DNA detection DOI: 10.1002/smll.200500322
Glass-Bead-Based Parallel Detection of DNA Using Composite Raman Labels Rongchao Jin, Y. Charles Cao, C. Shad Thaxton, and Chad A. Mirkin* Biomolecule detection has become increasingly important in biomedical research and disease diagnosis.[1–8] In particular, microarrays are quite promising because they allow one to carry out many assays simultaneously with rapid readout provided by a variety of labeling strategies (flourophores, chemiluminescent entities, or nanoparticles).[1–4] Such arrays have been widely used in genomics and proteomics research.[5] In a typical DNA microarray format, glass substrates are spotted with appropriate capture cDNA (> 200 nucleotides) or synthesized oligonucleotide strands (25– 80 nucleotides).[6] Fluorophore-labeled target DNA strands are captured via hybridization to the arrayed capture strands. Since each DNA spot in the microarray has been positionally encoded, the results can be easily determined, typically, by using a computer-controlled laser scanner system. An alternative to the use of spotted microarrays in detecting biomolecules is the random-array approach to detection.[7–13] This approach uses individual beads for an assay in lieu of a spot in a conventional microarray. In a typical assay, batches of beads for different but specific targets are prepared and then mixed. Reporter groups are used to identify the bead, the corresponding target, and whether or not a reaction with the target nucleic acid sequence has taken place. The advantages of the random array format over the microarray approach are that the beads exhibit faster hybridization kinetics, are easier and less expensive to fabricate, and do not require a sophisticated laser scanner system to obtain the results. The disadvantage is that one loses the positional encoding afforded by a microarray. As such, one needs ways of spectroscopically encoding each of the targetspecific beads involved in a particular assay. Fluorophores are commonly used as reporter groups in the random-array strategy,[8] however, due to their relatively broad emission bands and energy transfer between different dye molecules, the number of fluorescent dye labels that can be simultaneously detected in a multiplexing scheme is limited. In addition, one has to perform multiple laser-scanning events
[*] R. Jin, Y. C. Cao, C. S. Thaxton, Prof. C. A. Mirkin Department of Chemistry and International Institute for Nanotechnology, Northwestern University, 2145 Sheridan Road Evanston, IL 60208 (USA) Fax: (+ 1) 847-467-5123 E-mail:
[email protected] Supporting information for this article is available on the WWW under http://www.small-journal.com or from the author. small 2006, 2, No. 3, 375 – 380
since each dye label typically has a distinct excitation wavelength. Nanoparticle probes are beginning to offer the potential to circumvent some of the problems and limitations posed by conventional molecular fluorophore technology.[14–33] For example, there are now a variety of microarray formats that utilize nanoparticle probes for the high-sensitivity and highselectivity detection of DNA and protein targets. These assays provide many types of readout (electrical, Raman, light scatter, fluorescence, and colorimetric), and several of them have significant advantages over conventional fluorophore-labeling approaches with respect to sensitivity, selectivity, simplicity, and multiplexing capabilities.[34–38] Recently, we reported a new microarray format based upon nanoparticle probes and Raman spectroscopy for the multiplexed detection of DNA and protein targets.[4] In this assay, gold nanoparticles are functionalized with different types of Raman dyes that are coded for specific targets and then used as labels in a sandwich assay format. The particle probes are used to catalyze the deposition of silver from a developing solution comprised of an AgI salt and hydroquinone. This silver layer acts as an enhancing layer that allows one to probe the results of the assay in a highly sensitive manner using surface-enhanced Raman scattering (SERS) spectroscopy. The advantages of Raman spectroscopy in this format over fluorescence are twofold: First, one can use SERS to increase sensitivity,[17, 39, 40] allowing one to easily probe target concentrations in the femtomolar to attomolar range.[4] Second, the narrow bands associated with the Raman spectra of the dye-labeled particles, as compared with fluorescence spectra,[41] allow for greater degrees of multiplexing. Therefore, this approach might be exceptionally well-suited for the development of a random-array detection system. Multiplexing and dye-labeled encoding of beads are critical to the development of random arrays.