Solid-State NMR Study of Cysteine on Gold Nanoparticles - The

Sep 30, 2010 - Department of Chemistry, West Virginia University, Morgantown, West Virginia ... Solid-state NMR spectroscopy is used to characterize t...
1 downloads 0 Views 680KB Size
Download PDF
J. Phys. Chem. C 2010, 114, 18109–18114

18109

Solid-State NMR Study of Cysteine on Gold Nanoparticles Anuji Abraham, Eugene Mihaliuk, Bharath Kumar, Justin Legleiter, and Terry Gullion* Department of Chemistry, West Virginia UniVersity, Morgantown, West Virginia 26506, United States ReceiVed: July 29, 2010; ReVised Manuscript ReceiVed: September 15, 2010

Solid-state NMR spectroscopy is used to characterize the interaction of L-cysteine with gold nanoparticles. The experiments show that there are two types of cysteine in the gold-cysteine complex, with nearly equal populations. We postulate that cysteine forms a two-layer boundary around the gold nanoparticles. The first layer is made of cysteine molecules forming a thiolate bond with the gold surface and having its charged amino and carboxyl groups oriented away from the gold surface. The second layer has its amino and carboxyl groups oriented toward the first layer and its sulfur group oriented away from the gold particles. Introduction Cysteine is the only amino acid having a thiol group and thus is a representative thiol linker, which has the ability to bind to gold either in its molecular form or as part of a peptide or protein. The binding of molecular cysteine to gold substrates has been examined experimentally and through DFT calculations. Compared to the extensively studied alkanethiols on gold, cysteine is an interesting small molecule because it can occur as a zwittterion with charged amino and carboxyl groups and has the potential for hydrogen bonding with its neighbors.1 Most experimental structural studies have been made with cysteine deposited on specific Au(111) and Au(110) substrates, which were characterized by scanning tunneling microscopy.2-11 Scanning tunneling microscopy studies have been performed with substrates under ultrahigh vacuum conditions and with substrates in solution, with very different results. The solution experiments suggest the formation of a sulfur-gold bond at room temperature, whereas the ultrahigh vacuum experiments indicate a physisorbed molecule at room temperature that becomes chemisorbed at higher temperatures. DFT calculations have focused primarily on the sulfur binding at the bridge and fcc hollow sites and the associated molecular conformations.12-14 These calculations have been performed assuming high vacuum conditions and isolated cysteine molecules on the surface. NMR is a good structural tool in the characterization of organic molecules bound to metal surfaces because internal interactions such as the chemical shift, Knight shift, and quadrupolar coupling are sensitive to the electronic environment of the nuclei being examined.15-24 Slichter and co-workers used solid-state NMR to examine the binding of small molecules on platinum, ruthenium, and palladium surfaces.21-23 They were able to provide structural details of the binding of the molecules to the metal surfaces and to show that the metals donated electron density to the absorbed molecule, as indicated by substantial Knight shifts and through relaxation times dominated by the Koringa mechanism. NMR studies of thiols, particularly alkanethiols, bound to gold nanoparticles have been used to examine self-assembly and to provide evidence for the formation of sulfur-gold bonds.15-17 The NMR studies generally find that the 1H and 13C resonances for the respective nuclei proximal to the sulfur show noticeable shifts relative to the unbound * To whom correspondence should be addressed. Tel.: 304-293-3435ext. 6427. Fax 304-293-4904. E-mail: [email protected].

