Deposition of DNA-Functionalized Gold Nanospheres into

Angelika Niemz, Krisanu Bandyopadhyay, Eric Tan, Kitty Cha, and Shenda M. Baker ... Guping He , Thomas M. Bennett , Mohammad Alauhdin , Michael W. Fay...
0 downloads 0 Views 1014KB Size
Download PDF
4978

Langmuir 2006, 22, 4978-4984

Deposition of DNA-Functionalized Gold Nanospheres into Nanoporous Surfaces Krisanu Bandyopadhyay,†,§ Eric Tan,† Lin Ho,† Sarah Bundick,‡ Shenda M. Baker,*,‡ and Angelika Niemz*,† Keck Graduate Institute, Claremont, California 91711, and Department of Chemistry, HarVey Mudd College, Claremont, California 91711. ReceiVed December 23, 2005. In Final Form: March 16, 2006 We report the deposition of DNA-conjugated gold nanospheres into arrays of surface nanopores obtained from hexagonally ordered thin polystyrene-b-poly(methyl methacrylate) (PS-b-PMMA) diblock copolymer films on silicon. The deposition occurs spontaneously from aqueous solution and is driven by either electrostatic interactions or specific DNA hybridization events between the DNA nanospheres and the surface nanopores. To mitigate this spontaneous deposition, we have chemically modified the nanopores with either positively charged aminosilanes or oligonucleotide probe sequences. The deposition of DNA nanospheres into the surface nanopores was characterized by atomic force microscopy (AFM) and X-ray photoelectron spectroscopy (XPS). We have observed preferential immobilization of individual DNA nanospheres within the nanopores, based on the size matching between the two entities. The inclusion density and selectivity of DNA nanosphere deposition into the surface nanopores was found to depend predominantly on the methods through which the nanoporous surfaces were prepared and chemically functionalized.

Introduction Bio-inspired bottom-up self-assembly can be used to fabricate complex, nanoscopically ordered structures in a rapid, parallel manner using minimal instrumentation.1-6 In contrast, top-down nanoscopic surface patterning through photolithography is limited by the wavelength of light and the materials currently used, and fabricating features e70 nm imposes significant challenges.7 Smaller feature sizes can be fabricated by direct-write processes such as e-beam lithography8 and dip-pen nanolithography;9 however, these processes do not allow for rapid, parallel nanoscale patterning of entire macroscopic surfaces and mass production of nanoscale devices.10 The goal of this study is to demonstrate the feasibility of spontaneous deposition of individual DNAfunctionalized gold nanospheres into arrays of appropriately functionalized nanopores on a silicon surface, mediated through nonspecific electrostatic interactions or through specific biomolecular recognition based on DNA hybridization. DNA-functionalized gold nanospheres are versatile nanoscopic building blocks that have been used in a variety of bioassay and * Corresponding author. (A.N.) Address: Keck Graduate Institute, 535 Watson Drive, Claremont, CA 91711; phone: (909) 607-9854; e-mail: [email protected]. (S.M.B.) Address: Department of Chemistry, 301 East 12th Street, Harvey Mudd College, Claremont, CA 91711; phone: (909) 621-8643; e-mail: [email protected]. † Keck Graduate Institute. ‡ Harvey Mudd College. § Current address: Department of Natural Sciences, University of Michigan-Dearborn, 4901 Evergreen Road, Dearborn, MI 48128-1491. (1) Storhoff, J. J.; Mirkin, C. A. Chem. ReV. 1999, 99, 1849-1862. (2) Zhang, S. G. Nat. Biotechnol. 2003, 21, 1171-1178. (3) Niemeyer, C. M. Curr. Opin. Chem. Biol. 2000, 4, 609-618. (4) Liz-Marzan, L. M.; Mulvaney, P. J. Phys. Chem. B 2003, 107, 7312-7326. (5) Daniel, M. C.; Astruc, D. Chem. ReV. 2004, 104, 293-346. (6) Parak, W. J.; Gerion, D.; Pellegrino, T.; Zanchet, D.; Micheel, C.; Williams, S. C.; Boudreau, R.; Le Gros, M. A.; Larabell, C. A.; Alivisatos, A. P. Nanotechnology 2003, 14, R15-R27. (7) Ito, T.; Okazaki, S. Nature 2000, 406, 1027-1031. (8) Marrian, C. R. K.; Tennant, D. M. J. Vac. Sci. Technol. A 2003, 21, S207S215. (9) Ginger, D. S.; Zhang, H.; Mirkin, C. A. Angew. Chem., Int. Ed. 2004, 43, 30-45. (10) Parallel dip-pen nanolithography has been demonstrated, albeit sacrificing resolution for throughput.