[7, 8, 16] Herein, we report a new methodology that allows one to carry out SERS-based assays on microbeads using nanoparticles with co-adsorbed Raman labels as reporter groups and catalysts for developing a Raman-enhancing silver layer from the surfaces of beads that have captured specific targets (Scheme 1). To demonstrate that composite Raman spectroscopic labels can be generated from dye mixing, we mixed two alkylthiol-capped oligonucleotide strands with the same base sequences but with different Raman labels (TMR and Cy3), and used solutions of Raman-dye-functionalized oligonucleotides at various ratios (molar) to functionalize gold nanoparticles (diameter: 13 1.2 nm; Scheme 1 A). In order to obtain the Raman spectrum for each type of dye-labeled nanoparticle, the particle probes were hybridized to glass beads that were modified with capture strands (Scheme 1 B). At high target concentrations (> 1 nm), the glass beads exhibit a red color due to the high surface coverage of the gold nanoparticles on the bead surface. Below 100 pm they are colorless and the reaction cannot be detected by the naked eye. SERS measurements on the glass beads bearing only the gold nanoparticle probes show a featureless fluorescence background; no distinct SERS signals were observed even at high target concentrations (e.g., > 100 pm).
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Scheme 1. Composite Raman labels generated via dye mixing based on one type of nanoparticle (A) and their use in glass-microbeadbased DNA detection (B).
This is due to the low enhancing capabilities of the gold nanoparticles under the stated conditions. In order to achieve strong SERS enhancement, the beads with hybridized gold nanoparticles were immersed in a silver enhancing solution (Ted Pella, 8 min at room temperature). In this process, the gold nanoparticles act as seeds, and silver is deposited onto the gold-particle surface, resulting in surface morphologies that give strong SERS signals. After silver enhancement, the SERS spectra from beads with nanoparticles with different TMR to Cy3 ratios show unique spectroscopic fingerprints that can be differentiated from each other (Figure 1 A). Furthermore, it was found that the Raman-band intensities correlate well with the molar ratio of the dyes used to functionalize the gold nanoparticles. This suggests that the rates of adsorption for the different dyes are comparable, thus making surface loading of the dyes straightforward. Each composite Raman label can be uniquely identified by examining differences in relative SERS intensities of the bands associated with each surface-immobilized dye. For example, in a two-probe experiment involving Cy3-labeled particles and TMR-labeled particles, one can look at two non-overlapping Raman bands (I1 at 1590 cm 1 for TMR and I2 at 1650 cm 1 for Cy3) and compare their intensity ratios (I1/I2 ; two-peak index) to determine the spectroscopic labels and therefore targets (Figure 1 B).
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Figure 1. A) Raman spectra generated by co-adsorbing two Ramandye-functionalized oligonucleotides onto the Au nanoparticle surface (diameter 13 nm). Spectra a–m correspond to TMR:Cy3 ratios of 1:0, 5:1, 4:1, 3:1, 2:1, 1.5:1, 1:1, 1:1.5, 1:2, 1:3, 1:4, 1:5, 0:1, respectively. B) Two-peak indexing by comparing I1 (1590 cm 1)/I2 (1650 cm 1) intensity ratios.
The SERS spectra of the silver-coated nanoparticles exhibit excellent stability during the Raman measurement (e.g., the spectral intensity drops less than 10 % over a tenminute period). The peak deviation is less than 5 cm 1 from experiment to experiment, and fluctuations of relative intensity ratios for each of the most intense Raman peaks are less than 5 % for one set of developing conditions. Note that the absolute SERS intensity may vary from experiment to experiment and also depends on the silver developing conditions, hence, only the relative intensities are valid and useful for spectroscopic encoding. The strategy of dye mixing also has been extended to three dyes. Examples of new spectroscopic fingerprints generated by mixing three different Raman dyes (TMR, Cy3, and Cy3.5) are shown in Figure 2. In principle, because there are a large number of Raman chromophores with non-overlapping and spectroscopically unique features, a large number of new spectroscopic codes can be generated via this dye ratio approach.
