Generalized Approach to SERS-Active Nanomaterials via Controlled

Jul 7, 2009 - Gary B. Braun,† Seung Joon Lee,† Ted Laurence,‡ Nick Fera,† Laura ... Chemistry, Materials, Earth and Life Sciences Directorate,...
0 downloads 0 Views 3MB Size
Download PDF
13622

J. Phys. Chem. C 2009, 113, 13622–13629

Generalized Approach to SERS-Active Nanomaterials via Controlled Nanoparticle Linking, Polymer Encapsulation, and Small-Molecule Infusion Gary B. Braun,† Seung Joon Lee,† Ted Laurence,‡ Nick Fera,† Laura Fabris,† Guillermo C. Bazan, Martin Moskovits,*,† and Norbert O. Reich*,† Department of Chemistry and Biochemistry, UniVersity of California, Santa Barbara, California, 93106, and Chemistry, Materials, Earth and Life Sciences Directorate, Lawrence LiVermore National Laboratory, LiVermore, California 94550 ReceiVed: April 13, 2009; ReVised Manuscript ReceiVed: June 18, 2009

Over the past decade the emphasis on single-molecule sensitivity of surface-enhanced Raman spectroscopy (SERS) has brought to prominence the special role played by so-called SERS hot spots, oftentimes nanometerscale junctions between nanoparticles (NPs). In this report, molecular linkers are used to mediate the assembly of NPs to dimers and small clusters. When the SERS enhancement is optimized, the aggregation process is quenched by polymer and protein stabilizers that subsequently act as encapsulants resulting in SERS substates with unprecedented enhancement uniformity, reproducibility, and stability. The polymer-stabilized NP junctions were then imprinted with a variety of small molecules that permeated the polymer coat and displaced the linker from the hot spot. The average SERS enhancement of these SERS “nanocapsules” was found to be at least 300× greater than for single NPs, while the Raman/Rayleigh scattering ratio was 104 higher for linked NPs versus nonoptimized aggregates. Single-particle statistics showed that almost every nanocapsule produced intense SERS, suggesting that they are NP dimers and small clusters with the probe molecule resident in a hot spot. Nanocapsules were functionalized and shown to compete successfully with fluorescence imaging in multiplexed identification of cancer cell epitopes at the single-cell and single-nanotag level. Introduction Surface enhanced Raman scattering spectroscopy (SERS) is a highly enhanced form of Raman spectroscopy enabled exclusively by nanostructured systems which, as a vibrational spectroscopy, can potentially lead to molecular identification of species positioned on or near certain metallic nanostructures, often at nanosized junctions or sharp protrusions.1 The utility of SERS has been demonstrated for the detection of small molecules, nucleic acids, proteins, and cells through direct enhancement of the Raman scattering of the analyte molecule or by indirectly detecting prelabeled SERS nanoparticle tags.2-8 Particularly noteworthy are efforts to detect glucose levels in blood noninvasively3 and the application of nanoparticle conjugates for tumor imaging.8,9 Recently, the engineering of nanoscale junctions has become a key goal in designing high-quality SERS substrates, as they have been shown to generate the most enhancing fields.10,11 Although many promising structures have been proposed, creating them with near-optimal SERS activity and sufficient yield in a facile, reproducible, and scalable manner and with uniform response toward a library of molecular adsorbates, has not been achieved. Several groups have tackled the problem of reducing the ad hoc nature of the SERS signal following aggregation of colloidal nanoparticles. These efforts often build upon promising dimerlike “cluster” architectures to achieve highly intense SERS. Su et al. developed a system of adding dye during nanoparticle growth at elevated temperatures to produce Ag assemblies which were then protein coated and conjugated to antibodies for * To whom correspondence should be addressed. E-mail: reich@ chem.ucsb.edu (N.O.R.); [email protected] (M.M.). † University of California, Santa Barbara. ‡ Lawrence Livermore National Laboratory.

multiplexed immunoassays.8 Recently, Brown et al. used roomtemperature aggregation with NaCl followed by the synthesis of dye-tagged silica-coated nanoparticles to generate glasscoated clusters.10 We have previously explored molecular crosslinking strategies using bifunctional dithiol linkers to manage the aggregation of Ag nanoparticles,12,13 and DNA hybridization to assemble silver nanoparticles onto a DNA-coated metal surface.14 However, these strategies require the analyte or tag to be present during aggregation. This fact precludes a range of applications, including sensing small molecules out of the environment or the systematic analysis of engineered hot spots using probe molecules. In this report we describe a new set of linking, signaloptimization, encapsulation, and analyte infusion strategies that, used together, produce a new family of encapsulated SERSactive clusters (Figure 1). The two major improvements here over previous reports on nanoparticle aggregates that had been produced either purposely or inadvertently is, first, the development of a kinetically optimized nanoparticle linking/quenching protocol in which the SERS signal itself is used as the optimization parameter through each surface modification step and, second, a polymer-quenching step to structurally stabilize the nanoaggregate while allowing small molecules access to the junction regionsthe hot spotsat a later time. The materials show near-optimum, highly reproducible enhancements and great stability against further aggregation during the subsequent chemical and physical manipulations, characteristics that can be valuable in fundamental SERS studies as well as in a wide range of tagging, sensing, and analysis applications. The effort is generalized to a solution-based synthesis that is scalable to large-quantity preparation of encapsulated SERS clusters (nanocapsules) composed primarily of dimers and small clusters, which can be infused with small molecule probes at a later time.

