Controlled Permeability in Porous Polymer Nanocapsules Enabling

This is an open access article published under an ACS AuthorChoice License, which permits copying and redistribution of the article or any adaptations for non-commercial purposes.

Research Article www.acsami.org

Controlled Permeability in Porous Polymer Nanocapsules Enabling Size- and Charge-Selective SERS Nanoprobes Ying Jia,† Sergey N. Shmakov,‡ and Eugene Pinkhassik*,‡ †

Department of Chemistry, Saint Louis University, 3501 Laclede Ave., St. Louis, Missouri 63103, United States Department of Chemistry, University of Connecticut, 55 North Eagleville Road, Storrs, Connecticut 06269, United States

‡

S Supporting Information *

ABSTRACT: Nanoprobes for surface-enhanced Raman scattering (SERS) were prepared by creating nanorattles, or yolk−shell structures, containing gold or silver nanoparticles entrapped in porous hollow polymer nanocapsules. Controlled permeability of the shells of nanocapsules, achieved by controlling the pore size and/or shell surface functionalization, resulted in size- and chargeselective SERS analyses. For example, a trace amount of phenanthroline, a model analyte, was detected in human blood plasma without preprocessing of plasma samples. Comparison with commercially available nanoparticles showed superior performance of the newly prepared nanorattle structures.

KEYWORDS: size selectivity, charge selectivity, SERS, polymer nanocapsules, yolk−shell nanostructure

■

INTRODUCTION Surface-enhanced Raman scattering (SERS) has demonstrated great potential as an important tool for fundamental and applied studies with advantages of extremely low detection limit, high sensitivity, surface selectivity, and nondestructive analysis.1,2 Progress in the analysis of complex mixtures, such as biological fluids, is constrained by the presence of the multitude of interfering molecules. Physiological fluids, like blood or urine, contain proteins, various amino acids, metabolites, and other components that make the analysis especially complicated.1 The most common SERS substrates, such as citratestabilized gold or silver nanoparticles, are particularly vulnerable to protein binding that leads to the formation of the so-called “protein corona” on their surface.3−6 As a result, nanoparticles are effectively prevented from aggregating and forming hot spotsan essential prerequisite for the strong signal enhancement by nanoparticles.7 In addition, the “protein corona” hinders the access of analytes to the surface of nanoparticles. An important aspect in the analysis of biofluids is focusing on increasing SERS selectivity and specificity toward desired analytes. The most notable approaches include covering of the metal nanoparticle surface with analyte-selective labels2,7 or molecularly imprinted polymers.8−12 Other methods use selective coatings and often require preseparation of components in complex physiological or chemical mixtures before the SERS analysis.13−17 Materials with selective nanopores offer an attractive alternative to coatings by providing chemical or physical filtering of interfering molecules.18−20 This approach reduces the need for mixture pretreatment and improves selectivity. © XXXX American Chemical Society

Functionalization of sieving materials can further reduce the need for the pretreatment of complex mixtures.21 Yolk−shell structures that entrap clusters of nanoparticles in a semipermeable shell appear to be a particularly attractive, although currently underutilized, design for the creation of selective SERS nanoprobes. Entrapment of clusters of nanoparticles within a hollow capsule offers signal enhancement through interparticle coupling. Regulation of the permeability of the shell may control selectivity of the analysis. Encapsulation may be compatible with methods providing selectivity through surface coatings, thus leading to highly selective devices capable of multistage in situ sample processing. The main goal of this study is to test the hypothesis that selective permeability through the shells of nanocapsules will enable selective SERS measurements using encapsulated nanoparticles (Figure 1A). Here, we investigate SERS nanoprobes based on gold or silver nanoparticles entrapped in porous polymer nanocapsules with nanometer-thin shells. We focus on exploring size and charge selectivity due to tunable permeability across the shell boundary. Polymer nanocapsules used here were prepared by a straightforward vesicle-templating method.22,23 In this approach, monomers and cross-linkers are loaded into the interior of a bilayer formed by lipids or catanionic surfactants and polymerized to form a hollow polymer shell with a single-nanometer thickness. Unlike other yolk−shell or nanorattle structures, the shells of these capsules Received: May 9, 2016 Accepted: May 17, 2016

A

DOI: 10.1021/acsami.6b05522 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX

Research Article

ACS Applied Materials & Interfaces

metal nanoparticles of controlled size ranging from a single nanometer to tens of nanometers entrapped within hollow polymer capsules with typical diameters in the 100−200 nm range (Figure 1).33 We showed that encapsulation of metal nanoparticles in hollow nanocontainers with porous polymer walls resulted in size selectivity and improved catalyst recycling without compromising reaction kinetics.34 In this work, we investigated the performance of Au and Ag nanoparticles entrapped in polymer nanocapsules (AuNCs and AgNCs, respectively) as size- and charge-selective SERS nanoprobes.