biosensor applications.11-17 Template-assisted self-assembly of nanoparticles has recently attracted considerable interest as a method for facile production of composite surface structures that may be applied in creating novel sensors and devices.18-20 Deposition of spheres into arrays of surface pores has been demonstrated at the microscopic21,22 and nanoscopic level,18,19,23-29 based on capillary forces during solvent evaporation or dip coating 18,19,21-25, through spin-coating,26-28 or through application of an external electric field.29 None of these examples involve biofunctionalized nanospheres, and none rely on intrinsic molecular interactions between the spheres and the pores, such as attractive electrostatic forces or specific biomolecular recognition events. (11) Jin, R. C.; Wu, G.; Li, Z.; Mirkin, C. A.; Schatz, G. J. Am. Chem. Soc. 2003, 125, 1643-1654. (12) Storhoff, J. J.; Elghanian, R.; Mucic, R. C.; Mirkin, C. A.; Letsinger, R. L. J. Am. Chem. Soc. 1998, 120, 1959-1964. (13) Taton, T. A.; Mirkin, C. A.; Letsinger, R. L. Science 2000, 289, 17571760. (14) Tan, E.; Wong, J.; Nguyen, D.; Zhang, Y.; Erwin, B.; Van Ness, L. K.; Baker, S. M.; Galas, D. J.; Niemz, A. Anal. Chem. 2005, 77, 7984-7992. (15) Storhoff, J. J.; Lucas, A. D.; Garimella, V.; Bao, Y. P.; Muller, U. R. Nat. Biotechnol. 2004, 22, 883-887. (16) Cao, Y. W. C.; Jin, R. C.; Mirkin, C. A. Science 2002, 297, 1536-1540. (17) Park, S. J.; Taton, T. A.; Mirkin, C. A. Science 2002, 295, 1503-1506. (18) Cui, Y.; Bjork, M. T.; Liddle, J. A.; Sonnichsen, C.; Boussert, B.; Alivisatos, A. P. Nano Lett. 2004, 4, 1093-1098. (19) Cui, Y.; Banin, U.; Bjork, M. T.; Alivisatos, A. P. Nano Lett. 2005, 5, 1519-1523. (20) Liu, K.; Baker, S. M.; Tuominen, M.; Russell, T. P.; Schuller, I. K. Phys. ReV. B 2001, 63, article 060403. (21) Xia, Y. N.; Yin, Y. D.; Lu, Y.; McLellan, J. AdV. Funct. Mater. 2003, 13, 907-918. (22) Yin, Y. D.; Lu, Y.; Gates, B.; Xia, Y. N. J. Am. Chem. Soc. 2001, 123, 8718-8729. (23) Juillerat, F.; Solak, H. H.; Bowen, P.; Hofmann, H. Nanotechnology 2005, 16, 1311-1316. (24) Lee, S. H.; Diana, F. S.; Badolato, A.; Petroff, P. M.; Kramer, E. J. J. Appl. Phys. 2004, 95, 5922-5924. (25) Misner, M. J.; Skaff, H.; Emrick, T.; Russell, T. P. AdV. Mater. 2003, 15, 221-224. (26) Xia, D. Y.; Brueck, S. R. J. J. Vac. Sci. Technol. B 2004, 22, 3415-3420. (27) Xia, D. Y.; Biswas, A.; Li, D.; Brueck, S. R. J. AdV. Mater. 2004, 16, 1427-1432. (28) Xia, D. Y.; Brueck, S. R. J. Nano Lett. 2004, 4, 1295-1299. (29) Zhang, C. L.; Xu, T.; Butterfield, D.; Misner, M. J.; Ryu, D. Y.; Emrick, T.; Russell, T. P. Nano Lett. 2005, 5, 357-361.

10.1021/la0534773 CCC: $33.50 © 2006 American Chemical Society Published on Web 04/19/2006

Deposition of DNA-Au Nanospheres into Nanopores

To achieve the deposition of individual or a controllable, small number of particles per pore, the particle size and pore diameter must be matched.18,21,22 For the deposition of nanoparticles with a diameter less than or equal to 50 nm, the fabrication of nanostructured surface templates of appropriate dimensions through top-down approaches suffers the disadvantages discussed above. Alternatively, it is possible to fabricate surface nanopore arrays with pore diameters less than 50 nm covering the entire area of a macroscopic silicon substrate through a parallel, bottomup self-assembly approach, based on microphase separation within polystyrene-b-poly(methyl methacrylate) (PS-b-PMMA) diblock copolymer thin films.20,30-35 PS-b-PMMA diblock copolymers with a PS/PMMA volume ratio of 70:30 form PMMA cylinders within a PS matrix after annealing. By carefully controlling the surface interactions between the diblock copolymer and the substrate, it is possible to orient the PMMA cylinders perpendicular to the substrate.36 Exposing the annealed PS-b-PMMA thin film to UV radiation cross-links the PS and degrades the PMMA. Removal of the degraded PMMA through appropriate solvents leaves behind a hexagonally ordered nanoporous PS thin film. The center-to-center pore spacing and pore diameter of the nanoporous templates can be controlled through the molecular weight of the diblock copolymer.32 Deposition of semiconductor quantum dots from organic solvents into these types of nanoporous templates through dip coating or application of external electric fields has been demonstrated.25,29 We herein report how chemical functionalization of these nanoporous templates with aminosilanes or oligonucleotide probe sequences can be used to effect the spontaneous deposition of DNA-functionalized gold nanospheres into the surface nanopores based on electrostatic interactions or specific DNA hybridization, respectively. To our knowledge, the deposition of biofunctionalized nanoparticles, particularly DNA-functionalized gold nanospheres, into arrays of surface nanopores has thus far not been reported. Deposition of biofunctionalized nanoparticles into nanostructured surfaces based on intrinsic molecular interactions, in particular, bio-inspired, specific self-assembly, is expected to facilitate the fabrication of complex surface structures and enable the development of biosensor surfaces. Materials and Methods General Reagents and Instrumentation. All water used in these experiments was purified through a MilliQ Biocel system to a resistivity of 18 MΩ cm. Test-grade silicon wafers were purchased from International Wafer Service, Inc. (Santa Clara, CA). A random PS-PMMA copolymer (PS-r-PMMA; styrene fraction: 0.58) with a terminal surface reactive hydroxyl group (neutral brush)36 was obtained from the Russell Laboratory at the University of Massachusetts Polymer Science and Engineering Department (Amherst, MA). Two different PS-b-PMMA diblock copolymers were used, the first being of 77 kDa MW, with a PS/PMMA volume ratio of 72:28 and a polydispersity of 1.09 (Polymer Source, Inc., Dorval, Que´bec-Canada), and the second being of 147 kDa MW, with a (30) Thurn-Albrecht, T.; Steiner, R.; DeRouchey, J.; Stafford, C. M.; Huang, E.; Bal, M.; Tuominen, M.; Hawker, C. J.; Russell, T. AdV. Mater. 2000, 12, 787-791. (31) Guarini, K. W.; Black, C. T.; Yeuing, S. H. I. AdV. Mater. 2002, 14, 1290-1294. (32) Xu, T.; Kim, H. C.; DeRouchey, J.; Seney, C.; Levesque, C.; Martin, P.; Stafford, C. M.; Russell, T. P. Polymer 2001, 42, 9091-9095. (33) Black, C. T.; Guarini, K. W.; Milkove, K. R.; Baker, S. M.; Russell, T. P.; Tuominen, M. T. Appl. Phys. Lett. 2001, 79, 409-411. (34) Montero, M. I.; Liu, K.; Stoll, O. M.; Hoffmann, A.; Akermann, J. J.; Martin, J. I.; Vicent, J. L.; Baker, S. M.; Russell, T. P.; Leighton, C.; Nogues, J.; Schuller, I. K. J. Phys. D: Appl. Phys. 2002, 35, 2398-2402. (35) Baker, S. M.; Kolthammer, W. S.; Tan, J. B.; Smith, G. S. Z. Kristallogr. 2004, 219, 179-185. (36) Mansky, P.; Liu, Y.; Huang, E.; Russell, T. P.; Hawker, C. Science 1997, 275, 1458-1460.