10.1021/jp903399p CCC: $40.75  2009 American Chemical Society Published on Web 07/07/2009

Generalized Approach to SERS-Active Nanomaterials

J. Phys. Chem. C, Vol. 113, No. 31, 2009 13623

Figure 1. SERS nanocapsule synthesis. (a) Ag nanoparticles are cross-linked with the bifunctional linker 4-aminobenzenethiol (ABT, blue) or (b) 1,6-hexamethylenediamine (HMD, black), each then coated with a layer of PVPA. PEG thiol or streptavidin and bovine serum albumin proteins are then adsorbed (not shown). In b, the SERS tag (red) is infused through the polymer coat. The inset represents SERS from tags in the junction. (c) Structures of 4-mercaptobenzoic acid (MBA, red), 4-aminobenzenethiol (ABT), 5,5′-dithiobis(2-nitrobenzoic acid) (DTNB), biphenyl-4,4′-dithiol (BPDT), 2-mercaptopyridine (MPY), 2-naphthalenethiol (NPT), 4-amino-5-(4-pyridyl)-4H-1,2,4-triazole-3-thiol (APTT), phthalazine (PHTH), and 4-Amino-4′-dimethylamino azobenzene (DAAB). (d) Color-coded spectra of SERS nanocapsules with MBA and DTNB prepared as in b and ABT as in a.

We describe two strategies involving three stages of linker addition, polymer quenching, and filling the hot spot with analyte. Figure 1a shows schematically the synthesis of tagged clusters using a Raman active linker that possesses terminal thiolate or amino groups (4-aminobenzenethiol, ABT), which is used to optimize the overall approach, since the bifunctional linkers, which act as the analyte or Raman tag, automatically position themselves in hot spots as they form. After sufficient aggregation, polymer is added to quench the reaction by sterically protecting the clusters against further growth. Figure 1b illustrates a much more versatile strategy for SERS substrate preparation, which is the primary focus of this study. In this approach nanoparticle dimers and small aggregates are assembled by using a diamine aggregation agent (1,6-hexamethylenediamine, HMD) which can be displaced at a later stage by a molecule of interest without compromising the integrity of the prelinked nanocapsules. Importantly, the diamine has a small Raman cross-section and weak binding affinity, allowing the postinfusion and sensing of multiple Raman scatterers that may be inefficient aggregation agents in their own right. The polymer-quenching process stabilizes the hot spots between particles which are accessible in a subsequent infiltration step

by small molecule probes. Figure 1c shows a number of taggants infused into these prelinked nanocapsules (see also Supporting Information). Because the taggants used in building SERS nanocapsules must facilitate the tracking and multiplexing applications we have in mind, they must possess the following characteristics: (i) SERS spectra with well-defined Raman vibrational signatures suitable for multiplexed optical biosensing applications in which several species must be confidently detected simultaneously; (ii) large Raman cross sections to facilitate (near) single-particle detection; and (iii) the ability to produce a time-invariant SERS signal by passivating and excluding other molecules from entering the SERS hot spot. Likewise, the coating is selected to have one or more of the following properties: (i) a moderately compact architecture; (ii) an ability to sterically and/or electrostatically protect clusters from aggregation in high ionic strength buffers; and (iii) an appropriately useful chemical structure that can provide conjugation handles for subsequent functionalization with receptors or biomolecules allowing the nanocapsule to interact specifically with a desired chemical or biological target.

13624

J. Phys. Chem. C, Vol. 113, No. 31, 2009

Braun et al.