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Figure 2. Examples of new Raman spectra generated by co-adsorbing three Raman-dye-functionalized oligonucleotides onto the nanoparticle surface: a) TMR, b) Cy3, c) Cy3.5, d) TMR:Cy3:Cy3.5 = 20:20:1, e) TMR:Cy3:Cy3.5 = 5:5:1.
Note that the analogous fluorophore dye-mixing approaches are limited by the broad emission bands observed in fluorescence and energy transfer between molecules. To demonstrate the selectivity and suitability of these composite Raman labels for DNA detection, we focused our efforts on a glass bead-based sandwich detection format (Scheme 1 B). In such an assay, glass beads ( 300 mm) were first functionalized with the appropriate strands complementary to a specific target (Scheme 2). Multiple types of beads are then mixed, forming a random array for detecting multiple DNA targets. To test the selectivity of this bead-based approach, we first prepared a composite Raman label (TMR:Cy3 ratio = 1:1), pure TMR, and Cy3 nanoparticle probes to use in a single target assay. Three types of glass beads (beads a, b, and c; sequences are listed in Scheme 2) were mixed and immersed in a phosphate buffer solution (0.3 m NaCl, pH 7, 10 mm; designated as 0.3 m PBS below) containing target a (100 pM) for 2 h at room temperature. The glass beads were washed with a 0.3 m PBS buffer and further treated with a 0.3 m PBS buffer solution containing three types of nanoparticle probes (each 0.5 nm) functionalized with Raman labels associated with target a, b, or c (Scheme 2) for two hours, small 2006, 2, No. 3, 375 – 380
then washed with a phosphate buffer solution (0.3 m NaNO3, pH 7, 10 mm) to remove chloride ions and nonspecifically bound nanoparticles. The glass beads were then treated with silver enhancement solution for 8 min, which resulted in silver deposition where nanoparticle probes were present. Prior to Raman measurements, the microbeads were immersed in a 0.3 m PBS buffer for 10 min to further enhance Raman scattering.[4] When measuring the Raman spectra from randomly selected glass beads, we only observed the composite Raman spectrum corresponding to the 1:1 ratio of TMR to Cy3, which is coded for target a. No pure Cy3 or TMR spectroscopic fingerprints were found, demonstrating that no detectable cross-hybridization (i.e., target hybridized with non-complementary oligonucleotide strands) had taken place. We further tested the effectiveness of the composite nanoparticle probes for multiplexed parallel DNA detection in the bead-based format. Nine batches of glass beads were prepared (see Experimental Section), eight of which were respectively functionalized with eight different capture strands and one which was functionalized with non-complementary oligonucleotide strands (HS-AAA AAA AAA AAA AAA AAA AA, designated as HS-A20) containing no Raman labels and served as a control in the multiplexed assay (Scheme 2). The beads were mixed and immersed in a 0.3 m PBS solution that contained eight different targets (each 100 pM). After DNA hybridization, the glass beads were washed with a 0.3 m PBS buffer and further treated with a 0.3 m PBS buffer solution containing nine types of nanoparticle probes (each 0.5 nm), eight of which are respectively functionalized with composite Raman labels. The remaining probe was functionalized with HS-T20 and served as a control. After hybridization for 2 h, the beads were washed with phosphate buffer solution (0.3 m NaNO3, pH 7) and further immersed in a silver enhancement solution for 8 min, and then rinsed with Nanopure water (resistance 18.1 MW). After silver amplification, the microbeads (including the control beads) appeared as dark-grey spheres (Figure 3 A), and the