Langmuir, Vol. 22, No. 11, 2006 4979 PS/PMMA volume ratio of 77:23 and a polydispersity of 1.06 (from the Russell Lab, UMass Amherst). Chlorodimethyloctadecylsilane and 3-aminopropyl-trimethoxysilane (APTES, 97%) were purchased from Sigma-Aldrich (St. Louis, MO). (3-Trimethoxysilylpropyl)diethylenetriamine (DETA, 95%) was purchased from Gelest (Morrisville, PA). Oligonucleotides were purchased from Integrated DNA Technologies, Inc. (Coralville, IA) and were used for functionalization of the Au nanospheres (5′-SH-TTT TTT TTT CGG TCT GGC GCT-PO3-3′, obtained in disulfide-protected form), for functionalization of the silicon substrates (5′-GAT CGA CGA GAT TTT TTT TTT-NH2-3′) and as complementary (5′-ATC TCG TCG ATC AGC GCC AGA CCG- 3′) or noncomplementary (5′-CGA CTT CAA TGG AGC GCC AGA CCG-3′) bridging target sequences. Other general reagents were purchased through Fisher Scientific (Hampton, NH) or Sigma-Aldrich (St. Louis, MO). Spin coating was performed using a model PWM32 spin coater (Headway Research, Inc., Garland, TX). Atomic force microscopy (AFM) was performed using a Digital Instruments (Santa Barbara, CA) BioScope AFM, in air, mounted on an inverted optical microscope, with a Nanoscope IIIa controller, in tapping mode. Samples were UV exposed with a UVP model XX-15S Sterilaire lamp (Upland, CA). Neutral brush and diblock copolymer film thicknesses were measured using a variable-angle spectroscopic ellipsometer, model LSE (Gaertner Scientific Corporation, Skokie, IL). Contact angle measurements were performed using a NRL contact angle goniometer, model no. 100-00 (Rame-Hart, Inc., Mountain Lake, NJ). UV-visible (UV-vis) spectroscopy was performed using a SPECTRAmax PLUS 384 cuvette/plate reader (Molecular Devices, Sunnyvale, CA) and dynamic light scattering (DLS) was performed using a Protein Solutions DynaPro-MS/X system (12 µL quartz sample cell, Proterion Corporation, Piscataway, NJ). X-ray photoelectron spectroscopy (XPS) was performed using an M-probe surface spectrometer (Surface Science Instruments), with focused, monochromatic Al KR irradiation (1486.6 eV) at an angle of 55° relative to the surface normal. Preparation of Nanoporous Templates. Preparation of nanoporous templates was performed mostly according to literature procedures.31,32 Silicon wafers cut into approximately 1 cm2 pieces were cleaned for more than 4 h in aqua regia (concn HCl/concn HNO3, 3:1), rinsed in water, dried under a stream of argon, and then heated for ∼2 min on a hot plate to evaporate any remaining water. For surface neutralization using the “neutral brush” approach, a layer of neutral brush (solution of 1 wt %/wt in toluene) was spun onto the substrates, which were then annealed for 2 days at 165 °C in a vacuum oven. After removal of excess neutral brush through rinsing with toluene, the thickness of the neutral brush layer was determined by ellipsometry to be between 4 and 6 nm. For surface neutralization using the alkylchlorosilane approach, the silicon surfaces were refluxed in a 2% solution of chlorodimethyloctadecylsilane in toluene for approximately 2.5 h. Samples were then rinsed in toluene, dried with argon gas and allowed to cure overnight. Prior to further processing, the substrates were again rinsed with toluene. For surface neutralization using the hydrogen-passivation approach, silicon substrates were etched with 17% hydrofluoric acid for 3 min, rinsed with water for 5 min to hydrogen-passivate the exposed silicon surface, dried under argon, and used immediately. For the template fabrication, solutions of 77 or 147 kDa PSPMMA diblock copolymers (1 wt %/wt in toluene) were spin-coated onto the neutralized surfaces to obtain a 35-40 nm thick diblock copolymer thin film. Substrates were then thermally annealed at 165 °C for 2 days in a vacuum oven, exposed to 254 nm UV radiation (for 7-12.5 min, depending on template type), and sonicated in glacial acetic acid and water for 30 s each. Samples prepared on neutral brush or alkylchlorosilane substrates were then etched with 5% hydrofluoric acid for 5-10 s. Functionalization of Nanoporous Templates. For aminosilane surface modification, the nanoporous templates were immersed for 5-7 min in a 2% (v/v) solution of APTES or DETA in 95% ethanol. Following modification, surfaces were briefly rinsed with ethanol

