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Langmuir 2008, 24, 9966-9969
Ligand Customization and DNA Functionalization of Gold Nanorods via Round-Trip Phase Transfer Ligand Exchange Andy Wijaya† and Kimberly Hamad-Schifferli*,‡ Departments of Chemical Engineering, Biological Engineering, and Mechanical Engineering, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139 ReceiVed June 18, 2008 Customizable ligand exchange of gold nanorods (NRs) is described. NRs are synthesized with the cationic surfactant cetyltrimethylammonium bromide (CTAB) which is exchanged with thiolated ligands that enable suspension in buffer. Exchange is achieved by a two phase extraction. First, CTAB is removed from the NR-CTAB by extracting the NRs into an organic phase via the ligand dodecanethiol (DDT). The NR-DDT are then extracted into an aqueous phase by mercaptocarboxylic acids (MCA), HS-(CH2)n-COOH (n ) 5, 10, and 15). Ligands can be further customized to thiolated poly(ethylene glycol), PEGMW (MW ) 356, 5000, and 1000). Ligand-exchanged NRs (NR-MCA and NR-PEGMW) are stable in buffer, do not aggregate, and do not change size upon ligand exchange. They can be run in agarose gel electrophoresis with narrow bands, indicating uniform charge distribution and enabling quantitative analysis. DNA functionalization of NR-MCA is straightforward and quantifiable, with minimal nonspecific adsorption.
Gold nanorods (NRs) have been attractive for many biological applications such as gene delivery,1 cell imaging,2 and photothermal therapy.3 However, their ligand functionalization has been problematic for conjugation chemistries. Au NR synthesis results in a double layer of cetyltrimethylammonium bromide (CTAB) for passivation (NR-CTAB), which is problematic for bioconjugation, nonspecific adsorption of DNA, cytotoxicity, and stability. These factors have severely limited the use of NRs in biological applications, especially compared to Au nanoparticles (NPs).4 In order to tune the biological properties of the NR-DNA conjugate and minimize nonspecific adsorption, exchange must permit ligand customization. For gel electrophoresis to assay conjugation and DNA conformation on the NRs, NRs must have uniform charge distribution. There are reports of NR ligand exchange that separately permits conjugation to an antibody2,5,6,7 or gel electrophoresis.8 However, there has not yet been a ligand exchange method which enables ligand customization, biofunctionalization, and gel electrophoresis. Furthermore, customizable ligand chemistry would broadly enhance the versatility of NRs in biological applications. We show here a new approach for ligand exchange that utilizes “round-trip” phase transfer for ligand customization and DNA functionalization (Scheme 1). Resulting NRs are stable in physiological buffers and exhibit narrow bands in gel electrophoresis. DNA functionalization is straightforward, with minimal nonspecific adsorption on the NR. * To whom correspondence should be addressed. E-mail: schiffer@ mit.edu. † Department of Chemical Engineering. ‡ Departments of Biological Engineering and Mechanical Engineering. (1) Chen, C.-C.; Lin, Y.-P.; Wang, C.-W.; Tzeng, H.-C.; Wu, C.-H.; Chen, Y.-C.; Chen, C.-P.; Chen, L.-C.; Wu, Y.-C. J. Am. Chem. Soc. 2006, 128, 3709– 3715. (2) Oyelere, A. K.; Chen, P. C.; Huang, X. H.; El-Sayed, I. H.; El-Sayed, M. A. Bioconjugate Chem. 2007, 18(5), 1490–1497. (3) Huang, X.; Jain, P. K.; El-Sayed, I. H.; El-Sayed, M. A. Nanomedicine 2007, 2(5), 681–693. (4) Murphy, C. J.; San, T. K.; Gole, A. M.; Orendorff, C. J.; Gao, J. X.; Gou, L.; Hunyadi, S. E.; Li, T. J. Phys. Chem. B 2005, 109(29), 13857–13870. (5) Yu, C.; Irudayaraj, J. Biophys. J. 2007, 93(10), 3684–3692. (6) Li, P.-C.; Shieh, D.-B.; Wang, C.-R.; Wei, C.-W.; Liao, C.-K.; Ding, A.A.; Wu, Y.-N.; Poe, C.; Jhan, S. Nat. Preced. 2008. Available from Nature Precedings, http://hdl.handle.net/10101/npre.2008. 1687.1, 2008. (7) Liao, H.; Hafner, J. H. Chem. Mater. 2005, 17, 4636–4641.
