end of the ODN). These data allow the calculation of an average nucleotide phosphoramidite coupling yield per cycle of 99% during ODN solid-phase Langmuir 2010, 26(7), 4941–4950
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Figure 9. TEM pictures of the nanoparticles carrying ODN HBV Figure 8. Hybridization with HBV (hepatitis B virus) probe ODN-functionalized NOM assemblies. (A) Schematic representation of the hybridization assay. (B) Luminescence signal for complementary and noncomplementary targets.
synthesis. Considering this coupling yield, the purity of the final 23-mer oligonucleotide was estimated to be ∼70% in the crude material. TEM pictures further proved that neither the presence of ODN probes nor the deprotection step had a noticeable influence on the NOM stability (Figure S7, Supporting Information). This was also confirmed by SEM, which produced fuzzy pictures likely related to the presence of ODNs on the NOM surface. Nevertheless, these observations do not completely rule out the possible release of a few NPs from the microbeads. To further prove the presence of ODNs on the NPs, an HBV probe labeled with fluorescein at the 50 end was synthesized. Fluorescein was introduced at the end of ODN synthesis. The ODN-functionalized NOM assemblies obtained after the deprotection step (4 h in 0.05 M K2CO3/MeOH) were observed by optical microscopy. A fluorescent ring around the edge of the NOM was clearly distinguishable, indicating the effective anchoring of HBV probes onto the surface of NPs supported by the micrometric beads (Figure 7). To check the nature of the oligonucleotides synthesized from the NOM assemblies, a baso-cleavable monomer, referred to as the chemical phosphorylation reagent in the Experimental Section, was grafted onto the OH groups by automated synthesis before subsequent HBV 23-mer ODN synthesis. During treatment in 28% aqueous NH4OH for 16 h at 60 C, this particular monomer described by Urdea and Horn23 was able to release ODN-30 -phosphate into solution. Obviously, NPs were also released during the treatment, but this did not interfere with the mass spectrometry characterization of the crude ODN solution. MALDI-TOF mass spectrometry analysis confirmed that the main oligonucleotide in the crude solution corresponded to the expected 23-mer ODN-30 phosphate (Figure S8, Supporting Information). ODN-functionalized NOM assemblies were finally evaluated for the specific capture of single-stranded DNA fragments using a chemiluminescent (CL) assay (Figure 8A). This threestep process relies on (1) hybridization between HBV probe-NOM assemblies and a cDNA target bearing a biotin group, (2) the addition of a (streptavidin-horseradish peroxidase (HRP)) conjugate that specifically binds to biotin, and (3) the addition of a CL measurement solution containing all of the enzyme substrates. The light emitted during this reaction was measured using a CCD camera. This test was performed with complementary and noncomplementary (23-mer random sequence) biotin DNA targets. Figure 8B clearly shows that (23) Urdea, M. S.; Horn, T. Tetrahedron Lett. 1986, 27, 2933.
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probes.
the test was positive only with the complementary target. This result confirms the good accessibility of the probes present on the NOM assemblies for specific hybridization with the DNA target. Release of the Nanoparticles from Their Solid Support. The ultimate goal of this work is to ensure the release of the ODNfunctionalized NPs from their micrometric colloidal supports. Different conditions (solvent and temperature) were tested. As mentioned above in the text, DLS analysis of the assemblies before incubation in various solutions showed a single peak centered at 1 μm corresponding to the nanoparticle-bead NOM assemblies. After incubation in a medium such as water or aqueous ammonia, a second peak appeared, centered at 100 nm and corresponding to the NPs (Figure 6B,C,E,F). Another set of experiments showed the importance of temperature in the release kinetics. Although NPs appeared after 48 h at 20 C in water, only 90 min was necessary at 60 C (Figure 6F). To conclude, two parameters accelerated the NP release from NOM: ammonia concentration and temperature. Finally, to confirm the reversibility of the grafting, HBV probe-NOM assemblies were treated for 2 h in water at 60 C. Fast sedimentation of the beads occurred in 80 C. After optimization of the experimental conditions, well-defined nano-on-micro assemblies were obtained and this material was sufficiently stable during the oligonucleotide synthesis. Another interesting part of this work concerned the stability of the assemblies during the deprotection step following ODN synthesis and the subsequent release of the ODN-functionalized NPs under specific treatment (1% NH4OH in water for 1 h at 60 C). Our method provided silica NPs that were well functionalized with oligonucleotides, DOI: 10.1021/la903572q
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as demonstrated by hybridization experiments achieved with the cDNA target. Capillary electrophoresis experiments are now underway to optimize the detection of the target DNA sequence. In the context of miniaturization processes, the ultimate goal of the project presented here concerns the development of a lab-ona-chip diagnostic test to allow direct nucleic acid (DNA or RNA) multidetection without sample labeling. The final aim is to be able to use NOM assemblies based on luminescent nanoparticles and micrometric magnetic silica beads. After oligonucleotide synthesis, the resulting ODN-functionalized assemblies will enable DNA extraction and concentration through their magnetic properties and direct, label-less detection by luminescence, permitting the specific coding of various DNA targets. This final part of the project is under investigation in the laboratory. Acknowledgment. This work was supported by the Agence Nationale de la Recherche and the Lyonbiop^ole. We thank M. Becchi (IBCP, IFR 128 BioSciences Lyon Gerland) for mass spectrometry analyses, A. Baudouin and F. D’Agosto
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(UMR 5265, C2P2) for their help with NMR analyses, and C. Grossiord (Science et Surface S. A.) for XPS analyses. Supporting Information Available: Characterization of TESPHU. C 1s core spectra of TESPHU, and OH-functionalized silica nanoparticles. C1 spectrum subtraction of bare NPs from OH-functionalized NPs and of silica NOM assemblies. Infrared spectroscopic analyses of TESPHU and functionalized nanoparticles and TESPHU, microbeads, and NOM assemblies. 13C solid-state NMR spectrum of the propanediol-functionalized microbeads (model system) and corresponding curve fitting. DLS analyses of NOM assemblies after incubation in ODN synthesis reagents. SEM picture of NOM assemblies after a simulated 23-mer ODN synthesis without the nucleoside phosphoramidites in the synthesizer. TEM pictures of the NOM assemblies bearing HBV probe ODNs. MALDI-TOF MS of the 23-mer ODN 30 phosphate. TEM analyses of microbeads after treatment in 1% NH4OH/H2O for 1 h at 60 C. This material is available free of charge via the Internet at http://pubs.acs.org.
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