4980 Langmuir, Vol. 22, No. 11, 2006 and dried under a stream of argon. The samples were left overnight for curing of the silane layer. For the DNA functionalization of nanoporous templates, the aminemodified probe oligonucleotide was chemically activated using cyanuric chloride according to literature procedures.37 The activated probe sequence was purified through a NAP-5 size exclusion column with 0.1 M Na-borate as the mobile phase. APTES-modified nanoporous templates were then incubated with this activated probe sequence for 12 h at room temperature, rinsed with MilliQ water, and stored at 4 °C for no more than a week. Preparation of DNA-Functionalized Au Nanospheres. The synthesis of monodisperse 13 nm diameter gold nanospheres and their functionalization with the thiol-modified probe oligonucleotide was performed as previously reported.14 The DNA-Au nanospheres were characterized by UV-vis spectroscopy, and their concentration was determined using a molar extinction coefficient of 2.7 × 108 M-1 cm-1 at λ520.11 Monodispersity and average hydrodynamic radius were determined via DLS from the autocorrelation function of the scattered light of 25 to 40 readings using 10-30 s reading windows. Monodispersity was further evaluated using AFM imaging of DNA nanospheres adsorbed to a DETA-modified flat silicon substrate, and the diameter of the dehydrated DNA-Au nanospheres was determined from the particle z-height using section analysis of these AFM images. Deposition of DNA-Au Nanospheres into Nanoporous Templates. To deposit the DNA-Au nanospheres into DETA-functionalized nanoporous substrates based on electrostatic interactions, 40 µL samples of a 2-5 nM solution of DNA-Au nanospheres in TE buffer (10 mM Tris-HCl, 0.5 mM EDTA, pH 7.5) containing 0.01% sodium azide were placed onto the template surface and incubated for 1-2 h at room temperature. The samples were then washed with 2X phosphate-buffered saline (PBS; 40 mM NaPi, pH 7.4, 300 mM NaCl), rinsed with water, and dried under a stream of argon. To deposit the DNA-Au nanospheres into nanoporous templates functionalized with probe oligonucleotides based on specific DNA hybridization, 40 µL of a solution containing 2-5 nM of DNA-Au nanospheres plus 1 µM of the complementary or noncomplementary bridging target oligonucleotide in a standard high ionic strength hybridization buffer commonly used for DNA microarray experiments (4X SSC (60 mM Na-citrate and 600 mM NaCl); 500 mM Tris-HCl, pH 7.5; 0.2% SDS (sodium dodecyl sulfate)) was placed onto the template surface and incubated for 2 h at room temperature. The substrates were then washed with a solution containing 2X SSC (30 mM Na-citrate, 300 mM NaCl) and 0.2% SDS, followed by two washes with 0.5X SSC (7.5 mM Na-citrate and 75 mM NaCl), and drying under a stream of argon. Following DNA nanosphere deposition, samples were imaged via tapping-mode AFM, and characterized spectroscopically via XPS. To highlight the DNA-Au nanospheres deposited inside of the surface nanopores, manual coloration of AFM images was performed using Adobe Photoshop by selecting the regions of the images representing the DNA nanospheres deposited inside of the surface nanopores, and altering the color balance of these regions using the Adobe color balance tool. For comparison, the original uncolorized images are included in the Supporting Information.

Results and Discussion The nanoporous templates used in our studies are fabricated from thin films of PS-b-PMMA diblock copolymers with molecular weights of 77 or 147 kDa (Figure 1). We used three different approaches to achieve suitable surface neutralization. In the first and second method, the SiOx surface layer of the silicon substrate is chemically modified either with an appropriate PS-PMMA random copolymer referred to as neutral brush,36

(37) Vanness, J.; Kalbfleisch, S.; Petrie, C. R.; Reed, M. W.; Tabone, J. C.; Vermeulen, N. M. J. Nucleic Acids Res. 1991, 19, 3345-3350.

Bandyopadhyay et al.

Figure 1. Tapping-mode AFM height images of nanoporous templates prepared from thin films of PS-b-PMMA diblock copolymers with molecular weights of 77 kDa (images a, c, and e) and 147 kDa (images b, d, and f). The templates shown in images a and b were prepared on silicon substrates functionalized with a layer of “neutral brush” (PS-r-PMMA random copolymer, PS weight fraction 0.59). The templates shown in images c and d were prepared on silicon substrates functionalized with a submonolayer of chlorodimethyloctadecylsilane.38 The templates shown in images e and f were prepared on silicon substrates that were HF etched to remove the SiOx layer, followed by hydrogen passivation. Scale bars of all images: 200 nm.

or with a long chain alkylchlorosilane.38 In the third approach, the SiOx surface layer is removed using hydrofluoric acid (HF) etching followed by hydrogen passivation of the underlying silicon surface.32 By using 77 or 147 kDa PS-b-PMMA, we obtained nanoporous surfaces with average center-to-center pore spacings of 51 ( 5 and 66 ( 11 nm, and with estimated pore diameters of 23 ( 4 and 33 ( 7 nm, respectively, determined by AFM as shown in Figure 1. The exact pore diameters cannot be determined by AFM because of convolution of the tip with the narrow pores. Nanoporous films generated on hydrogen-passivated silicon surfaces using 77 kDa PS-b-PMMA were found to be less regular than the analogous 77 kDa templates on neutral brush or alkylchlorosilane surfaces. We were unable to generate wellordered templates from 147 kDa PS-b-PMMA thin films on hydrogen-passivated silicon surfaces. For optimal annealing, the thickness of the polymer thin films is restricted to between 30 and 45 nm, depending on the copolymer molecular weight and the surface neutralization approach used. Consequently, the (38) Bandyopadhyay, K.; Cha, K.; Tan, E.; Niemz, A.; Baker, S. M. To be submitted for publication, 2006.