Scheme 1. Method for NR Ligand Exchange and DNA Conjugation
Au NRs were synthesized by literature methods.9 Transmission electron microscopy (TEM) was employed to determine mean NR-CTAB dimensions of 43.5 ( 11.9 nm (Figure 2a). Roundtrip ligand exchange first utilizes aqueous-to-organic phase transfer.10,11 NR-CTAB at high concentration (2 × 10-8-5 × 10-8 M) in water was put into contact with dodecanethiol (DDT) (Scheme 1, left). After addition of acetone, NRs were extracted into DDT by swirling the solution for a few seconds, upon which the aqueous phase became clear, indicating that no NRs remained (Scheme 1, middle). Next, organic-to-aqueous phase transfer was performed.12,13 Excess DDT was removed by diluting the DDT coated NRs (NR-DDT) in toluene (1×) and an excess of methanol (5×), and then spun down and resuspended in 1 of mL toluene by brief (8) Hanauer, M.; Pierrat, S.; Zins, I.; Lotz, A.; Sonnichsen, C. Nano Lett. 2007, 7(9), 2881–2885. (9) Sau, T. K.; Murphy, C. J. Langmuir 2004, 20(15), 6414–6420. (10) Gaponik, N.; Talapin, D. V.; Rogach, A. L.; Eychmuller, A.; Weller, H. Nano Lett. 2002, 2(8), 803–806. (11) Jebb, M.; Sudeep, P. K.; Pramod, P.; Thomas, K. G.; Kamat, P. V. J. Phys. Chem. B 2007, 111(24), 6839–6844. (12) Gittins, D. I.; Caruso, F. ChemPhysChem 2002, 3(1), 110–113.
10.1021/la8019205 CCC: $40.75 2008 American Chemical Society Published on Web 08/22/2008
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sonication. The NR-DDT in toluene was then added to 9 mL of 0.01 M mercaptocarboxylic acid (MCA) in toluene at elevated temperature and vigorously stirred. Three MCA ligands (mercaptohexanoic acid (MHA), mercaptoundecanoic acid (MUDA), and mercaptohexadecanoic acid (MHDA)) were used. Operating temperatures were 95 °C for MHA and 70 °C for MUDA and MHDA.14–16 Reflux and stirring continued until visible aggregation was observed (within ∼15 min), and then the solution was allowed to settle and cool to room temperature. Aggregation indicated that NRs were successfully coated by MCA (NR-MCA), which are insoluble in toluene. The aggregates were washed 2× with toluene via decantation and then once with isopropanol to deprotonate the carboxylic acid. The aggregates spontaneously redispersed in 1× tris-borate-EDTA buffer (TBE) and were no longer soluble in toluene (Scheme 1, right), suggesting residual DDT on the NR is minimal. Once resuspended in TBE, NR-MCA could have their MCA coating optimized, be ligand-exchanged with another species, or be conjugated to DNA (Scheme 1). We incubated NR-MCA with 1 mM MCA in H2O or a H2O/ethanol mixture for further optimizing the MCA coating. We also performed further ligand exchange of NR-MHA by incubating them in 1 mM aqueous solution of poly(ethylene glycol) (PEG)-thiols (HS-PEG356, HSPEG1000, and HS-PEG5000). Lastly, we conjugated NR-MHA with fluorescently labeled thiolated DNA 40-mers (5′ HS-TTTTT TTTTT TTTTT