■

EXPERIMENTAL SECTION

Materials and Methods. Chemicals were purchased from SigmaAldrich unless noted otherwise. 1,2-Dimyristoyl-sn-glycero-3-phosphocholine (DMPC) was purchased from Avanti Polar Lipids, Inc. as a dry powder. All chemicals were used as received, unless noted otherwise. Butyl methacrylate (BMA), tert-butyl methacrylate (t-BMA), and ethylene glycol dimethacrylate (EGDMA) were passed through a freshly activated alumina column to remove the inhibitor shortly before the polymerization. Solvents (Fisher Scientific) were HPLCgrade and used as received. Glass and quartz slides were purchased from Fisher Scientific and Quartz Scientific, Inc. respectively. Amicon Ultra-0.5 mL centrifugal filters (cutoff 3 kDa) were purchased from EMD Millipore. All glassware was washed with aqua regia prior to synthesis of gold nanoparticles. Hydrodynamic diameter measurements and zeta potential measurements were performed on a Malvern Nano-ZS Zetasizer (Malvern Instruments Ltd., Worcestershire, U.K.). Scanning electron microscopy (SEM) and scanning transmission electron microscopy (STEM) images were acquired on an FEI Inspect F50 microscope. Transmission electron microscopy (TEM) was carried out on an FEI Tecnai T12. Cryo-TEM micrographs were recorded on an FEI Tecnai Arctica. The SERS spectra were recorded with a Renishaw inVia (Renishaw plc, U.K.) equipped with a Leica DM2500 M microscope with a 50× objective (N.A. = 0.75) and CCD camera. A 785 nm diode laser (300 mW) was used for experiments with blood plasma. A He−Ne 633 nm laser (17.5 mW) was used for the rest of the measurements. The gratings used were 1800 lines/mm for 633 nm and 1200 lines/mm for 785 nm lasers. Calibration was performed using a reference 520 cm−1 vibrational band of an internal silicon standard prior to every experiment. UV/vis measurements were performed on an OLIS absorbance spectrophotometer equipped with CLARiTY integrating cavities. Synthetic Procedures. Synthesis of Nanocapsules. Nanocapsules were prepared using previously described procedures.22,25,26,32 Aqueous stock solutions containing monomers and cross-linkers mixed with (S1) cetyltrimethylammonium tosylate (CTAT) and (S2) sodium dodecylbenzenesulfonate (SDBS) were prepared as follows. Solution S1: t-BMA (64 μL, 0.4 mmol), BMA (64 μL, 0.4 mmol), EGDMA (64 μL, 0.34 mmol), 2,2-dimethoxy-2phenyl-acetophenone, DMPA, (1 mg, 4 μmol), and CTAT (200 mg, 0.438 mmol) were added to a test tube, followed by addition of 20 mL of 1 mM aqueous tannic acid solution. Solution S2: t-BMA (64 μL, 0.4 mmol), BMA (64 μL, 0.4 mmol), EGDMA (64 μL, 0.34 mmol), DMPA, (3 mg, 0.01 mmol), and SDBS (200 mg, 0.57 mmol) were added to a test tube, followed by addition of 20 mL of 1 mM aqueous tannic acid solution. Solutions were kept at 35−40 °C for 15 min, then shaken or briefly sonicated to give a homogeneous dispersion. Then, solutions were quickly mixed in desired proportions. In a typical experiment, solutions were mixed in the 2:8 S1:S2 ratio, i.e., 2 mL of S1 and 8 mL of S2. The mixture was agitated on a vortexer several times and then incubated undisturbed for 30 min at 35 °C. The resulting suspension containing vesicles of different sizes was extruded 4−5 times at 35 °C through a track-etched polyester membrane (Sterlytech) with a 0.2 μm pore size using a Lipex stainless steel extruder (Northern Lipids). The extruded sample was irradiated (λ = 254 nm) in a photochemical reactor equipped with a stirrer (10 lamps