Deposition of DNA-Au Nanospheres into Nanopores

resulting surface nanopores have an aspect ratio of approximately 1:1. Lower molecular weight films give rise to better long-range hexagonal order. Various approaches have been reported to obtain better long-range order and domain alignment for diblock copolymer thin films.39-45 To obtain DNA-Au nanospheres, we synthesized gold colloids through the reduction of HAuCl4 with sodium citrate.46 We then functionalized these “bare”, that is, citrate-stabilized, Au nanospheres with thiol-terminated oligonucleotides (Figure 2a) according to established literature procedures,11 with a few modifications.14 The bare and DNA-functionalized Au nanospheres exhibit a plasmon resonance λmax of 520 and 522 nm, respectively, as determined by UV-vis spectroscopy. Through DLS we found that the bare and DNA-functionalized Au nanospheres were monodisperse, with average hydrodynamic diameters of 13.2 ( 0.9 and 30.1 ( 7.3 nm, respectively. This hydrodynamic diameter represents the diameter of the nanoparticle plus the hydration sphere, that is, the solvent layer that moves along with the particle. For the DNA-functionalized Au nanospheres, it also includes the extended and fully hydrated oligonucleotides, thus resulting in the significant size increase. We also used AFM section analysis to determine the diameter of the surface immobilized nanospheres from the particle z-height. The dehydrated diameters of the bare and DNA-functionalized Au nanospheres were found to be 11.4 ( 1.9 and 18.6 ( 2.5 nm, respectively. For the DNA-Au nanospheres, this dehydrated diameter represents the size of the particles with the oligonucleotides collapsed onto the surface following the removal of solvent molecules, and is therefore considerably smaller than the hydrodynamic diameter. Our approach for the deposition of individual DNA-Au nanospheres into the surface nanopores is based on introducing intrinsic attractive interactions through chemical modification of the nanoporous templates, as illustrated in Figure 2c-d. To effect the electrostatic immobilization of negatively charged DNA-functionalized gold nanospheres, we modified the nanoporous templates with a positively charged aminosilane. To effect the immobilization of DNA-functionalized gold nanospheres mediated by DNA hybridization, we further functionalized aminosilane-modified nanoporous templates with probe oligonucleotides. Results for the deposition of DNA-Au nanospheres into nanoporous templates based on electrostatic interactions are shown in Figures 3 and 4. We found that surface functionalization with a silane containing multiple positively charged amine groups, such as DETA, results in significantly greater electrostatic immobilization efficiencies compared to those obtained for surface functionalization with a silane containing a single amine group, such as APTES.47 Nanoparticles were deposited from low ionic strength buffer to optimize the electrostatic interactions. Once (39) Black, C. T.; Bezencenet, O. IEEE Trans. Nanotechnol. 2004, 3, 412415. (40) Morkved, T. L.; Lu, M.; Urbas, A. M.; Ehrichs, E. E.; Jaeger, H. M.; Mansky, P.; Russell, T. P. Science 1996, 273, 931-933. (41) Thurn-Albrecht, T.; DeRouchey, J.; Russell, T. P.; Jaeger, H. M. Macromolecules 2000, 33, 3250-3253. (42) De Rosa, C.; Park, C.; Thomas, E. L.; Lotz, B. Nature 2000, 405, 433437. (43) Park, C.; De Rosa, C.; Thomas, E. L. Macromolecules 2001, 34, 26022606. (44) Rockford, L.; Mochrie, S. G. J.; Russell, T. P. Macromolecules 2001, 34, 1487-1492. (45) Sundrani, D.; Sibener, S. J. Macromolecules 2002, 35, 8531-8539. (46) Grabar, K. C.; Freeman, R. G.; Hommer, M. B.; Natan, M. J. Anal. Chem. 1995, 67, 735-743. (47) Niemz, A.; Baker, S. M. Self-Assembled DNA Nanoarrays. MRS Spring Meeting Proceedings; Materials Research Society: Warrendale, PA, 2004; Vol. 818, M10.3.

Langmuir, Vol. 22, No. 11, 2006 4981

Figure 2. (a) Schematic representation of the synthesis of DNAAu nanospheres. (b) Tapping-mode AFM height image and section analysis (z-height trace) of DNA-Au nanospheres deposited onto a DETA-modified silicon substrate. Particles are monodisperse with an average z-height of 18.6 ( 2.5 nm. Scale bar: 200 nm. (c,d) Schematic representations of template functionalization and DNA nanosphere deposition based on electrostatic interactions and specific DNA hybridization, respectively. Deposition is mediated through functionalization of the nanopore’s bottom surface with DETA or with APTES followed by coupling to an activated probe oligonucleotide. Schemes represent cross-sections of the nanoporous templates, showing the SiOx bottom surfaces and cross-linked PS walls of the surface nanopores.