molecules. This is not surprising considering that there should be electron density donated from the gold particles to the chemisorbed molecules. The 1H and 13C resonances of these complexes in solution are generally broader than those of free molecules. The line widths increase with particle size, a fact that has been attributed to longer rotation correlation times, with concomitant shortening of T2.16,24 The rotational correlation time, τc, is proportional to the cube of the radius of the particle, so as the nanoparticles become larger the solution NMR spectra linewidths become broader.25 13C resonances in solution NMR of thiols bound to nanoparticles are unobservable (because of strong broadening) for particles as small as 2 nm. Magic-angle spinning (MAS) 13C NMR experiments on cysteine bound to 6.6 nm gold nanoparticles are reported in this paper. Solid-state NMR was chosen over solution NMR because of the large size of the nanoparticles, which potentially could lead to excessive broadening of solution-state 13C resonances because of long rotational correlation times of the nanoparticles. The NMR results on the solid-phase system indicate chemisorbed and unbound populations of cysteine. The NMR results show that the amino and the carboxyl groups have no significant interactions with the gold surface. Experimental Section Sample Preparation. Gold nanoparticles were prepared26,27 by reducing 120 mL of 0.5 mM gold(III) chloride trihydrate (purchased from Sigma-Aldrich) solution with 0.015 g of sodium borohydride (purchased from Fluka). The resulting solution changed to a ruby red color. 400 mL of 1 mM L-cysteine (U-13C3, 98%; 15N, 98%) (purchased from Cambridge Isotope Laboratories, Inc.) was added to this solution. The mixture was stirred for 30 min and then left undisturbed overnight. Approximately 10 mL of 5 mM solution of aluminum chloride hexahydrate was added to enhance the separation of the nanoparticles from solution via centrifugation. L-cysteine-coated gold nanoparticles were centrifuged at 20,000 G for 30 min on a Fisher Scientific AccuSpin 3R centrifuge. The supernatant was removed and the nanoparticles were washed with water several times before being dried overnight at 333 K. Approximately 1.5 mg of sample was obtained. A sample of gold nanoparticles coated with L-cystine was also prepared, starting with 120 mL of 0.5 mM gold nanoparticles as prepared by the procedure mentioned above. To that solution 400 mL of 1 mM L-cystine (U-13C6, 98%; U-15N2,

10.1021/jp107112b  2010 American Chemical Society Published on Web 09/30/2010

18110

J. Phys. Chem. C, Vol. 114, No. 42, 2010

Figure 1. {1H}13C{15N} REDOR pulse sequence. Continuous-wave decoupling on the proton channel was applied during the dipolar evolution period, t1, and the signal detection period, t2. Following 1 H-13C cross-polarization (CP), a four-rotor-cycle dipolar evolution period lasting for 1.28 ms was applied. 15N dipolar dephasing π pulses with xy-4 phasing were applied to obtain the reduced spectrum, Sr, and the full spectrum, S, was obtained by omitting the 15N π pulses.

98%) (purchased from Cambridge Isotope Laboratories, Inc.) was added. The mixture was stirred for 30 min and then left undisturbed overnight. Ten ml of 5 mM solution of aluminum chloride hexahydrate was added, and the solution was centrifuged at 20,000 G for 30 min. The nanoparticles were washed several times and dried overnight at 333 K. Approximately 1.5 mg of sample was obtained. Solid-State NMR Spectroscopy. MAS NMR spectra were obtained using a triple-channel custom-built spectrometer with a Tecmag Libra pulse programmer and a 3.55 T magnet (proton frequency of 151.395 MHz). The triple-channel probe is a transmission-line design, and it incorporates a Chemagnetics 7.5 mm pencil rotor spinning assembly with a 14 mm long, 8.65 mm inner diameter, 6 turn coil made of 14 gauge wire. Radio-frequency field strengths were 114 kHz for 1H decoupling, 49 kHz for 1H-13C cross-polarization, 49 kHz for 13C pulses, and 48 kHz for the 15N pulses. The 13C and 15N power amplifiers were under active control. All spectra were obtained with 1 s recycle delays and 1 ms matched 1H-13C Hartmann-Hahn cross-polarization transfers. Samples were spun at 3125 Hz and controlled to (0.2 Hz. 13C spectra are referenced such that the 13 C resonance of the methyl carbon of L-alanine occurs at 20.0 ppm. The 15N isotropic chemical shifts are referenced such that the 15N isotropic shift for CH3NO2 is 0 ppm. Rotational-Echo Double-Resonance. REDOR28-32 (Rotational-Echo, Double-Resonance NMR) was used to restore the dipolar couplings between heteronuclear pairs of spins that are usually suppressed by magic-angle spinning. The 13C{15N} REDOR pulse sequence used in this work is shown in Figure 1, and it is the version that has a single refocusing π pulse on the 13C channel. This variant of the experiment was used to minimize recoupling of the 13C-13C homonuclear dipolar interaction that could be caused by a rotor-synchronous 13C π pulse train, in a SEDRA-like (Simple Excitation for the Dephasing of Rotational-echo Amplitudes) fashion,33 for the uniformly 13C-labeled samples. The REDOR experiment is done in two parts, once with rotor-synchronized 15N dipolar dephasing π pulses (dipolar dephased or reduced echo, Sr) and once without (full echo, S). The full echo sequence (no 15N pulses and no net dipolar dephasing) generates an echo of the 13C crosspolarization34 signal at the beginning of data acquisition and serves as a control experiment. The 15N dipolar dephasing π pulses toggle the sign of the 13C-15N heteronuclear dipolar coupling, thereby interfering with the spatial averaging of the dipolar interaction caused by the sample rotation. Consequently, the acquired dipolar dephased NMR signal is reduced in amplitude relative to the full signal. The difference in signal