TTTTT TCGGC CCGTA TAATT-TMR 3′) using the charge screening method for DNA functionalization of Au NPs.17 The width of the longitudinal surface plasmon resonance can be used to directly probe the stability and aggregation of NRs. UV-vis spectra of ligand-exchanged NRs with MCA show shifts of the longitudinal plasmon with no significant broadening, indicative of no aggregation (Figure 1a). The peak shift is linear with chain length (Figure 1a, inset) as expected from the change in refractive index with increasing alkyl chain length.18–20 The longitudinal plasmon of NRs functionalized with PEG-SH (Figure 1b) also showed no significant broadening. We found that the ligand-exchanged NRs were stable even after 3 or 4 months of storage at high concentration (∼2 × 10-8 M), and the plasmon peaks exhibited no significant changes in peak width or position (Figure 1c). Heating gold nanoparticles near the boiling point of the solvent in the present of surface-active ligands such as alkanethiols over a certain period of time may result in a reduction in the average size of the particle due to digestive ripening.21 TEM imaging was used to probe any change in size of the NR dimensions upon ligand exchange (Figure 2). TEM size analysis determined mean NR-MHA dimensions of 43.2 × 11.8 nm (Figure 2b), indicating no significant size change with ligand exchange. Fourier transform infrared (FTIR) spectroscopy was performed to probe the nature of the ligands on the surface of gold nanorods before and after the ligand exchange. FTIR spectroscopy of NR-CTAB (Figure 3a, black) showed peak at (13) Aldana, J.; Lavelle, N.; Wang, Y. J.; Peng, X. G. J. Am. Chem. Soc. 2005, 127(8), 2496–2504. (14) Hostetler, M. J.; Templeton, A. C.; Murray, R. W. Langmuir 1999, 15(11), 3782–3789. (15) Shon, Y. S.; Lee, T. R. J. Phys. Chem. B 2000, 104(34), 8192–8200. (16) Garg, N.; Carrasquillo-Molina, E.; Lee, T. R. Langmuir 2002, 18(7), 2717–2726. (17) Zhang, J.; Song, S. P.; Wang, L. H.; Pan, D.; Fan, C. Nat. Protoc. 2007, 2(11), 2888–2895. (18) Ehler, T. T.; Malmberg, N.; Noe, L. J. J. Phys. Chem. B 1997, 101(8), 1268–1272. (19) Templeton, A. C.; Pietron, J. J.; Murray, R. W.; Mulvaney, P. J. Phys. Chem. B 2000, 104(3), 564–570. (20) Sun, Y. G.; Xia, Y. N. Anal. Chem. 2002, 74(20), 5297–5305.
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Figure 1. UV-vis spectra of NRs upon ligand functionalization. (a) UV-vis spectra of NR-CTAB (black) before and after ligand exchange with MCA, NR-MHA (red), NR-MUDA (blue), and NR-MHDA (green) [inset: longitudinal surface plasmon peak as a function of ligand length for SH-(CH2)n-COOH]; (b) UV-vis spectra of NR-MHA (black) before and after functionalization with PEG-SH, NR-PEG356 (red), NR-PEG1000 (green), NR-PEG5000 (blue); and (c) UV-vis spectra of ligand-exchanged NRs right after exchange (solid line) or after 3-4 months of storage at high concentration (dashed line).