measured Raman spectra from randomly selected glass beads displayed the correct spectroscopic fingerprints corresponding to each target. A representative SERS spectrum from an individual bead (#1 in Figure 3 A) is shown in Figure 3 B. All the spectra matched well with the standard ones in Figures 1 and 2 (relative peak intensity error < 5 %; peak deviation < 5 cm 1). To simplify the analysis, we assigned a color (circles) to each Raman-labeled probe (Figure 3 A). For ease of Raman analysis, the glass beads also can be aligned along a trough on a glass cover slide, allowing for the rapid acquisition of a Raman signal (Figure 3 C). Note that in the multiplexed assay, a potential problem pertains to the exchange of the surface dye-capped oligonucleotides between nanoparticle probes (see Scheme 1 A in the Supporting Information), which may reduce the fidelity of the nanoparticle probes and hence result in assay errors (see Scheme 1 B in the Supporting Information). Our results demonstrate that this type of exchange does not occur to a significant extent as no detectable Raman signals were observed from the control beads hybridized with the control nanoparticle. In other
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communications search) was used to incorporate Cy3 into the oligonucleotide. For 3’-alkanethiolcapped, TMR, or Cy3.5-labeled oligonucleotides, ultramild base monomers were used to allow for the deprotection reaction under mild conditions. After synthesis, the CPG beads were transferred to a 2 mL reaction vial and treated with 1 mL of 0.05 m potassium carbonate in anhydrous methanol for 4 h at room temperature. Then the supernatant was collected and neutralized with 1.5 mL of 2 m triethylammonium acetate. The oligonucleotides were further Scheme 2. The nucleotide sequences of the target, probe, and capture strands. The synthetic target sequen- purified by preparative reverse-phase HPLC using an ces are adapted from sequence data specific to the respective organism: a) Hepatitis A (HVA), b) Hepatitis B (HVB), c) HIV, d) Ebola virus (EV), e) Variola virus (smallpox; VV), f) Bacillus anthracis (BA), g) Francisella HP ODS Hypersil column tularensis (FT), g) hog cholera segment (VC). Their full sequences are available on the website of the Nation- (5 mm, 250 : 4 mm) with al Center for Biotechnology Information (NCBI): http://www2.ncbi.nlm.nih.gov/Genbank/index.html. Raman 0.03 m triethylammonium labels (R) are TMR:Cy3 = 1:1, TMR, Cy3, TMR:Cy3 = 5:1, 3:1, 1:3, 1:5, and TMR:Cy3:Cy3.5 = 5:5:1 for probe acetate (TEAA, pH 7) and a strands corresponding to targets (a–h), respectively. The control nanoparticles and beads (group i) are 1 % min 1 gradient of 95 % respectively functionalized with HS-T20 and HS-A20 containing no Raman dyes, which is designed to CH 3CN/5 % 0.03 m TEAA at a investigate the potential dye-capped oligonucleotide exchange between different types of nanoparticle flow rate of 1 mL min 1, while probes. monitoring the UV signal of DNA at 254 nm and the absorption of the Raman dyes at words, the HS-T20 oligonucleotide strands on the control 550 nm for Cy3 and 590 nm for Cy3.5. After purification, the 5’probes do not exchange to a measurable extent with the DMT was removed by dissolving the oligonucleotide in an 80 % dye-capped oligonucleotides on the nanoparticle probes acetic acid aqueous solution (reaction: 30 min). After evaporaused for the assay (Figure 3 A and B). tion of the solvent, the oligonucleotide was redispersed in In summary, we have shown that composite Raman 500 mL of water and extracted with ethyl acetate three times labels generated via Raman dye mixing on nanoparticle (300 mL each). After water evaporation, the oligonucleotide probes can be used in a bead-based random-array format. was redispersed in 500 mL of Nanopure water (resistance In principle, a large number of spectroscopically unique 18.1 MW). dyes can be generated via this approach. For a two-dye Functionalization of glass beads: The 250–300-mm diameter system, if five levels of relative peak intensities are assumed, glass beads (Polysciences) were functionalized with capture DNA one can obtain 25 (5 A 5) different combinations for twofollowing a modified literature method.