Abraham et al.

Figure 2. L-cysteine- and L-cystine-coated gold nanoparticles were deposited on mica for size analysis by AFM. Representative AFM images of gold nanoparticles coated with a) L-cysteine and c) L-cystine are shown. The height of individual particles over several images was measured and used to construct size histograms of nanoparticles coated with b) L-cysteine and d) L-cystine. Gold nanoparticles coated with monomeric L-cysteine were characterized by heights of 6.6 ( 2.7 nm (number of particles examined, n, was 968), and gold nanoparticles coated with L-cystine were characterized by heights of 4.0 ( 1.3 nm (n ) 3,918).

intensity, ∆S ) S - Sr, for the observed spin in the two parts of the REDOR experiment is directly related to the corresponding distance to the dephasing spin. The ratio ∆S/S is typically reported. For fixed dipolar evolution times, the stronger the dipolar coupling (shorter internuclear distance) the larger the difference signal and ∆S/S. All REDOR spectra were collected with standard xy-431,32 phase cycling on the dephasing channel. Short dipolar evolution periods of 1.28 ms were purposely used (4 rotor periods) in the 13C{15N} REDOR experiments reported here to assign 13C resonances associated with the Cβ and Cγ carbons of cysteine. Atomic Force Spectroscopy. Cysteine-coated and cystinecoated gold nanoparticles were deposited on freshly cleaved mica that had been pretreated with 20 µL of a 0.01% V/V solution of 300 kDa poly-L-lysine (Sigma Aldrich),35 washed with 200 µL of nanopure water, and dried under a gentle stream of nitrogen. Each sample was imaged in the tapping mode using a MultiMode V atomic force microscope equipped with a Nanoscope V controller and a closed loop vertical engage J scanner (Veeco, Santa Barbara, CA). Images were taken with silicon cantilevers with nominal spring constants of 40 N/m, resonance frequency of ∼300 kHz, and tip radius of ∼10-20 nm. Typical imaging parameters were drive amplitude 150-500 mV with set points of 0.7-0.8 V, drive frequencies of 70-90% below resonance,36 scan frequency of 2 Hz, image resolution 512 by 512 points, and scan size of 2-5 µm. At least 10 images were taken from each deposition for image analysis of aggregate size. Images were analyzed using custom scripts written in MatLab (Mathworks, Natick, MA) equipped with the image processing toolbox. Results Particle Size Determination by Atomic Force Microscopy. To determine their particle sizes, cysteine-coated gold nanoparticles and cystine-coated gold nanoparticles were deposited on mica and images with tapping mode AFM were obtained. Representative images are shown in parts a and c of Figure 2. Because of the finite shape and size of the AFM tip dilating observed lateral dimensions of individual particles, height is the most accurately measured feature and was used to character-