Figure 1. (A) Schematic representation of size-restricted permeation of analytes through the ultrathin porous shells of hollow polymer nanocapsules containing entrapped gold nanoparticles (AuNCs): analytes smaller than the pore size (shown on the right) can enter the nanocapsule and interact with gold nanoparticles, whereas analytes larger than the pore size (shown on the left) are not able to enter the nanocapsule and have no contact with gold nanoparticles. Cryo-TEM micrographs of empty liposomes (B) and monomer-loaded liposomes before the polymerization of monomers in the hydrophobic interior of the lipid bilayer (C). STEM (D) and SEM (E) of typical AuNC used in this study showing individual gold nanoparticles entrapped within a hollow polymer nanocapsule.

can be easily functionalized by imprinting nanopores with different sizes or chemical environment.24−26 In addition, the bilayer-templating approach permits decoration of the shell surface with different functional groups.27 Permeability of the shells of these nanocapsules can be controlled by controlling the size and chemical environment of imprinted nanopores.28 Molecules and ions smaller than the pore size exhibited ultrafast transport through the pores, as expected for semipermeable membranes with the thickness in the single-nanometer range.29−31 In contrast, molecules larger than the pore size are not able to cross the shell as found in the same studies. Recently, we used vesicle-templated nanocapsules for the synthesis of polymer yolk−shell structures with metal cores.32,33 These structures can contain single or multiple B


Research Article


with trimethylamine in diethyl ether at 0 °C, as described by Michas et al. (yield, 72%).35,36 1H NMR (D2O): δ 6.12 (s, 1H), 5.71 (s, 1H), 4.21 (t, 2H), 3.31 (m, 2H), 3.1 (s, 9H), 1.91 (s, 3H), 1.85−1.55 (m, partially overlapped, 6H) 1.35 (m, 12H). LC/MS (ESI) m/z: calcd [M + Na]+ = 401 and [M − Br]+ = 298, found 401 and 298. 11-Bromoundecyl Methacrylate.35,37 11-Bromoundecanol (10 g, 40 mmol) and 150 mL of anhydrous THF were mixed in a flask at 0 °C under N2, followed by the addition of 6 mL (61 mmol) of methacryloyl chloride. The mixture was purged with N2 at room temperature for 2 h and stirred for 12 h. The unreacted methacryloyl chloride and THF were removed under reduced pressure. The residue was dissolved in ethyl ether and washed with saturated sodium hydrogen carbonate solution until the aqueous layer became basic. After evaporation of ethyl ether, a viscous yellowish liquid of 11bromoundecyl methacrylate was obtained with a yield of 89%. GC/MS m/z: [M]+ = 318. SERS Analysis. Analytical experiments were carried out on glass or quartz slides. Size-Selective SERS. A 0.1 mL portion of 1 μM 4-nitrothiophenol (4-NTP) was mixed with SERS substrates of 0.1 mL of the Au nanoparticles (Au NPs) dispersion in PBS (∼7.2 × 1010 NPs/mL) or 0.1 mL of the AuNCs solution prepared as described above, respectively. Rhodamine 6G (0.1 mL, 20 μM), R6G, was mixed with the correspondent SERS substrate as mentioned above. The same mixing procedure was repeated for the mixture of 4-NTP and R6G with two substrates. The sample (10 μL) was then dropped on the glass slide after a 15 min equilibration. Droplets were rapidly dried under the stream of N2 gas and analyzed. A laser with 633 nm excitation, 3 s exposure time, 0.0875 mW of power, under a 50× objective was used. Reconstituted filtered plasma was mixed with SERS substrates (Ag NPs in citrate buffer or AgNCs prepared as described above) in a ratio of 1:9 (plasma:Ag substrates). The Ag NPs concentration was 0.02 mg/mL. Filtered and unfiltered plasma was spiked with 1,10phenanthroline. This mixture was added to SERS substrates (Ag NPs or AgNCs) in a ratio of 1:9. The sample (10 μL) was then dropped on the quartz slide within 1−2 minutes after mixing. Droplets were rapidly dried under the stream of N2 and analyzed. A laser with 785 nm excitation, 3 s exposure time, 1.5 mW of power, under a 50× objective was used. Charge-Selective SERS. Solutions of 8 μM 4-aminothiophenol (4ATP) were prepared in Britton-Robinson buffers at pH 3.3 and pH 7. 0.1 mL; each stock solution was added to 0.1 mL of SERS substrate dispersions (Au NPs in citrate buffer or AuNCs prepared as described above). The Au NPs concentration was ∼7.2 × 1010 NPs/mL. The sample (10 μL) was then dropped on the glass slide after a 15 min equilibration. Droplets were rapidly dried under the stream of N2 and analyzed using a laser with 633 nm excitation, 3 s exposure time, 0.875 mW of power, and a 50× objective. Data Analysis. All spectroscopic data were acquired and processed using the software WiRE 4.1 (Renishaw). To produce mapping plots, spectra were baseline corrected and selected peaks were fitted to determine their intensity. To build mapping plots, peaks at 1078 cm−1 for 4-ATP, 1330 cm−1 for 4-NTP, and 1363 cm−1 for R6G were selected.