[42] Glass beads (2 g) were peak indexing; after eliminating the redundant and nonexisfirst immersed in 15 mL of piranha solution (H2O2(30 %):H2SO4, tent SERS spectra, 13 distinguishable Raman spectra are 1:3 (v/v); CAUTION: strongly corrosive) in a glass petri dish for available. Similarly, for a six-, ten-, or 14-dye system with 15 min, and then rinsed repeatedly with Nanopure water. The non-overlapping SERS spectra, one could obtain approxiglass beads were then immersed in 15 mL of an aqueous solumately 8 A 103, 5 A 106, and 3 A 109 labels, respectively. Note tion of 1 % (by volume) 3-aminopropyltrimethoxysilane (APS, Althat this Raman-labeling strategy can be readily extended drich) with 0.01 % (by volume) glacial acetic acid in a polystyto RNA and protein detection.[43] rene dish. The sample was rotated on an orbital shaker for 30 min at room temperature. The silane-modified glass beads were then washed three times with H2O, dried under N2, and Experimental Section baked in an oven at 120 8C for 10 min. The beads were promptly treated with 10 mL of 1 mm succinimidyl 4-(maleimidophenyl) DNA synthesis: Raman-dye-labeled oligonucleotides capped butyrate (SMPB, Sigma) in a 4:1 (v/v) EtOH:DMSO solution in a with 3’-protected alkanethiols were synthesized on a 1-mmol glass dish. The coupling reaction was allowed to proceed for scale using standard phosphoramidite chemistry starting with a 2 h. The beads were then rinsed three times with EtOH and thiol-modifier C3 S-S CPG (controlled pore glass) solid support dried under N2. Coupling of freshly deprotected 5’-thiol-DNA to using a commercial synthesizer (Expedite). The Cy3-CE phosphorthe SMPB-modified glass bead surface was carried out using amidite (indodicarbocyanine 3, 1’-O-(4-monomethoxytrityl)-1-O2 mm thiolated DNA in phosphate (NaH2PO4/Na2HPO4) buffer so(2-cyanoethyl)-(N,N-diisopropyl)-phosphoramidite, from Glen Re-
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DTT). The deprotected DNA was purified by passing the solution through a desalting NAP-5 column (Sephadex G-25 medium, DNA grade, Pharmacia Biotech). The Au nanoparticles were functionalized using literature procedures (the Raman-labeled oligonucleotides were used in place of those without labels).[44] Raman spectroscopic measurements: Raman measurements on silver-coated glass beads were performed using a Raman 633 spectrometer coupled to a fiber-optic probe with a 0.65 N.A. microscope objective (Concurrent Analytical, Inc.). The laser (He-Ne, 633 nm) power was 30 mW and the beam diameter was 25 mm. Prior to Raman measurements, the glass beads were treated with silver enhancement solution (Ted Pella, Inc.) for 8 min, washed with Nanopure water, and then dried under N2. The glass beads were spread on a microscope glass slide (25 mm : 75 mm) and studied by Raman spectroscopy. The collection time for each spectrum was typically 0.1 s.
Acknowledgements C.A.M. acknowledges the Homeland Security Advanced Research Projects Agency, the Air Force Office of Scientific Research, the Defense Advanced Research Projects Agency (DARPA), NIH, and the NSF for support of this research. C.A.M. is grateful for a NIH Director’s Pioneer Award.