Solid-State NMR Study of Cysteine

J. Phys. Chem. C, Vol. 114, No. 42, 2010 18111

Figure 3. a) {1H}13C{15N} REDOR full spectrum of L-cysteine, (b) {1H}13C{15N} REDOR reduced spectrum of L-cysteine, and (c) {1H}13C{15N} REDOR difference spectrum of L-cysteine. The chemical shift scale was set such that the methyl resonance of L-alanine appears at 20 ppm. Unlabeled peaks are spinning sidebands. 10,000 acquisitions were obtained for the full and reduced spectra. The insert shows the zwitterion structure of L-cysteine.

TABLE 1:

13

C and 15N Isotropic Chemical Shifts and REDOR ∆S/S Ratiosa L-cysteine

sites

δ (ppm) 173.5 ( 0.1 54.0 ( 0.1 27.6 ( 0.1

CR Cβ Cγ C*β C*γ N a

15

-217.4 ( 0.1 13

Au-cysteine ∆S/S

δ (ppm)

∆S/S

0.08 ( 0.01 0.66 ( 0.01 0.06 ( 0.01

173.2 ( 0.1 53.8 ( 0.4 ∼30 66.2 ( 0.4 43.0 ( 0.1 -215.6 ( 0.1

0.09 ( 0.01 0.32 ( 0.01 0.11 ( 0.01 0.49 ( 0.01 0.11 ( 0.01

L-cystine

Au-cystine

δ (ppm)

δ (ppm)

174.8 ( 0.1 51.1 ( 0.1 34.1 ( 0.1

174.8 ( 0.1 51.1 ( 0.2 34.1 ( 0.1

-216.5 ( 0.1

-216.7 ( 0.1

13

The C isotropic chemical shifts are referenced such that the C isotropic shift for the methyl carbon of solid L-alanine is 20 ppm. The N isotropic chemical shifts are referenced such that the 15N isotropic shift for CH3NO2 is 0 ppm.

ize the size of the gold nanoparticles. Several images were captured for each sample and analyzed so that large data sets of measured nanoparticle heights could be compiled into histograms. The size histograms for the cysteine-coated gold nanoparticles and cystine-coated gold nanoparticles are shown in parts b and d of Figure 2, respectively. Gold nanoparticles coated with cysteine had heights of 6.6 ( 2.7 nm (n ) 968), and gold nanoparticles coated with cystine had heights of 4.0 ( 1.3 nm (n ) 3,918). Solid-State NMR Spectroscopy of L-cysteine. 13C{15N} REDOR spectra were obtained on a sample of L-cysteine (U-13C3, 98%; 15N, 98%). The REDOR spectra and chemical structure of cysteine are shown in Figure 3. The REDOR full spectrum shows the 13C resonances of the CR, Cβ, and Cγ carbons are at 173.5 ppm, 54.0 ppm, and 27.6 ppm, respectively. The chemical shifts are summarized in Table 1. Because of the uniform 13C labeling, the linewidths of the 13C resonances are broader than their respective counterparts obtained from a CPMAS (Cross-Polarization Magic-Angle Spinning) spectrum of natural-abundance L-cysteine (not shown), and this broadening comes from the homonuclear dipolar interaction. X-ray crystallography provides CR-N, Cβ-N, and Cγ-N internuclear distances of 2.863 Å, 1.486 Å, and 2.868 Å, respectively.1 These distances correspond to 13C-15N dipolar couplings of 130 Hz for the 13CR-15N spin pair, 929 Hz for the 13 Cβ-15N spin pair and 129 Hz for the 13Cγ-15N spin pair. The