of 32 W each; 10 cm distance between the lamps and the sample) for 60 min. Methanol (10 mL) was added, and the precipitate was washed 3−5 times with methanol and then 3 times with DI water over a period of 24 h. Surface-Charged Nanocapsules. Surface-charged nanocapsules were prepared similarly to the procedure used to prepare noncharged nanocapsules. An extra step was introduced: the extrusion was followed by a dropwise addition of a 1.5 wt % aqueous solution of methacryloyloxyundecyl trimethylammonium bromide (MUTB) to the suspension of vesicles within 4−5 min at constant gentle vortex mixing. The amount of MUTB added corresponded to 10 mol % of the total combined surfactant content (CTAT + SDBS). Polymer Nanocapsules with Entrapped Gold Nanoparticles (AuNCs). A slurry of precipitated nanocapsules with encapsulated tannic acid prepared as described above (ca. 5 mg of dry material) was transferred to a screw-capped test tube equipped with a stir bar and dispersed in 2 mL of Milli-Q water. Then, 20 μL of a 10 mM aqueous HAuCl4 solution was added to the nanocapsules dispersion, and the mixture was agitated for 30 min at ambient temperature. Then, 200 μL of 10 mM HAuCl4 was added. After 1 h, 8 mL of water was added in the mixture. The resulting colored precipitate was washed with water. In these washing steps, nanocapsules were precipitated using a centrifuge (3 min at approximately 2000g), the supernatant was decanted, and the process was repeated five times. The final precipitate was reconstituted with water to 2 mL of the suspension. All batches of AuNCs were combined to form a stock solution used for the SERS measurements. Polymer Nanocapsules with Entrapped Silver Nanoparticles (AgNCs). Polymer nanocapsules with entrapped silver nanoparticles were prepared by a liposome-templated method developed earlier.32 In a typical experiment, t-BMA (43 μL, 0.193 mmol), BMA (42 μL, 0.199 mmol), EGDMA (32 μL, 0.17 mmol), and DMPA, (1 mg, 3.9 μmol) were added to a 0.4 mL solution of DMPC (160 mg, 0.236 mmol) in CHCl3. The CHCl3 was evaporated using a stream of argon to form a lipid/monomer mixture on the wall of a test tube. The film was further dried under vacuum for 5 min to remove traces of CHCl3. The dried film was hydrated with 8 mL of a 0.15 M aqueous solution of AgNO3 to give a dispersion of multilamellar vesicles. During the hydration of the lipid/monomer mixture, the test tube was briefly agitated on a vortexer every 5 min. The suspension was extruded 16 times at 35 °C through a track-etched polyester Nucleopore membrane (Sterlytech) with 0.2 μm pores using a Lipex stainless steel extruder (Northern Lipids). Prior to polymerization, nonentrapped silver ions were removed from the mixture by size-exclusion chromatography on a Sephadex G-50 column. Oxygen was removed by passing argon through the solution. Polymerization was initiated with UV irradiation as described above. Methanol (10 mL) was added, and the precipitate was washed 3−5 times with methanol and then 3 times with DI water. In these washing steps, nanocapsules were precipitated using a centrifuge (3 min at approximately 2000g), the supernatant was decanted, and the process was repeated. The final precipitate was reconstituted with water to 8 mL of the suspension. All batches of AgNCs were combined to form a stock solution used for the SERS measurements. Size of Au and Ag Nanoparticles. The size of Au and Ag nanoparticles was determined from TEM and STEM micrographs. Size distributions were calculated by measuring 200−300 nanoparticles from a collection of several micrographs. In these measurements, several micrographs were taken from random areas of the sample. Each nanoparticle within each of these micrographs was measured, and the results were plotted on a histogram. Processing and Storage of Reconstituted Blood Plasma. Plasma from human blood (Sigma-Aldrich) was reconstituted to the indicated volume with pH 7 PBS buffer. Aliquots of plasma were stored (unfiltered samples), while the rest of aliquots were centrifuged (Amicon Ultra Centrifugal filters, 3 kDa cutoff) at 14 000g, room temperature, for 20 min. All samples were stored at −80 °C prior to analysis. Methacryloyloxyundecyl Trimethylammonium Bromide (MUTB). MUTB was synthesized by reacting 11-bromoundecyl methacrylate