Keywords: detection · DNA · nanobiotechnology · nanoparticles · Raman spectroscopy
Figure 3. Raman spectroscopic identification of microbeads after hybridization with target DNA and nanoparticles after silver enhancement: A) Random array on a cover slip. B) Raman spectra from individual bead #1 (corresponding to Cy3 labels, associated target: HIV) and #2 (control for oligonucleotide exchange). C) Aligned beads. For ease of interpretation, each type of target was assigned a color that corresponds to different Raman labels. In (A), the row of colored circles from left to right correspond to the respective Raman spectrum of TMR:Cy3 = 1:1, TMR, Cy3, TMR:Cy3 = 5:1, 3:1, 1:3, 1:5, and TMR:Cy3:Cy3.5 = 5:5:1. Both images (A and C) were taken by an optical microscope equipped with a CCD.
lution (0.1 m NaCl, pH 7, 10 mm; for simplicity designated as 0.1 m PBS). The reaction was allowed to proceed overnight at room temperature, followed by rinsing of the beads three times with 0.1 m PBS. The oligonculeotide-functionalized beads were stored in 0.1 m PBS buffer prior to use. Functionalization of Au nanoparticles: Au nanoparticles (13 1.2 nm diameter, 10 nm in particle concentration) were used for all experiments. Prior to the functionalization of Au nanoparticles, the protected thiol functionalities on the Raman-dye-functionalized oligonucleotides were first deprotected by treating the ologonculeotides with 0.1 m dithiothreitol (DTT) in phosphate buffer (0.17 m, pH 8) for 2 h (typically, 10 OD of DNA and 40 mL small 2006, 2, No. 3, 375 – 380
[1] C. A. Mirkin, R. L. Letsinger, R. C. Mucic, J. J. Storhoff, Nature 1996, 382, 607. [2] M. Schena, D. Shalon, R. W. Davis, P. O. Brown, Science 1995, 270, 467. [3] H. Akhavan-Tafti, L. V. Reddy, S. Siripurapu, B. A. Schoenfelner, R. S. Handley, A. P. Schaap, Clin. Chem. 1998, 44, 2065. [4] Y. W. Cao, R. Jin, C. A. Mirkin, Science 2002, 297, 1536. [5] C. A. Afshari, Endocrinology 2002, 143, 1983. [6] D. S. Rickman, C. J. Herbert, L. P. Aggerbeck, Nucleic Acids Res. 2003, 31, 109. [7] M. Han, X. Gao, J. Z. Su, S. Nie, Nat. Biotechnol. 2001, 19, 631. [8] D. R. Walt, Science 2000, 287, 451. [9] A. P. Goodey, J. T. McDevitt, J. Am. Chem. Soc. 2003, 125, 2870. [10] T. Buranda, J. Huang, V. H. Perez-Luna, B. Schreyer, L. A. Sklar, G. P. Lopez, Anal. Chem. 2002, 74, 1149. [11] J. Wang, R. Polsky, A. Merkoci, K. L. Turner, Langmuir 2003, 19, 989. [12] X. Zhao, S. A. Shippy, Anal. Chem. 2004, 76, 1871. [13] K. Sato, M. Tokeshi, T. Odake, H. Kimura, T. Ooi, M. Nakao, T. Kitamori, Anal. Chem. 2000, 72, 1144. [14] R. Elghanian, J. J. Storhoff, R. C. Mucic, R. L. Letsinger, C. A. Mirkin, Science 1997, 277, 1078. [15] M. Bruchez, Jr., M. Moronne, P. Gin, S. Weiss, A. P. Alivisatos, Science 1998, 281, 2013. [16] S. R. Nicewarner-PeÇa, R. G. Freeman, B. D. Reiss, L. He, D. J. PeÇa, I. D. Walton, R. Cromer, C. D. Keating, M. J. Natan, Science 2001, 294, 137. [17] S. Nie, R. Emory, Science 1997, 275, 1102. [18] Y. Cui, Q. Wei, H. Park, C. M. Lieber, Science 2001, 293, 1289.