dipolar dephased spectrum obtained with a four rotor cycle dipolar evolution period is shown in part b of Figure 3. The intensity of the 13C resonance for the Cβ carbon is much lower than that in the full spectrum because of the strong 13Cβ-15N dipolar coupling. The intensities of the 13C resonances from the CR and Cγ carbons are only slightly lower than those found in the full spectrum due to the short dipolar evolution period and weaker dipolar couplings. The differences in intensities are perhaps better illustrated in the difference spectrum (top), where the difference intensity arising from the Cβ carbon is high and the difference intensities coming from the CR and Cγ carbons are low. The purpose of using a short four-rotor-cycle dipolarevolution period in the REDOR experiment is further illustrated in the difference spectrum. The carbon directly bonded to the nitrogen makes a strong contribution to the difference spectrum, whereas the Cγ carbon (being farther from the nitrogen) makes a small contribution to the difference spectrum. The ∆S/S ratios for the three carbon sites of L-cysteine are summarized in Table 1. The longer CR-N and Cγ-N distances yield small ∆S/S ratios for their respective 13C signals, whereas the short Cβ-N distance results in a large ∆S/S ratio for the 13C signal arising from the Cβ carbon. The significant contrast in ∆S/S values between the Cβ and Cγ carbons can be used to determine the complex spectral features of the gold-cysteine sample.

18112

J. Phys. Chem. C, Vol. 114, No. 42, 2010

Abraham et al.

Figure 4. a) 13C spectrum of L-cysteine from part a of Figure 3, b) {1H}13C{15N} REDOR full spectrum of Au-cysteine (713,100 acquisitions), c) {1H}13C{15N} REDOR reduced spectrum of Au-cysteine (713,100 acquisitions), and d) {1H}13C{15N} REDOR difference spectrum Au-cysteine. Unlabeled peaks are spinning sidebands.

Solid-State NMR Spectroscopy of L-cysteine on Gold Nanoparticles. Figure 4 shows the REDOR spectra of cysteine on gold nanoparticles (referred to as Au-cysteine) and, for comparison, the spectrum of L-cysteine. The REDOR full spectrum shows the 13C resonance for the CR carbon of the Au-cysteine sample is not shifted from its position found in the L-cysteine spectrum, indicating that the CR carbon does not interact with the gold surface. The most notable feature of the full spectrum is that instead of two distinct resonances in the low frequency region, as in the L-cysteine sample, there is now a wider distribution of resonances with several sharp features. The four-rotor-cycle REDOR experiment helps determine the assignment of the features in the low frequency region of the spectrum. The reduced spectrum is shown in the figure and is used to produce the difference spectrum. The 13C resonances for the Cβ and C*β labeled peaks make strong contributions to the difference spectrum and have large ∆S/S ratios (summarized in Table 1), indicating that these peaks arise from carbon atoms directly bonded to nitrogen. The peak labeled C*γ and the broad low field region designated as Cγ make small contributions to the difference spectrum and have small ∆S/S ratios, so these features arise from carbons directly bonded to the sulfur. A deconvolution is shown in the full spectrum to help visualize the Cγ peak, which shows no sharp feature in the spectrum like the other three peaks. The 13C isotropic chemical shifts of peaks labeled CR, Cβ, Cγ, C*, γ in Figure 4 are listed in Table 1. β and C* Solid-State NMR Spectroscopy of L-cystine. 13C CPMAS spectra were obtained on a sample of L-cystine (U-13C6, 98%; U-15N2, 98%). The CPMAS spectra and chemical structure of cystine are shown in part a of Figure 5. The spectrum is similar to the 13C spectrum of L-cysteine but the 13C resonances for the Cβ and Cγ carbons are shifted relative to the corresponding 13C