958 cm-1 (arrow, 1) due to the quarternary amine stretch of CTAB (Figure 3a, green). NR-MHA (Figure 3b, red) exhibited a COO- stretch (1585 cm-1, arrow, 2), shifted from the COOH stretch (1690 cm-1, arrow, 3) for MHA (Figure 3b, blue), which is due to deprotonation of carboxylic acid. NR-MHA lack a S-H stretch (2613 cm-1, arrow, 4) but still have a C-S stretch (706 cm-1, arrow, 5).22–25 Gel electrophoresis was performed with 0.5% agarose in 0.5× TBE (Figure 4a). NR-CTAB aggregate in buffer and do not move from the well (lane 1). NR-MHDA (lane 2), NR-MUDA (lane 3), and NR-MHA (lane 4) all run in the positive direction, indicating that NRs are negatively charged. The increasing trend in mobility is most likely due to a decrease in the hydrodynamic radius (RH) of the NRs as a result of decreasing alkyl chain (21) Prasad, B. L. V.; Stoeva, S. I.; Sorensen, C. M.; Klabunde, K. J. Langmuir 2002, 18(20), 7515–7520. (22) Nelson, G. J. Lipids 1968, 3(1), 104. (23) Xing, S. X.; Chu, Y.; Sui, X. M.; Wu, Z. S. J. Mater. Sci. 2005, 40(1), 215–218. (24) Morales-Cruz, A. L.; Tremont, R.; Martinez, R.; Romanach, R.; Cabrera, C. R. Appl. Surf. Sci. 2005, 241(3-4), 371–383.
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Figure 3. FTIR spectra of (a) CTAB (green) and NR-CTAB (black) before ligand exchange and (b) MHA (blue) and NR-MHA (red) after ligand exchange.
Figure 2. TEM images and size analyses of (a) NR-CTAB before ligand exchange and (b) NR-MHA after ligand exchange.
length. NR-PEG356 (lane 5), NR-PEG1000 (lane 6), and NRPEG5000 (lane 7) all ran in the positive direction, which could be due to residual MHA on the NR surface. Mobility decreased with increasing PEG chain length, most likely due to the increase in RH with longer PEG. Furthermore, bands of all of the ligand functionalized NRs were narrow, exhibiting clear mobility shifts with surface functionalization, enabling reliable quantitative Ferguson analysis to find the hydrodynamic radius8,26 and ζ-potential.27 (25) Hostetler, M. J.; Stokes, J. J.; Murray, R. W. Langmuir 1996, 12(15), 3604–3612. (26) Zanchet, D.; Micheel, C. M.; Parak, W. J.; Gerion, D.; Alivisatos, A. P. Nano Lett. 2001, 1(1), 32–35.
Figure 4. Gel electrophoresis (0.5% agarose in 0.5× TBE) of (a) before and after ligand exchange: lane 1, NR-CTAB; lane 2, NR-MHDA; lane 3, NR-MUDA; lane 4, NR-MHA; lane 5, NR-PEG356; lane 6, NRPEG1000; lane 7, NR-PEG5000. (b) DNA functionalization of NR-MHA (UV image): lane 8, NR-MHA; lane 9, NR-MHA incubated with nonthiolated DNA; lane 10, NR-MHA incubated with thiolated DNA; lane 11, nonthiolated DNA; lane 12, thiolated DNA. All DNA was functionalized with a 3′ TMR (TAMRA).
After ligand exchange, the resulting NR-MHA was conjugated to HS-DNA as described above. Figure 5a shows the UV-vis spectra of before and after DNA conjugation. DNA conjugation (Figure 5a, red solid line) did not broaden or shift the longitudinal surface plasmon resonance (SPR) significantly relative to the NR-MHA peak (black dashed line), indicating no aggregation after DNA conjugation. Furthermore, the UV-vis spectra of the NR-DNA conjugate were taken 7 days after the conjugation was performed, indicating its long-term stability. Gel electrophoresis was performed to confirm the DNA-NR conjugation via Au-S bonding. NR-DNA conjugates (Figure 4b, lane 10) ran slower relative to NR-MHA (lane 8), indicating a RH increase. Bands are narrow enough to permit Ferguson analysis of the NR-DNA. The narrow band also indicates no aggregation and a uniform
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Figure 5. DNA conjugation. (a) UV-vis spectra of NR-MHA (black dashed line), NR-MHA + nonthiolated DNA (black solid), and NRMHA + thiolated DNA (red line). (b) Fluorescence spectroscopy of supernatants of NRs conjugated with thiolated DNA (red solid line) or nonthiolated DNA (black solid line). Fluorescence spectroscopy of controls (DNA without NRs) of thiolated DNA (red dashed line) or nonthiolated DNA (black dashed line). (c) Fluorescence spectroscopy of supernatants of MCH treated NRs conjugated with thiolated DNA (red solid line) and nonthiolated DNA (black line). Spectrum of supernatant of NR conjugated with thiolated DNA without MCH treatment (red dashed line).