■

RESULTS AND DISCUSSION The selectivity in the yolk−shell nanoprobes investigated here is achieved by the permeability through the pores of nanocapsules and should be independent of the nature, size, or size distribution of the encapsulated SERS substrates. We used the most common SERS substrates, gold and silver nanoparticles, to evaluate their utility in the construction of yolk−shell selective nanoporbes. Our experiments, described in detail below, focused on the interactions between the analytes and the shells of nanocapsules in achieving selective SERS measurements. C


Research Article

ACS Applied Materials & Interfaces By choosing reaction conditions (temperature, time, reducing agents), we can tune the size of metal nanoparticles in yolk−shell nanostructures, as reported previously.33 In this work, we used tannic acid (TA) as a sacrificial molecule to initiate and grow gold nanoparticles exclusively inside polymer nanocapsules. The concept of site-specific synthesis recently gained popularity because it affords a clean final product and eliminates separation steps.38 The average size of Au nanoparticles synthesized by this approach was 23 ± 12 nm (Figures S1 and S2). The size distribution of nanoparticles should have no effect for this study since the selectivity is achieved by the permeability of analytes through the shells of nanocapsules, and the spectra are analyzed for patterns rather than absolute intensity of the signals. Commercially available Au nanoparticles in citrate or PBS buffer solution were used as a reference. We chose reference nanoparticles 40 nm in diameter to ensure a strong and reliable SERS signal (Figure S3). To test SERS activity of AuNCs, we chose 4-nitrothiophenol (4-NTP) and Rhodamine 6G (R6G) that are popular model analytes in SERS analysis.20,39,40 The smallest dimension of 4-NTP is approximately 0.6 nm. R6G has the smallest dimension of approximately 1 nm. In our previous work, we showed that ultrathin shells of Au nanoreactors did not hinder the diffusion of 4-nitrophenol that has the same smallest dimension as 4-NTP while effectively preventing the diffusion of 3,5-di-tert-butyl benzaldehyde, whose smallest dimension is similar to that of R6G.33 All SERS measurements reported here were performed in the dry state on drop-casted samples. All samples were allowed to equilibrate for at least 15 min before drop-casting, unless noted otherwise, to ensure that all the molecules that could diffuse through the shells entered the nanocapsules and adsorbed on the surface of nanoparticles. We found previously that molecules and ions smaller than the pore diffuse through the nanopores at a very high rate. For example, investigation of entrapped pH-sensitive dyes revealed that the full equilibration occurred on the millisecond time scale or faster with mixing of two solutions being the rate-limiting step.29 Previous studies showed no evidence of nanorattles being ruptured upon drying.25 In preliminary experiments, we saw no difference in the outcome of SERS measurements when using equilibration times ranging from 5 min to 2 h. This experimental setup alleviates any differences that might have been caused by specific adsorption characteristics of molecules in the mixture. Because the selectivity is determined by the permeability through the shells and because time-dependent experiments confirmed rapid equilibration, the only important parameter in these measurements is the smallest dimension of analyte molecules, or their ability to cross the shells of the nanocapsules. To avoid the influence of intrinsic nonhomogeneity of drop-casted samples on the measurements, multiple spectra were acquired in different locations and representative spectra from different samples were normalized to offer meaningful comparison of patterns of signals in the spectra. SERS spectra of R6G and 4-NTP collected on nonentrapped Au nanoparticles are shown in Figure 2, spectra A and B, respectively. In experiments with individual analytes, we found that AuNCs only enhanced the signal from 4-NTP and not from R6G (Figure S4), suggesting that R6G was too big to enter the nanocapsules, in agreement with expected size selectivity offered by limited permeability through the capsule shells. To test the size selectivity further, we measured SERS