resonances in L-cysteine. The 13C isotropic chemical shifts of are summarized in Table 1. Solid-State NMR Spectroscopy of L-cystine on Gold Nanoparticles. Part b of Figure 5 shows the 13C CPMAS spectrum of L-cystine on gold nanoparticles (referred to as Au-cystine). The spectrum lacks the complexity of the Au-cysteine spectrum. Instead, aside from a broad feature (indicated by the dashed curve) underlying the two intense resonances in the low frequency region of the spectrum, the three intense 13C resonances are very similar to those found in solid L-cystine. 15 N CPMAS NMR Spectroscopy. 15N CPMAS spectra were obtained for L-cysteine, Au-cysteine, L-cystine, and Au-cystine. The 15N spectra of the four samples were similar, consisting of single centerbands. The spectra are not shown, but the 15N isotropic chemical shifts are summarized in Table 1. The 15N isotropic shift for the Au-cysteine sample is slightly more shielded than the 15N isotropic chemical shift for L-cysteine. L-cystine

Discussion How cysteine interacts with gold nanoparticles is the primary focus of this work because of the potential to attach cysteineterminated proteins to gold. The REDOR full spectrum of Au-cysteine shown in part b of Figure 4 differs significantly from the REDOR full spectrum of L-cysteine (part a of Figure 3 and part a of Figure 4). Instead of two 13C resonances for the Cβ and Cγ carbons, there four 13C resonances for the Au-cysteine sample in the low frequency region of the spectrum (Cβ,C*,C γ, β Three of these are readily apparent and the fourth is and C*). γ the broad shoulder to the right of the sharper features, the visualization of which is aided by the deconvolution. Assignment of the 13C resonances in the low frequency region of the

Solid-State NMR Study of Cysteine

J. Phys. Chem. C, Vol. 114, No. 42, 2010 18113

Figure 5. a) {1H}13C CPMAS spectrum of L-cystine (7,386 acquisitions), and b) {1H}13C CPMAS spectrum of Au-cystine (235,900 acquisitions). The inset shows the zwitterion form of L-cystine.

spectrum is determined with the REDOR experiment. The ∆S/S ratios for the 13C resonances at 54 ppm (Cβ) and 66 ppm (C*) β are large, indicating that these 13C resonances arise from carbons directly attached to nitrogen. The ∆S/S ratios for the sharp peak at 43 ppm (C*) γ and the broad region centered around 30 ppm (Cγ) are small, indicating that these 13C resonances come from carbons not attached to the nitrogen but, because of their position in the spectrum, from carbon atoms attached to the sulfur atom. The results of the REDOR experiment suggest that there are two types3,6,12 of cysteine molecules interacting with the gold nanoparticles. One type of cysteine has 13C resonances identified as C*β and C*γ that are deshielding by about 12 ppm relative to the same resonances of pure solid L-cysteine. Deshielding of this magnitude has been observed in alkanethiols chemisorbed via a sulfur-gold bond.15 Accordingly, we propose that the C*β and C*γ resonances are associated with cysteine molecules chemisorbed to the gold surface. The other pair of 13C resonances, labeled Cβ and Cγ, are not shifted much from the respective positions found in solid L-cysteine. Hence, the cysteine molecules associated with these 13C resonances could be molecules physisorbed to the gold surface or could be molecules interacting with the chemisorbed cysteine molecules through hydrogen bonding between the charged amino and carboxyl groups of the zwitterions. We believe the latter of the two possibilities is most likely. First, the 13C isotropic chemical shifts of the CR, Cβ, and Cγ peaks in Au-cysteine are very similar to those of solid L-cysteine, which is a network of hydrogen-bonded molecules. Second, the 15N resonance of the Au-cysteine sample is only slightly shifted relative to the 15N resonance of solid L-cysteine. Further support for the second group of molecules interacting with the chemisorbed molecules is that the chemisorbed and unbound molecules appear to contribute equally to the 13C full spectrum, as suggested by an intensity ratio of the 13C resonances of Cβ and C*β that is nearly 1:1. Additional evidence for two types of cysteine molecules in Au-cysteine is provided by thermogravimetric analysis (TGA). TGA was performed on a Au-cysteine sample made of naturalabundance cysteine (no 13C or 15N isotopic enrichment). The TGA results, provided in the Supporting Information, show a component coming off the sample around 110 °C and another component coming off around 280 °C. The masses of the components are comparable. On the basis of the nearly equal populations of the two types of cysteine molecules and the observed 13C and 15N isotropic chemical shifts, we propose the following model. The gold nanoparticles are covered with a layer of chemisorbed cysteine