immobilized, the nanoparticles are not removed upon washing with high ionic strength buffers, such as 300 mM NaCl. Successful deposition of DNA nanospheres was observed via AFM using nanoporous templates prepared from thin films of 147 kDa PSb-PMMA diblock copolymer on either neutral brush- or alkylchlorosilane-functionalized substrates, and from thin films of 77 kDa PS-b-PMMA diblock copolymer on hydrogenpassivated substrates. In each case, the deposition was carried out for 1 h, with little further improvement observed for longer deposition times. No or very little deposition could be observed for nanoporous templates prepared using 77 kDa PS-b-PMMA diblock copolymer on neutral brush- or alkylchlorosilanefunctionalized substrates. For nanoporous templates prepared on a layer of neutral brush or alkylchlorosilane, we observed that an additional brief etch in 5% HF, following the regular sonication in acetic acid and water and prior to modification with DETA, increases the inclusion density for the electrostatic deposition of DNA nanospheres. Prolonged HF etching of nanoporous PS templates on Si/SiOx substrates is known to dissolve the SiOx

4982 Langmuir, Vol. 22, No. 11, 2006

Figure 3. Tapping-mode AFM images following exposure of DETAfunctionalized (a,c,e) and unfunctionalized (b,d,f) nanoporous templates to DNA-Au nanospheres. DNA-Au nanospheres are deposited only into the DETA-functionalized nanoporous templates, based on intrinsic electrostatic interactions. Images are manually colorized to highlight in red the DNA-Au nanospheres deposited into the nanopores. Original images are included in the Supporting Information. Nanoporous templates were fabricated from 147 kDa PS-b-PMMA thin films on a “neutral brush”-functionalized substrate (a,b), 147 kDa PS-b-PMMA on a chlorodimethyloctadecylsilanefunctionalized substrate (c,d), and 77 kDa PS-b-PMMA on a hydrogen-passivated substrate (e,f). Templates a-d were exposed to a brief HF etch following UV exposure and regular solvent processing. Scale bars of all images: 200 nm.

surface layer and detach the nanoporous PS thin film.25,29 We shortened the HF etching time to a point at which no damage to the nanoporous template is observable by AFM. Additional HF etching was not found to be necessary or beneficial for nanoporous templates prepared on hydrogen-passivated Si substrates, which do not involve an organic neutralization layer. We confirmed the surface immobilization of DNA-modified gold nanospheres using XPS through the presence of the Au 4f7/2 and 4f5/2 peaks at binding energies of 84 and 88 eV (Figure 4a). AFM section analysis (z-height trace) confirms that the DNAAu nanospheres are deposited inside of the surface nanopores (Figure 4c). AFM images show that, in general, no more than a single nanosphere is immobilized per nanopore. On the basis of the approximately 1:1 aspect ratio of the nanopores, it can be assumed that the nanospheres are not stacked on top of each other. Using AFM and XPS, we confirmed that the DNA nanospheres do not deposit onto the unmodified nanoporous PS templates (Figures 3b,d,f and 4b). This observation supports the assumption that the deposition of DNA nanospheres is caused

Bandyopadhyay et al.

Figure 4. XPS Au 4f region of nanoporous templates (147kDa Ps-b-PMMA, neutral-brush substrates) that were (a) DETAfunctionalized or (b) left unfunctionalized, followed by exposure to DNA-Au nanospheres for 1 h. Deposition of gold nanospheres is only observed for the DETA-functionalized template. Excerpts of tapping-mode AFM images and corresponding section analysis of (c) the DETA-modified template, showing DNA-Au nanosphere deposition into the surface nanopores (indicated by dashed circles and black arrows), and (d) the unmodified template, showing a typical height profile of empty surface nanopores. Scale bars of AFM images: 100 nm.

Figure 5. Tapping-mode AFM images of nanoporous templates fabricated from 77 kDa PS-b-PMMA thin films on hydrogenpassivated substrates that were functionalized with probe DNA and then exposed for 2 h to (a) DNA-Au nanospheres plus a complementary bridging target oligonucleotide and (b) DNA-Au nanospheres plus a noncomplementary bridging target oligonucleotide. DNA-Au nanospheres are deposited only in the presence of a complementary bridging target oligonucleotide, based on specific DNA hybridization interactions. Images are manually colorized to highlight in red the DNA-Au nanospheres deposited into the nanopores. Original images are included in the Supporting Information. Scale bars: 200 nm.

by electrostatic interactions introduced through functionalization with DETA. However, we observed varying degrees of deposition of DNA-Au nanospheres on top of DETA-functionalized templates (Figure 4a,c,d). Results for the deposition of DNA-Au nanospheres into nanoporous templates based on specific hybridization interactions are shown in Figure 5. To immobilize DNA probe sequences within the surface nanopores, we functionalized the nanoporous templates with the monoaminosilane APTES, and then coupled

Deposition of DNA-Au Nanospheres into Nanopores

Langmuir, Vol. 22, No. 11, 2006 4983 Table 1.

sample

inclusion density a,c

selectivity b,c

147 kDa copolymer, neutral brush surface, electrostatic deposition 147 kDa copolymer, alkylchlorosilane surface, electrostatic deposition 77 kDa copolymer, H-passivated surface, electrostatic deposition 77 kDa copolymer, H-passivated surface, DNA hybridization deposition

18 ( 2% 7 ( 1% 34 ( 2% 8 ( 2%

73 ( 9% 52 ( 4% 62 ( 3% 85 ( 4%

a Inclusion density ) number of DNA nanospheres inside the pores divided by the total number of pores (in %). b Selectivity ) number of DNA nanospheres inside the pores divided by the total number of deposited DNA nanospheres (in %). c Average and standard deviation values derived from four representative 1 × 1 µm areas.