? 2006 Wiley-VCH Verlag GmbH & Co. KGaA, D-69451 Weinheim
www.small-journal.com
379
communications [19] I. Willner, F. Patolsky, J. Wasserman, Angew. Chem. 2001, 113, 1913; Angew. Chem. Int. Ed. 2001, 40, 2261. [20] H. Mattoussi, J. M. Mauro, E. R. Goldman, G. P. Anderson, V. C. Sundar, F. V. Mikulec, M. G. Bawendi, J. Am. Chem. Soc. 2000, 122, 12 142. [21] C. M. Niemeyer, Angew. Chem. 2001, 113, 4254; Angew. Chem. Int. Ed. 2001, 40, 4128. [22] X. Zhao, R. Tapec-Dytioco, W. Tan, J. Am. Chem. Soc. 2003, 125, 11 474. [23] H. Li, L. J. Rothberg, Anal. Chem. 2004, 76, 5414. [24] S. Howorka, S. Cheley, H. Bayley, Nat. Biotechnol. 2001, 19, 636. [25] R. C. Bailey, J.-M. Nam, C. A. Mirkin, J. T. Hupp, J. Am. Chem. Soc. 2003, 125, 13 541. [26] L. Olofsson, T. Rindzevicius, I. Pfeiffer, M. Kall, F. Hççk, Langmuir 2003, 19, 10 414. [27] B. Dubertret, M. Calame, A. J. Libchaber, Nat. Biotechnol. 2001, 19, 365. [28] R. Chakrabarti, A. M. Klibanov, J. Am. Chem. Soc. 2003, 125, 12 531. [29] A. J. Haes, S. Zou, G. C. Schatz, R. P. van Duyne, J. Phys. Chem. A 2004, 108, 6961. [30] L. R. Hirsch, J. B. Jackson, A. Lee, N. J. Halas, J. L. West, Anal. Chem. 2003, 75, 2377. [31] Y. Sun, Y. Xia, Anal. Chem. 2002, 74, 5297. [32] M. Culha, D. Stokes, L. R. Allain, T. Vo-Dinh, Anal. Chem. 2003, 75, 6196.
380
www.small-journal.com
[33] P. Alivisatos, Nat. Biotechnol. 2004, 22, 47. [34] T. A. Taton, G. Lu, C. A. Mirkin, J. Am. Chem. Soc. 2001, 123, 5164. [35] L. He, M. D. Musick, S. R. Nicewarner, F. G. Salinas, S. J. Benkovic, M. J. Natan, C. D. Keating, J. Am. Chem. Soc. 2000, 122, 9071. [36] J. Reichert, A. Csaki, J. M. Kohler, W. Fritzsche, Anal. Chem. 2000, 72, 6025. [37] X. Zhou, J. Zhou, Anal. Chem. 2004, 76, 5302. [38] J. J. Storhoff, A. D. Lucas, V. Garimella, Y. P. Bao, U. R. MOller, Nat. Biotechnol. 2004, 22, 883. [39] A. M. Michaels, J. Jiang, L. Brus, J. Phys. Chem. B 2000, 104, 11 965. [40] D. Graham, B. J. Mallinder, D. Whitcombe, N. D. Watson, W. E. Smith, Anal. Chem. 2002, 74, 1069. [41] M. J. Hessner, X. Wang, K. Hulse, L. Meyer, Y. Wu, S. Nye, S.-W. Guo, S. Ghosh, Nucleic Acids Res. 2003, 31, 14. [42] L. A. Chrisey, G. U. Lee, C. E. O’Ferrall, Nucleic Acids Res. 1996, 24, 3031. [43] Y. W. C. Cao, R. Jin, J.-M. Nam, S. Thaxton, C. A. Mirkin, J. Am. Chem. Soc. 2003, 125, 14 676. [44] R. Jin, G. Wu, Z. Li, C. A. Mirkin, G. C. Schatz, J. Am. Chem. Soc. 2003, 125, 1643. Received: September 1, 2005 Revised: October 24, 2005 Published online on January 10, 2006
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