molecules bound to the gold surface through coordination at the sulfur site. The outer boundary of this layer consists of charged amino and carboxyl groups. A second layer of cysteine molecules forms on top of the chemisorbed layer. The outermost layer has its charged amino and carboxyl groups oriented toward the chemisorbed layer such that the charged groups of each layer are interacting through hydrogen bonding. The outermost boundary of the system consists of sulfur groups. A similar model had been proposed previously based on X-ray photoelectron spectroscopy.6,12 To rule out that the sample made from adding L-cysteine to gold results in the formation of cystine on gold, 13C CPMAS experiments were performed on samples of L-cystine and Au-cystine. The 13C CPMAS spectrum of Au-cystine (part b of Figure 5) is similar to that of L-cystine (part a of Figure 5) and does not show the more complicated low-frequency features found in the Au-cysteine sample. Hence, we rule out the possibility that cystine molecules were formed on the surface of the sample specifically prepared with cysteine. Whereas the 13 C spectrum of the Au-cystine sample is not as detailed as that for the Au-cysteine sample, we suggest a structure of the Au-cystine system. The sharp 13C resonances from the Cβ and Cγ carbons come from cystine molecules that interact with cystine molecules that are either chemisorbed or physisorbed to the gold nanoparticles. The contribution of the chemisorbed/ physisorbed molecules to the low frequency region of the 13C spectrum is the broad feature indicated by the dashed line. Conclusion Solid-state NMR spectroscopy was used to characterize cysteine on gold nanoparticles. The NMR data is consistent with there being two types of cysteine molecules. One type is attached to the gold surface by the sulfur and its charged amino and carboxyl groups are oriented away from the surface. The other type of cysteine molecules forms an outer layer and the charged amino and carboxyl groups of the outer layer interact with their counterparts in the inner layer. Acknowledgment. This work was supported by grant CHE0846583 from the National Science Foundation. We thank Aaron Routzahn for assistance with the TGA measurement. Supporting Information Available: Thermogravimetric analysis data for Au-cysteine. This material is available free of charge via the Internet at http://pubs.acs.org.

18114

J. Phys. Chem. C, Vol. 114, No. 42, 2010

References and Notes (1) Gorbitz, C. H.; Dalhus, B. Acta Crystallogr., Sect. C: Cryst. Struct. Commun. 1996, 52, 1756. (2) Kuehnle, A. Curr. Opin. Colloid Interface Sci. 2003, 14, 157. (3) Kuehnle, A.; Linderoth, T. R.; Hammer, B.; Besenbacher, F. Nature 2002, 415, 891. (4) Kuehnle, A.; Linderoth, T. R.; Schunack, M.; Besenbacher, F. Langmuir 2006, 22, 2156. (5) Uvdal, K.; Bodo, P.; Liedberg, B. J. Colloid Interface Sci. 1992, 149, 162. (6) Dodero, G.; Michieli, L.; De.; Cavalleri, O.; Rolandi, R.; Oliveri, L.; Dacca, A.; Parodi, R. Colloids Surf., A 2000, 175, 121. (7) Dakkouri, A. S.; Kolb, D. M.; Edelstein-Shima, R.; Mandler, D. Langmuir 1996, 12, 2849. (8) Zhang, J.; Chi, Q.; Nielsen, J. U.; Friis, E. P.; Andersen, J. E. T.; Ulstrup, J. Langmuir 2000, 16, 7229. (9) Xu, Q.-M.; Wan, Li.-J.; Wang, C.; Bai, C-Li.; Wang, Z.-Y.; Nozawa, T. Langmuir 2001, 17, 6203. (10) Nazmutdinov, R. R.; Zhang, J.; Zinkicheva, T. T.; Manyurov, I. R.; Ulstrup, J. Langmuir 2006, 22, 7556. (11) Nazmutdinov, R. R.; Manyurov, I. R.; Zinkicheva, T. T.; Jang, J.; Ulstrup, J. Russ. J. Electrochem. 2007, 43, 328. (12) Felice, R. Di.; Selloni, A. J. Chem. Phys. 2004, 120, 4906. (13) Felice, R. Di.; Selloni, A.; Molinari, E. J. Phys. Chem. B 2003, 107, 1151. (14) Hakkinen, H.; Barnett, R. N.; Landman, U. Phys. ReV. Lett. 1999, 82, 3264. (15) Badia, A.; Demers, L.; Dickinson, L.; Morin, F. G.; Lennox, R. B.; Reven, L. J. Am. Chem. Soc. 1997, 119, 11104. (16) Zelakiewicz, B. S.; Dios, A. D. de.; Tong, YuYe. J. Am. Chem. Soc. 2003, 125, 18. (17) Zelakiewicz, B. S.; Lica, G. C.; Deacon, M. L. Tong, YuYe. J. Am. Chem. Soc. 2004, 126, 10053.