those primary amine groups with a probe oligonucleotide bearing a 3′-terminal amine group, which was activated with cyanuric chloride,37,48,49 as illustrated in Figure 2d. This protocol was found to be compatible with preserving the integrity of the nanoporous templates. Since the probe sequences immobilized on the DNA nanospheres and on the surface are designed to be noncomplementary, the DNA nanospheres are not expected to deposit into the probe-functionalized nanopores unless a complementary bridging target sequence is added.50 Indeed, we observed DNA nanosphere deposition into DNA-functionalized nanoporous templates in the presence of the complementary bridging target sequence (Figure 5a), but not in the presence of a noncomplementary control sequence (Figure 5b), nor in the absence of target oligonucleotides, which verifies the specificity of the process. We found that a brief rinse of the nanoporous template in 0.1 N NaOH following APTES modification and prior to exposure to the cyanuric chloride-activated probe sequences significantly reduces the deposition of DNA nanospheres on top of the template. As shown in Figure 5a, most of the DNA nanospheres are deposited into the surface nanopores, with very little deposition on top of the template. The templates shown in Figure 5 were prepared from 77 kDa MW diblock copolymer thin films on hydrogen-passivated substrates. Similar DNA nanosphere deposition, albeit at a slightly lower inclusion density, was observed using nanoporous templates prepared from 147 kDa MW diblock copolymer thin films on an alkylchlorosilane neutralization layer exposed to a brief HF etch, as described above, followed by APTES and DNA functionalization. For the DNA nanosphere deposition into the different types of nanoporous templates on the basis of either electrostatic interactions or specific DNA hybridization, we calculated the inclusion density, defined as the percentage of the total number of pores containing a DNA nanosphere deposited inside the pore, and the selectivity, defined as the percentage of the total number of deposited DNA nanospheres that reside inside of, as opposed to on top of, the template (Table 1). In these preliminary studies, we observed relatively low inclusion densities and moderate selectivities, which nevertheless provided proof for the feasibility of this approach. We intend to increase the inclusion density and selectivity by optimizing the conditions for template preparation, functionalization, and DNA nanosphere deposition. Achieving a high inclusion density requires proper matching between the diameters of the DNA nanospheres and nanopores to avoid steric hindrance. For the nanoporous templates prepared using 77 kDa PS-b-PMMA diblock copolymer, the pore diameter estimated using AFM (23 ( 4 nm) is commensurate with the dehydrated diameter of the DNA nanospheres (18.6 ( 2.5 nm), but is smaller than the hydrodynamic diameter of the fully (48) Steinberg, G.; Stromsborg, K.; Thomas, L.; Barker, D.; Zhao, C. F. Biopolymers 2004, 73, 597-605. (49) Lee, P. H.; Sawan, S. P.; Modrusan, Z.; Arnold, L. J.; Reynolds, M. A. Bioconjugate Chem. 2002, 13, 97-103. (50) Taton, T. A.; Mucic, R. C.; Mirkin, C. A.; Letsinger, R. L. J. Am. Chem. Soc. 2000, 122, 6305-6306.

hydrated DNA nanospheres in solution (30.1 ( 7.3 nm, determined by DLS). Nanoporous templates prepared using 147 kDa PS-b-PMMA diblock copolymer have larger pore diameters, as estimated using AFM (33 ( 7 nm), which should minimize steric hindrance. However, the highest inclusion density observed so far was obtained for the electrostatic deposition into nanoporous templates prepared from 77 kDa PS-b-PMMA diblock copolymer on hydrogen-passivated silicon substrates. This observation suggests that the dehydrated particle diameter is the more relevant factor, although one would expect that the required partial desolvation of DNA-Au nanospheres, which is captured by the size difference between the hydrodynamic and the dehydrated particle diameters, would impede the deposition of DNA nanospheres into the surface nanopores. An alternative explanation is the occurrence of a “sieving effect”, wherein the smaller particles are selected for deposition from solution. Steric hindrance alone also does not explain the low deposition yield observed for templates prepared from 77 kDa PS-b-PMMA on silicon substrates functionalized with neutral brush or alkylchlorosilane layers, even though the nanopore diameters of these templates are slightly smaller than the nanopore diameters of templates prepared from the same molecular weight diblock copolymer on hydrogen-passivated substrates. Achieving a high inclusion density further requires effective functionalization of the nanopores’ bottom surfaces to introduce the necessary attractive interactions. Effective functionalization in turn requires that any residual polymer or other organic matter has been removed from the pores, and that the nanopores’ bottom surfaces indeed consist of exposed SiOx. The increase in inclusion density observed for nanoporous templates on neutral brush- or alkylchlorosilane-functionalized silicon substrates upon exposure to a brief HF etch provides indirect evidence that the regular UV exposure and solvent processing, even under optimized conditions, is insufficient for the complete removal of the underlying chemically bound neutralization layer. The short HF etch presumably dissolves the surface layers of the partially exposed SiOx bottom surface of the nanopores enough to aid in the removal of attached organic remnants without detaching the template. This hypothesis is further supported by the observation that a similar HF etch step is not required for nanoporous templates fabricated on hydrogen-passivated surfaces, which do not contain an organic neutralization layer. Selective DNA nanosphere deposition inside the surface nanopores, as opposed to on top of the template, is based on the assumption that the nanopores’ walls, which consist of crosslinked PS, are intrinsically inert to the DNA nanospheres, and remain unfunctionalized during the silanization and probemodification steps. It has ben reported that UV-cross-linked PS does not associate with DNA under neutral or slightly basic conditions.51 This is in agreement with our observation that DNA nanospheres do not deposit on top of unfunctionalized nanoporous templates (Figure 3b,d,f) or on top of control surfaces consisting (51) Allemand, J. F.; Bensimon, D.; Jullien, L.; Bensimon, A.; Croquette, V. Biophys. J. 1997, 73, 2064-2070.