Experimental Methods SERS nanocapsules with one or a few interparticle junctions were synthesized by self-assembly induced by a combination of phosphate buffer destabilization and addition of either of two linker molecules (Figure 1a,b). The first strategy utilized ABT, which is a commonly used Raman tag possessing a large Raman cross-section, and the second was HMD, which produces a rather weak Raman spectrum. The clusters were then diluted in excess polyvinylpyrrolidone (PVP) or polyvinylpyrrolidone-poly(acrylic acid) (PVPA) followed by PEG-thiol or bovine serum albumin (BSA) and Cy3-streptavidin (STV) for stability in high salt systems and for affinity labeling. Notably, the Cy3 dye excitation is nonresonant with 633 nm excitation and therefore does not impact the background signal levels but rather serves as a particle label during fluorescence imaging. HMD-linked nanocapsules were exposed to a variety of Raman active molecules to explore their use as a generic sensing or labeling material. Using a 96-well plate, 50-100 µL of 500 pM colloid was placed in each well and interrogated with 0.1-1 mW of 633 nm laser light focused through a 10× objective. Spectra were recorded with 1 s acquisition, yielding ∼104 counts/s · mW for the benzene Raman band at ∼1050 cm-1 (methanol yields ∼40 counts/ s · mW). ABT Direct Linking. Ag nanoparticles were prepared by Lee and Meisel’s method.15 To find the optimum synthesis conditions, a well-plate format was used in conjunction with a confocal Raman system setup in a backscattering geometry. Many synthesis conditions were screened by systematically changing the reagents, the sequence of their addition, and the incubation times. Raman spectra were collected at 5 s intervals, allowing the effect of aggregation rate on SERS activity to be determined. An example data set collected from a system consisting of Ag nanoparticles linked by ABT is shown in Figure 2a. Ag colloid (100 µL) in 10 mM sodium phosphate buffer (PB) was mixed with various amounts of ABT (50 µM) in methanol to cause additional aggregation, followed by addition of PVPA to 0.2 mg/mL and dilution to approximately twice the volume with deionized (DI) water. After 3 min polyethyleneglycol (PEG) was added at 30 µM, centrifugation was performed at up to 1500g and redispersion in 5% 8000 Da PEG in DI water to the initial volume, or into pure DI water for TEM imaging. For PEI polymer overcoating the PVPA clusters were first brought up in 3% poly(acrylic acid) in 5% 8000 Da PEG in DI water (note there is a color change to gray), centrifuged, and redispersed in 3% polyethyleneimine (PEI) in DI water (note the colloid color returns to green). This was followed by two wash cycles into in 5% 8000 Da PEG in DI water, or pure DI water for TEM. 1% tetraethyl orthosilicate (TEOS) solution in water with 10 mM NH4OH were combined with PEI-coated clusters to generate silica coated clusters, returned to pH 7 with 100 mM PB, and further cleaned by centrifugation and washing with DI water. Prelinked Nanocapsules. Ag starting colloid was centrifuged and brought up at 0.3 absorption units (1 mm path length) in PB then combined with a small volume of ∼1 mM HMD in DI water (up to 200 µM final concentration) to initiate aggregation. PB and HMD concentrations were combined in various ratios to optimize the cluster formation terminated 90-300 s after HMD addition. This process generally is required to be repeated between citrate Ag batches and is between ∼20 mM PB with 0 µM HMD to 0.1 mg/mL total protein. Raman-active molecules in methanol were added after at least 3 min for polymer-coated samples and >15 min for PVP/STV, at 0.001-1 mM final analyte concentration depending on the molecular affinity. Minimal color change is generally observed if the protein coating is stable. After 30 min the products were centrifuged at 3K RPM several times and washed with PB, or phosphate buffered saline (PBS) with 0.1% BSA for STV samples. Cell Binding. Biotinylated TAT peptide (for HeLa cell line CCL2) or antibodies (for B-cell line CCL159) were added to Cy3-STV-coated nanocapsules at a concentration of ∼3 nM and incubated for 30 min on ice. A portion of this product was added to cells suspended in PBS and incubated at room temperature. After 30 min the cells were centrifuged, washed with PBS, and placed on a glass slide with glass coverslip. The cells were imaged using an Olympus BX-41 microscope equipped with a Hg lamp, FITC, and Cy3 filter sets, and a HORIBA Jobin Yvon