Abraham et al. (18) Duncan, T. M.; Yates, J. T.; Vaughan Jr, R. W. J. Chem. Phys. 1980, 70, 1219. (19) Tirendi, C. F.; Mills, G. A.; Dybowski, C. J. Phys. Chem. 1984, 88, 5765. (20) Gay, I. D. J. Catal. 1987, 108, 15. (21) Wang, Po.-K.; Slichter, C. P. Phys. ReV. Lett. 1984, 53, 82. (22) Ansermet, J. PH.; Slichter, C. P.; Sinfelt, J. H. Prog. NMR spectroscopy 1990, 22, 401. (23) Wang, Po.-K.; Ansermet, J. PH.; Rudaz, S. L.; Wang, Z.; Shore, S.; Slichter, C. P.; Sinfelt, J. H. Science 1986, 234, 35. (24) Hostetler, M. J.; Wingate, J. E.; Zhong, C.-J.; Harris, J. E.; Vachet, R. W.; Clark, M. R.; Londono, J. D.; Green, S. J.; Stokes, J. J.; Wignall, G. D.; Glish, G. L.; Porter, M. D.; Evans, N. D.; Murray, R. W. Langmuir 1998, 14, 17. (25) Bloembergen, N. Nuclear Magnetic Relaxation, Frontiers in Physics; Pines, D., Ed.; Benjamin, W.A., Inc.: New York, 1961. (26) Sarangi, S. N.; Hussain, A. P. M.; Sahu, S. N. Appl. Phys. Lett. 2009, 95, 73109. (27) Mayya, K. M.; Anand Gole, A.; Jain, N.; Phadtare, S.; Langevin, D.; Sastry, M. Langmuir 1995, 117, 12537. (28) Gullion, T.; Schaefer, J. J. Magn. Reson. 1989, 81, 196. (29) Gullion, T.; Schaefer, J. AdV. Magn. Reson. 1989, 13, 57. (30) Gullion, T. Concepts in Magnetic Resonance 1998, 10, 277. (31) Gullion, T.; Baker, D. B.; Conradi, M. S. J. Magn. Reson. 1990, 89, 479. (32) Gullion, T.; Schaefer, J. J. Magn. Reson. 1991, 92, 439. (33) Gullion, T.; Vega, S. Chem. Phys. Lett. 1992, 194, 423. (34) Gerstein, B. C. Dybowski, C. R. Transient Techniques in NMR of Solids; Academic Press: New York, 1985. (35) Vesenka, J.; Manne, S.; Giberson, R.; Marsh, T.; Henderson, E. Biophys. J., 1993, 65, 992. (36) Kowalewski, T.; Legleiter, J. J. Appl. Phys. 2006, 99, 064903.

JP107112B