4984 Langmuir, Vol. 22, No. 11, 2006

of PS thin films on hydrogen-passivated silicon, which were thermally annealed, UV-exposed, and solvent-processed in a manner analogous to that of the nanoporous templates (data not shown). We observed, however, deposition of DNA nanospheres, particularly on top of DETA-modified nanoporous templates (Figure 3a,c,e) and on top of PS thin film control surfaces that were subjected to the same processing and DETA modification protocols as the nanoporous templates. This indicates that either DETA sol particles are deposited onto the cross-linked PS during the functionalization process, or the aminosilane DETA reacts with cross-linked PS, which may have been oxidized during the UV processing step. We are currently exploring alternate modification protocols and methods to remove deposits from the top of the templates or to passivate the template walls. We observed a higher selectivity for DNA nanosphere deposition based on specific DNA hybridization on APTESmodified and probe-functionalized templates compared to that for DNA nanosphere deposition based on electrostatic interactions on DETA-modified templates. A significant improvement in selectivity can be achieved if the nanoporous templates are subjected to a brief rinse in 0.1 N NaOH following APTES modification and prior to exposure to the cyanuric chlorideactivated probe sequences. It has been reported that silane monolayers are etched off of a substrate upon exposure to 0.1 N NaOH for several minutes.52 A brief rinse in 0.1 N NaOH is therefore expected to remove via hydrolysis any APTES that may have physisorbed or chemisorbed onto the cross-linked PS walls of the nanopores, and the exposure is kept short enough to avoid degradation of the silane layer at the bottom of the surface nanopores. We are exploring similar approaches to improve the selectivity of electrostatic DNA nanosphere deposition. The higher selectivity for DNA nanosphere deposition based on specific DNA hybridization interactions can also, in part, be attributed to the use of a high ionic strength buffer, which maximizes specific DNA-hybridization and screens unwanted electrostatic interactions. The inclusion density for DNA nanosphere deposition based on specific hybridization was found to be lower than that for deposition into the same type of template based on electrostatic interactions. The precise nature of the observed difference requires further investigation, but we expect that the inclusion density can be improved by optimizing the conditions for DNA probe functionalization of surface nanopores and for hybridization of DNA-Au nanospheres to the probefunctionalized nanoporous templates.

Conclusions We have demonstrated the spontaneous deposition of individual DNA-functionalized Au nanospheres into surface nanopores of nanoporous templates prepared from PS-b-PMMA diblock copolymer thin films. The deposition is driven by intrinsic electrostatic interactions or by specific DNA hybridization. On the basis of the size matching between the nanospheres and nanopores, we are able to achieve the deposition of predominantly (52) Wasserman, S. R.; Tao, Y. T.; Whitesides, G. M. Langmuir 1989, 5, 1074-1087.

Bandyopadhyay et al.

individual nanospheres in the surface nanopores. To our knowledge, this is the first reported example for the deposition of biofunctionalized nanospheres into nanoporous surfaces, and the first example of nanosphere deposition based on intrinsic interactions, especially involving biomolecular recognition. The entire process, including the preparation of the nanoporous templates used herein, is based on self-assembly and can be conducted in a parallel and scalable manner using minimal instrumentation. Both the electrostatic- and hybridization-based DNA nanosphere depositions into surface nanopores occur relatively rapidly, within 1-2 h, which is an advantage over the lengthy process required for deposition based on solvent evaporation. Both approaches further use small volumes of DNAAu nanospheres at relatively low concentration in aqueous buffer, an advantage over nanosphere deposition based on dip coating or spin coating, which require large volumes (>5 mL as opposed to 50 µL) or high concentrations (1 wt %) of the nanosphere solution that are difficult to achieve using biofunctionalized nanospheres. We are optimizing the processes to increase the inclusion density and to decrease deposition on top of the nanoporous template. We will further use these DNA nanosphere nanoarrays as capture surfaces for the secondary self-assembly of other DNAfunctionalized nanoscopic entities based on hybridization through short bridging oligonucleotides. Such nanoporous structures used in secondary self-assembly can include another set of DNA-Au nanospheres, as well as DNA-functionalized quantum dots,53 or carbon nanotubes.54,55 The confined space of the surface nanopores is expected to render the deposited DNA nanospheres highly anisotropic, and to result in discrete 1:1 assembly with other entities in solution, as opposed to the generally observed dendritic growth.50 Our goal is to characterize the optical and electronic properties of these DNA nanosphere arrays or of any other structures obtained by secondary self-assembly, keeping in mind their potential use as biosensor surfaces. Acknowledgment. This work was supported by the National Science Foundation through Research Grants BES-0304675, ECS-0501629, DMR-0213695, and DMR-0109077, and through REU Site Grants EEC-0243910 and CHE-0353662, in addition to research funds from the Keck Graduate Institute. We thank Annie Tan and Travis McQueen for assistance with DNA nanosphere synthesis and nanotemplate preparation, respectively. Supporting Information Available: Original noncolorized images of Figures 3 and 5 are provided for comparison. This material is available free of charge via the Internet at http://pubs.acs.org. LA0534773 (53) Parak, W. J.; Gerion, D.; Zanchet, D.; Woerz, A. S.; Pellegrino, T.; Micheel, C.; Williams, S. C.; Seitz, M.; Bruehl, R. E.; Bryant, Z.; Bustamante, C.; Bertozzi, C. R.; Alivisatos, A. P. Chem. Mater. 2002, 14, 2113-2119. (54) Keren, K.; Berman, R. S.; Buchstab, E.; Sivan, U.; Braun, E. Science 2003, 302, 1380-1382. (55) Zheng, M.; Jagota, A.; Strano, M. S.; Santos, A. P.; Barone, P.; Chou, S. G.; Diner, B. A.; Dresselhaus, M. S.; Mclean, R. S.; Onoa, G. B.; Samsonidze, G. G.; Semke, E. D.; Usrey, M.; Walls, D. J. Science 2003, 302, 1545-1548.