Generalized Approach to SERS-Active Nanomaterials

J. Phys. Chem. C, Vol. 113, No. 31, 2009 13625

Aramis Raman system equipped with 10×, 50×, and 100× objectives, a He-Ne 633 nm laser and mapping stage. Results and Discussion Linking and Polymer Quenching. To examine the kinetics of linking16 and optimize the SERS we first followed the scheme shown in Figure 1a, where ABT, a linker with a large Raman cross-section,17 is used to aggregate the Ag nanoparticles. The assembly of the direct-linked ABT nanocapsules (Figure 1a) was optimized by monitoring the time evolution of the SERS intensity for varying concentrations and incubation times of the nanoparticles, buffer, adsorbates, and polymer (Figure 2a). Following the addition of ABT the SERS signal increases dramatically after a short induction period. SERS intensity reaches a constant value after dilution with polymer solution. At this stage, centrifugation results in a redispersible product. In contrast, if no polymer is added the aggregation process continues unchecked, forming an insoluble macroscopic precipitate. Polymer addition was also followed by protein adsorption before centrifugation, which further functionalizes the product for biological assays. Interestingly in some cases, the SERS intensity rises to a maximum and then drops to a constant lower value. While this decrease may result from aggregation kinetics, a similar effect was previously observed18,19 and explained to arise from the simultaneous increase in signal intensity due to the increasing surface concentration of adsorbate and the accompanying field depolarization effect which is greatest when the surface species density is near monolayer coverage. The PVP, PEG thiol, and protein coating protocols were optimized by probing the constancy of the SERS intensity and the dispersion stability following various centrifugation and sonication procedures interspersed with periodic SERS measurements. Stability was found to improve when the linker coverage on the nanoparticle was kept low; a concentration yielding kinetics between the green and blue curves Figure 2a was optimal, possibly as a result of the increased availability of surface to the polymer and protein which competes with the linker for Ag surface. Following this reasoning, PB was added prior to the linker to coordinate and increase the propensity for aggregation, lowering the number of linkers necessary for forming clusters, knowing that phosphate is more easily exchanged by both polymer and protein when they are added in turn. Cluster size analysis was carried out on TEM images from samples quenched at optimal SERS intensity (Figure 3), along with slight color change and UV-visible absorption spectra as an indicator for degree of aggregation (Supporting Information). The cluster distribution of the polymer coated product as shown by TEM analysis typically comprises ∼45% monomers, 25% dimers, 10% trimers, 5% tetramers, and decreasing amounts of larger cluster sizes. Strategies that are effective for collecting individual cluster fractions will be described in a separate report. Briefly, samples were centrifuged to the point where the yellowcolored supernatant, which consists primarily of monomer, gives only weak SERS. The supernatant is discarded, the pellet is redispersed and the resulting solution is passed through a 0.22 µm syringe filter to remove large clusters. The PVP and PEG thiol were chosen for their ability to quench the aggregation by a combination of coordination to, and dilution of, the particles and linker, imparting long-term stability during storage (Supporting Information). The PVP is expected to work through a large number of weakly coordinated sites on the cluster surface, being susceptible to displacement

Figure 3. Cluster population. (a) TEM analysis of 200 nanocapsules from a sample quenched at near-peak SERS intensity, showing a high population of dimers and trimers. A power law curve is overlaid as guide to the eye. Inset: TEM field view. (b) close-up images of a polymer-coated dimer and pentamer and silica-coated ABT-linked particles.

Figure 4. Histogram of intensities per event where a nanoparticle or nanocluster passes through the focal volume of the confocal instrument. Raman/Rayleigh (IRm/IRy) and Raman/background (IRm/IRbkgd) ratios are simultaneously measured from freely diffusing Ag nanoparticles randomly aggregated with 10 µM MBA (red) and for optimized 2 µM ABT/PVPA nanocapsules (green) isolated by centrifuging. Color intensity is proportional to the number of events. The two materials have a difference of 104 in the ratio of Raman/Rayleigh, and greater signal-to-noise (Raman/background).

by taggant molecules, while the PEG is expected to bind irreversibly at one end, taking up a large footprint per attachment point and leaving room for subsequent small-molecule infusion and adsorption. We also note that single-strand DNA alkyl thiolates, as pioneered by Mirkin,20 and carboxy- or aminoterminated PEG thiolates ∼3-5 kDa9 are also excellent stabilizers for the metal clusters while allowing analyte infusion and functionality (data not shown). Furthermore, as an alternative to porous polymer/protein coating, clusters were further coated with cationic PEI and a silica shell was grown with full retention of the SERS activity from the ABT embedded within (Figure 3b, Supporting Information), as a route to a temperature- and solvent-resistant SERS tag.21,22 The intense SERS observed during the aggregation process can also be used to analyze the impact of aggregation on SERS