charge distribution. A control with fluorescently labeled nonthiolated DNA exhibited no significant mobility shift, indicating minimal nonspecific adsorption (lane 9). The fluorescent DNA band of NR-DNA (lane 10) decreases in intensity compared to a control sample of the thiolated DNA by itself (lane 12) while the nonthiolated DNA does not (lane 9 vs lane 11), also supporting covalent attachment. Fluorescence spectroscopy also indicated NR-DNA conjugation. Supernatant fluorescence decreased (Figure 5b, red solid line) from its original value (red dashed line), indicating that DNA was removed from solution by conjugation to the NRs. Nonthiolated DNA showed a smaller
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decrease (Figure 5b, black line, solid vs dashed line), indicating that covalent conjugation was favored over nonspecific adsorption. The NR concentration was estimated using the extinction coefficients of the longitudinal plasmon band peak, ε ) 4.6 × 109 M-1 cm-1.28 Fluorescence spectra quantified ∼41 DNA/ NR. NR-DNA bands are not at the same position as the free DNA bands (lanes 9 and 10), so the band can be cut to extract the purified sample. Conjugated DNA was also quantified by displacement with mercaptohexanol (MCH).29 Purified NR-DNA were incubated in 1 mM MCH overnight, displacing the DNA from the NRs. Free DNA was separated from the NRs by centrifugation and quantified by fluorescence (Figure 5c, red solid line). As a control, a fluorescence scan was taken of the supernatant of the same NR-DNA solution without MCH treatment (red dashed line). The scan was taken 9 days after the conjugation, indicating there was no significant detachment of DNA from NR-DNA conjugates over this period of time. The gel-purified NRs incubated with nonthiolated DNA were also treated with MCH, and the supernatant exhibited no significant fluorescent peak (black solid line), indicating minimal nonspecific adsorption. Quantification of fluorescence determined ∼28 HSDNA/NR. This DNA loading translates to approximately 2 pmol DNA/cm2, which is the same order of magnitude for DNA on gold nanowires30 and nanoparticles.31 These results show that ligand exchange of Au NRs by the round-trip phase transfer method enables ligand customization and straightforward DNA conjugation. Ligand-exchanged NRs have uniform charge distribution, enabling their stability in physiological buffers and quantitative analysis by gel electrophoresis. This method for modifying the surface chemistry of gold NRs will enhance their versatility in biological applications such as therapy, sensing, and imaging. Acknowledgment. We would like to thank the MIT Center for Materials Science and Engineering for use of their facilities. Supporting Information Available: Additional TEM images and size histograms, and FTIR spectra. This material is available free of charge via the Internet at http://pubs.acs.org. LA8019205 (27) Park, S.; Hamad-Schifferli, K. J. Phys. Chem. C 2008, 112, 7611–7616. (28) Orendorff, C. J.; Murphy, C. J. J. Phys. Chem. B 2006, 110(9), 3990– 3994. (29) Park, S.; Brown, K. A.; Hamad-Schifferli, K. Nano Lett. 2004, 4(10), 1925–1929. (30) Mbindyo, J. K. N.; Reiss, B. D.; Martin, B. R.; Keating, C. D.; Natan, M. J.; Mallouk, T. E. AdV. Mater. 2001, 13(4), 249. (31) Hurst, S. J.; Lytton-Jean, A. K. R.; Mirkin, C. A. Anal. Chem. 2006, 78(24), 8313–8318.