13626

J. Phys. Chem. C, Vol. 113, No. 31, 2009

enhancement. In Figure 2b the approximate intensity advantage of nanoparticle clusters over monomers is estimated by referencing an actively aggregating system to one which is at an early stage of aggregation, and hence dominated by Ag nanoparticle monomers. Upon ABT addition, the second sample showed a delayed onset of SERS intensity growth, due to a lower concentration of aggregates attributed to a combination of decreased electrostatic screening and nanoparticle collision rate. The rate of ABT adsorption onto the nanoparticles is assumed to be similar in the two systems (i.e., we assume that the ABT concentration is large enough for the system to be in the saturation region of the isotherm in both cases). The timevarying SERS intensity is presented in Figure 2b along with the SERS intensity ratio between the two systems, corrected for initial nanoparticle concentration. The observed maximum ratio ∼300 suggests that the increase in SERS enhancement for the clusters is at least that much larger than for an equal number of isolated nanoparticles at 633 nm (Supporting Information). This ratio then reverts approximately to unity at long times, as expected. Because the slowly aggregating sample does not consist entirely of monomers but certainly contains some, although a much lower number, of aggregates which would contribute disproportionately to the signal, the estimated 300-fold increase is a lower limit. This also provides an empirical gauge of the benefit one derives from the effort expended in controlling the formation of clusters. The brightness of the SERS nanocapsules was compared to the SERS signal resulting from randomly aggregated silver clusters using a rapid spectroscopic analysis of nanoparticles freely diffusing in a capillary.23 The apparatus used to carry out these measurements can simultaneously monitor with single particle sensitivity, the Raman, Rayleigh, and background components of light scattered following 633 nm illumination. The Raman signal is monitored over a small wavenumber range near the 1055 cm-1 (679 nm) in which intense Raman modes both of 4-mercaptobenzoic acid (MBA) and ABT occur (Figure 1d). The Raman/Rayleigh ratio calculated for each signal burst due to the entry of a one or more cluster into the detection volume of the confocal microscope, provides a measure of the SERS intensity from that particle. The solution is dilute, so that most events represent a single particle or cluster. The results shown in Figure 4 compare the behavior of Ag aggregates produced by adding 10 µM MBA to citrate Ag NPs (in red) and that obtained with nanocapsules synthesized using optimized ABT concentrations, polymer stabilized as in Figure 3a, and centrifuged to remove the majority of monomers and large aggregates (in green). Particle detection events located in the upper right-hand quadrant of Figure 4 represent the highest SERS activity compared to Rayleigh and background levels (i.e., optimal SERS substrates). The SERS signals recorded with the optimized nanocapsules are intense and uniform (ABT-linked particle synthetic reproducibility is demonstrated in Figures S3, S10, and S11). Using the single-particle statistical method23 (Figure 4) we find that nearly every particle in the optimized ABT sample is a highly SERS-active substrate, as the vast majority of events have a similarly high Raman/Rayleigh ratio (green). The Raman/background ratio indicates that the SERS from the taggant has high signal-to-noise. The histogram obtained from the distribution of clusters aggregated with 10 µM MBA shows that sample be a poor SERS system by comparison (red), reflecting the broad distribution of cluster sizes and the random occurrence of the adsorbate in the hot spots. In this nonoptimized sample the majority of events have both low Raman/Rayleigh

Braun et al. and Raman/background ratios. Notably, although higher concentrations of MBA produced larger aggregates, the Raman/ Rayleigh ratio was not increased over that of the ABT system. In all cases the nonuniformity in the recorded signal arises not only from the inherent distribution in SERS enhancement but also from the rotational (polarization) sensitivity of SERS and the random paths taken through the focal volume. Linking and Small Molecule Infusion. Adsorbates such as MBA are not as effective as bifunctional linkers for controllably aggregating Ag nanoparticles (Figure 4, Supporting Information). This is why salt is traditionally added as an aggregating agent when the adsorbate does not promote aggregation efficiently.1,10 The method described in Figure 1b allows SERS detection at low concentration of molecules that do not themselves promote sufficient aggregation. Sodium phosphate and HMD are used to aggregate the Ag nanoparticles, the clusters are diluted (as before) and coated using the polymer solution, forming a stable nanocapsule solution. Because the structure is encapsulated the remaining phosphate and HMD can then be displaced by the molecule of interest without loss of SERS efficacy, even months later (Supporting Information). During incubation with HMD the SERS intensity increases linearly with time (using the phosphate bands as references) until polymer quenching is carried out (Figure 5a). This process is highly reproducible. The polymer coating which stabilizes the cluster nanostructure is nevertheless sufficiently porous to allow small molecules such as MBA or ABT to enter the interparticle interstices without losing the cluster’s strong SERS enhancement or disrupting the stabilizing effect of the polymer shroud. Furthermore, adsorption of BSA through the polymer layer does not appear to interfere with the binding of small molecules to silver in the interparticle junction region, presumably because of the large globular nature of the protein or because of partial desorption. We should note that when only PVP is used to stabilize the nanocapsules, there is a limit to the quantity of thiol tags one can add before aggregation resumes once again. This is probably due to the tag inducing sufficient polymer desorption, owing to its greater affinity to silver, to undermine the quenching effect of the polymer. The addition of PEG-thiol or protein added after the PVP has quenched the initial linking step, greatly reduces this effect. Moreover, the SERS bands from the polymers are very weak, likely due to the fact that the polymer does not penetrate the junction effectively, as well as to its low Raman cross section. A number of Raman tag candidates were tested using the postinfusion approach. Most yielded similar SERS intensities (Figures 1c, 5a, and S1), suggesting that the geometry of the SERS clusters (and the hot spots they inhabit) are reproducible, stable, and not greatly affected by the addition of these species, most of which have similar Raman cross sections. It also implies that for the list of molecules shown, an approximately equal number of molecules enter the hot spot, although their infiltration rate through the polymer coating varies greatly. Also, despite the fact that the starting colloid is not monodispersed and the aggregates are made through random linkages, the resulting aggregate distributions are sufficiently constant to lead to highly reproducible SERS performance. Moreover, by striving to make the nanoparticle dimer, with its single hot spot, the dominant SERS-active species, one further reduces the influence of aggregate geometry and particle size on the SERS performance of the resulting substrates (so long as the latter is much smaller than the wavelength). With ABT used as Raman tag, added after BSA encapsulation, the SERS intensity of ABT was found to be only ∼30%

Generalized Approach to SERS-Active Nanomaterials

J. Phys. Chem. C, Vol. 113, No. 31, 2009 13627

Figure 5. (a) Time series over a typical synthesis of prelinked STV-coated clusters obtained by sequential additions of HMD, PVPA, and STV with BSA to Ag colloid. Portions of the stock are then combined with a number of Raman active molecules; for each molecule a resulting time segment is shown for spectroscopic comparison. ABT* is when ABT is added before the STV in a separate experiment, to show the effect on the 1380 and 1425 cm-1 bands of the sequence of addition. (b) Mixed-tagged STV clusters by sequential addition of BPDT and MBA, spectra normalized to the shared band at 1075 cm-1. An increasing amount of BPDT is added and allowed to reach SERS saturation, followed by the addition of a fixed MBA concentration to backfill remaining sites (0, 0.03, 0.06, 0.1, 1 relative units of BPDT), showing an isosbestic behavior (inset). (c) Kinetic trace of BPDT, APTT, and MBA adsorption onto protein coated SERS nanocapsules and fit to exponential or biexponential time-functions. MBA* indicates slightly faster adsorption through PVPA in the absence of protein.

lower than for nanocapsules prepared directly with ABT as the linker as per Figure 1a (see Figures 5a, ABT vs ABT*, and S8). This indicates that the junction is not significantly restricted to incoming molecules, justifying the prelinking approach. Interestingly, a large difference in the relative intensity of certain bands24 associated with chemical enhancement was found for samples that were exposed to BSA and NaCl before ABT (Figure S8). Such a difference resulting from order of addition could be used to better understand the fundamental aspects of surface chemical modification and molecular orientation on SERS enhancement. On close inspection, for ABT and other thiolate probes, the phosphate (∼230, 565, 924, and 1084 cm-1) and Ag-chloride (230 cm-1) Raman bands decreased upon tag addition to the SERS clusters, while the intensity of Raman modes associated with the incoming molecule simultaneously increased, an observation that is useful in determining the degree of exchange (Supporting Information). By using combinations of tags to backfill aliquots of prelinked clusters, a large number of unique barcodes for bioassays could be produced by controlling the ratio of tags. This was tested by synthesizing SERS clusters containing mixtures of benzene thiol moieties (BPDT and MBA) of varying ratios in their hot spots (Figure 5b), produced by adding a small quantity of BPDT to the polymer coated nanocapsules (as in Figure 1b), and allowing the SERS intensity to increase to saturation. An MBA solution was then added to fill the remaining available Ag surface sites. This process was repeated with an initial solution of varying concentration of BPDT. The resulting SERS spectra, which consist of superpositions of the SERS spectra of MBA and BPDT in varying proportions, reveal an isosbestic-like behavior (Figure 5b), suggesting that the total number of available sites in the hot spot is approximately constant (Supporting Information Figure S22) and that the Raman cross sections of the two labels are more or less equal. Moreover, the tags do not appear to interact on the surface, so that the spectra observed are linear superpositions of the SERS spectra of the singly tagged clusters. This will likely not be the case for all combinations of molecules.

In synthesizing the Ag SERS nanocapsules described above, the binding kinetics of a number of molecules were observed by following the time evolution of the SERS intensity after their addition. Most of them followed Langmuir-like single- or doubleexponential kinetics (Figure 5c). Kinetics of this sort can be used for fundamental studies on submonolayer molecular orientation and reorganization, mixing effects, functional group interactions, surface chemical reactions, competitive surface binding of various molecular species, and the transport of small or moderately sized molecules through the polymer25 and protein shell. Microscopy Applications. For multiplexing and cellular studies, such as delivery of nanoparticles with various cargos, nanocapsules can be used as barcoded Raman tags and simultaneously tracked in real time by wide-field fluorescence imaging. To first examine their multiplexing capacity, three Raman tagged nanocapsule solutions were mixed and deposited on a glass slide (Figure 6). The identification of each cluster and occupation from site to site was obtained through Raman mapping over the area of interest and deconvolution using LabSpec 6. The resultant deconvolution into ABT (red), MBA (green), and DTNB (blue) fundamental spectra in Figure 6 demonstrates that multiple clusters can be sensitively and independently detected at the single nanocapsule level. In Figure 7 we show Cy3-STV-TAT-coated nanocapsules imaged through Cy3 fluorescence microscopy bound to the surface of HeLa cells. We also tested antibody-functionalized ABT and MBA nanocapsules, which recognize two cell surface markers present on a human B-cell line (Figure 7b,c). Further development of a variety of bioactive SERS nanocapsules with such noninterfering fluorescence labeling should enable rapid determination of particle positioning during cell binding/uptake using high-speed fluorescence imaging, while particles of interest could be identified as needed by resolving the unique Raman spectral barcode. A wide variety of polymers, proteins, or nucleic acids may be chosen so as to bestow on the SERS nanocapsule the desired solubility, stability, fluorescence, and affinity functionalities, similar to other colloidal nanomaterials such as surface-protected

13628

J. Phys. Chem. C, Vol. 113, No. 31, 2009

Braun et al.

Figure 6. Deconvolution mapping. A mixture of ABT, MBA, and DTNB SERS nanocapsules dispersed on a glass slide and Raman mapped at 2 µm resolution. Fitting the overall SERS spectrum at each pixel to the standard SERS spectra of the individual components, the percent composition of the three component spectra was determined and displayed in a color-coded format. (a) Overlaid image with the composition at each position represented in RGB color space. (b, c) Two typical observed spectra and their decomposition into the three constituent nanocapsules with percent composition.

monomer nanoparticles and quantum dots. The SERS cross sections of the resulting SERS nanocapsules are intense enough for routine single-nanocapsule detection even in complex biological environments, and we are currently extending the controlled assembly technique to smaller nanoparticles. Taken together these results indicate the linking strategy is highly reproducible. By using a suitable quenching/stabilization step to separating the linking from the Raman tag adsorption one is able ensure a structurally robust yet chemically defined surface. We may now summarize the observed qualities of these encapsulated clusters. ABT-linked nanocapsule tunability is presented in Figures 2a, S10, and S11, single-nanocapsule (and hot spot) statistical uniformity in Figure 4, batch to batch

reproducibility in Figure S3, intensity advantage over monomers and effect of order of addition in Figures 2b, 5a (ABT vs ABT*), S2, and S10-S11. The infusion technique using HMD prelinking allows the facile screening of molecular adsorbates and numerous physical analyses on the fundamental aspects of SERS. This ultimately provides a route to generating multiplexing nanotags, some of which are shown in Figures 5a and S1, with demonstrated performance in Figures 6 and S16. Synthetic versatility allows spectral analysis or tracking of adsorbate kinetics during chemical modifications, presented in Figures 5b and c, S4-S8, S9, S12-S13, and S22. Finally, long-term stability is demonstrated in Figure S21, while Figures 7 and

Generalized Approach to SERS-Active Nanomaterials

J. Phys. Chem. C, Vol. 113, No. 31, 2009 13629 with chemical selectivity derived from the nature of the coating or by tags or receptors positioned near the hot spot. Acknowledgment. This work was supported by funding from the Institute for Collaborative Biotechnologies (ICB) through Grant No. DAAD19-03-D-0004 from U.S. Army Research Office, from Lawrence Livermore National Laboratories through a UCDRD grant. Funding from the UC Discovery program and Tamarisc Diagnostics to N.R. is acknowledged. Extensive use of the MRL Central Facilities at UCSB supported by the National Science Foundation under Awards No. DMR-0080034 and DMR-0216466 for electron microscopy. We thank Dr. Stanley M. Parsons (UCSB) for use of the Olympus epifluorescent microscope. Supporting Information Available: Additional experimental details, figures, spectra, and discussions of peripheral findings. This material is available free of charge via the Internet at http://pubs.acs.org. References and Notes

Figure 7. Cell binding using antibodies. (a) Fluorescence image of biotin-TAT peptide-functionalized Cy3-STV-labeled MBA nanocapsules bound to HeLa cancer cells. (b, c) B-cells (cell line 155) expressing both CD49e and CD19 antigens bound with ABT and MBA SERS clusters coated with antibody (Ab) Ab-CD49e and Ab-CD19, respectively. 10× objective SERS map overlay of deconvoluted ABT (red) and MBA (green) channels is shown along with optical image of the same region. Scale bars are 20 µm.

S18 exploit their protein functionalization possibilities and resilience toward biological buffers. Conclusions In summary, we show that by controlling the linker-mediated assembly, then using a polymer which subsequently acts as an encapsulant to quench the aggregation process when the SERS enhancement is optimized, SERS materials are produced with highly reproducible enhancements, long-term stability, and scalability to large-scale, batch synthesis. The adsorption of linkers, coatings, and tags onto the nanoparticles were monitored in real time, allowing the surface chemistry to be investigated at every stage of the process, including the infiltration of thiolate tags into the hot spot and their adsorption. The latter follows Langmuir-like kinetics. The sequential addition of phosphate, linker, polymer, streptavidin, Raman tag, and biotinylated affinity label could be carried out without purification through the judicious choice of binding strength, concentration, and molecular size. This strategy was used to demonstrate the multiplexed labeling of cancer cells, competitive with commonly used fluorescence multiplexing. Future efforts, besides fundamental studies on SERS enhancement, ligand adsorption, and generation of more unique and stable tags for multiplexing, include the engineering of uniformly responsive SERS substrates

(1) Michaels, A. M.; Jiang, J.; Louis Brus, L. Mol. J. Phys. Chem. B 2000, 104, 11965–11971. (2) Skadtchenko, B. O.; Aroca, R. Spectrochim. Acta 2001, A57, 1009– 1016. (3) Haynes, C. L.; Yonzon, C. R.; Zhang, X.; Van Duyne, R. P. J. Raman Spectrosc. 2005, 36, 471-484. (4) Kneipp, K.; Kneipp, H.; Kartha, V. B.; Manoharan, R.; Deinum, G.; Itzkan, I.; Dasari, R. R.; Feld, M. S. Phys. ReV. E 1998, 57, R6281– 6284. (5) Gearheart, L. A.; Ploehn, H. J.; Murphy, C. J. J. Phys. Chem. B 2001, 105, 12609–12615. (6) Ni, J.; Lipert, R. J.; Dawson, G. B.; Porter, M. D. Anal. Chem. 1999, 71, 4903–4908. (7) Cao, Y. C.; Jin, R.; Nam, J. M.; Thaxton, C. S.; Mirkin, C. A. J. Am. Chem. Soc. 2003, 125, 14676–14677. (8) Su, X.; Zhang, J.; Sun, L.; Koo, T.-W.; Chan, S.; Sundararajan, N.; Yamakawa, M.; Berlin, A. A. Nano Lett. 2005, 5, 49–54. (9) Qian, X.; Peng, X.-H.; Ansari, D. O.; Yin-Goen, G.; Chen, G. Z.; Shin, D. M.; Yang, L.; Young, A. N.; Wang, M. D.; Nie, S. Nat. Biotechnol. 2008, 26, 83–90. (10) Brown, L. O.; Doorn, S. K. Langmuir 2008, 24, 2277–2280. (11) Lee, S. J.; Guan, Z.; Xu, H.; Moskovits, M. J. Phys. Chem. C 2007, 111, 17985–17988. (12) Braun, G.; Pavel, I.; Morrill, A. R.; Seferos, D. S.; Bazan, G. C.; Reich, N. O.; Moskovits, M. J. Am. Chem. Soc. 2007, 129, 7760–7761. (13) Fabris, M.; Dante, M.; Nguyen, T.-Q.; Tok, J. B.-H.; Bazan, G. C. AdV. Funct. Mater. 2008, 18, 2518–2525. (14) Braun, G.; Lee, S. J.; Dante, M.; Nguyen, T.-Q.; Moskovits, M.; Reich, N. J. Am. Chem. Soc. 2007, 129, 6378–6379. (15) Lee, P. C.; Meisel, D. J. Phys. Chem. 1982, 86, 3391–3395. (16) Moskovits, M.; Vlcˇkova´, B. J. Phys. Chem. B 2005, 109, 14755– 14758. (17) Hu, X.; Wang, T.; Wang, L.; Dong, S. J. Phys. Chem. C 2007, 111, 6962–6969. (18) Murray, C. A.; Bodoff, S. Phys. ReV. B 1985, 32, 671–688. (19) Blue, D.; Helwig, K.; Moskovits, M. J. Phys. Chem. 1989, 93, 8080–8089. (20) Rosi, N. L.; Giljohann, D. A.; Thaxton, C. S.; Lytton-Jean, A. K. R.; Han, M. S.; Mirkin, C. A. Science 2006, 312, 1027–1030. (21) Mulvaney, S. P.; Musick, M. D.; Keating, C. D.; Natan, M. J. Langmuir 2003, 19, 4784–4790. (22) Yuan, J. J.; Zhu, P. X.; Fukazawa, N.; Jin, R. H. AdV. Funct. Mater. 2006, 16, 2205–2212. (23) Laurence, T. A.; Braun, G.; Talley, C.; Schwartzberg, A.; Moskovits, M.; Reich, N. O.; Huser, T. J. Am. Chem. Soc. 2009, 131, 162–169. (24) Osawa, M.; Matsuda, N.; Katsumasa, Y.; Uchida, I. J. Phys. Chem. 1994, 98, 12702–12707. (25) Alvarez-Puebla, R. A.; Contreras-Ca´ceres, R.; Pastoriza-Santos, I.; Pe´rez-Juste, J.; Liz-Marza´n, L. M. Angew. Chem., Int. Ed. 2009, 48, 138–143.

JP903399P