Figure 2. Representative SERS spectra of 10 μM R6G (A), 0.5 μM 4NTP (B), and a mixture of R6G and 4-NTP (C, D). Spectra were collected on Au nanoparticles (A, B, C) and AuNCs (D). Spectra were normalized with respect to the tallest peak.

signals of the 4-NTP/R6G mixture. Nonentrapped Au nanoparticles did not discriminate molecules by size, predictably enhancing signals of both components in the mixture (Figure 2, spectrum C). Because of the inherently stronger response to 4-NTP, we used a 20:1 ratio of R6G to 4NTP in the mixture (10 μM R6G vs 0.5 μM 4-NTP) to observe equally strong signals from both compounds in control measurements (Figure 2, spectrum C). The resulting complex SERS spectrum featured peaks from both molecules (Figure 2, spectrum C). In contrast, AuNCs only showed 4-NTP in the same mixture (Figure 2, spectrum D), confirming size-selective SERS measurements due to selective access of smaller 4-NTP to the nanocapsules and the ability to keep out the larger R6G. Spectra obtained with AuNCs did not reveal discernible signals from the polyacrylate shells of nanocapsules. The lack of interference from the shell is in agreement with our expectation that the main mechanism for signal enhancement is due to interparticle coupling caused by the aggregation of nanoparticles in the nanocapsules. In this work, more than 50% of nanocapsules contained between 3 and 10 nanoparticles with the majority of nanoparticles being smaller than 20 nm. Since particle/shell interactions are virtually unavoidable in yolk− shell structures, not seeing the signals from the shell is useful for practical applications. Because the selectivity is achieved by the controlled permeability through the shells, size distribution of encapsulated nanoparticles does not affect the pattern of observed signals. To demonstrate the differences in size selectivity of Au nanoparticles and AuNCs on a larger scale, we mapped the sample of the R6G/4-NTP two-component mixture (Figure 3). Au nanoparticles showed both 4-NTP and R6G (Figure 3A,B; Figure S5A), while AuNCs only revealed 4-NTP (Figure 3C,D; Figure S5B). These findings showed successful size-selective SERS analysis due to controlled permeability of nanocapsules. We then tested size-selective performance of polymer nanocapsules using human blood plasma spiked with a model molecule as a complex mixture. In the past, silver nanoparticles were shown to be a substrate of choice for SERS analysis of blood plasma.8,17,41 Similarly to these recent studies, we used a 785 nm laser for measurements performed in plasma. D


Research Article


blood plasma spiked with Phen (50 nM) for the reference nanoparticles and AgNCs to test the degree of biofouling interference. In our study, apparent biofouling completely suppressed the signal from Phen on the reference nanoparticles in a control experiment performed without nanocapsules (Figure 4, red). In contrast, SERS data collected for AgNCs under the same experimental conditions revealed the presence of Phen analyte (Figure 4, green). The experimental data obtained for conventional substrates, Ag nanoparticles (Figure 4, red curve), are consistent with previous reports highlighting the difficulties in performing SERS analysis of blood plasma without preprocessing.44 A common preprocessing method for the analysis of blood plasma has been ultrafiltration using membranes with a 3 kDa molecular weight cutoff that removes most proteins and other large biomolecules.44 When the filtered plasma was mixed with nonencapsulated Ag nanoparticles in a control experiment, signals from various components of blood plasma appeared (Figure 5, black), which Figure 3. SERS signal intensity (counts) of 0.5 μM 4-NTP (blue) and 10 μM R6G (red) on the map of two-component mixture. Maps show the same region for Au nanoparticles (A, B) and AuNCs (C, D), respectively. Pixel size is 1 μm2.

We synthesized silver nanoparticles inside nanocapsules by the liposome-templated method.32 This approach yielded nanoparticles with the average diameter of 8.2 nm (Figures S6 and S7). Citrate-stabilized Ag nanoparticles with an average diameter of 40 nm were used as a reference (Figure S8). In our experiments, we used 1,10-phenanthroline (Phen), which is a planar molecule with the smallest dimension of ca. 0.7 nm. It has been a popular model SERS analyte for silverbased substrates with a strong signal due to its large Raman cross section (Figure 4).42,43 Furthermore, its SERS signals overlap with signals from blood plasma, making it especially suitable for evaluating selective analysis methods. Previous studies of human blood plasma reported biofouling of silver nanoparticles by plasma proteins.14,44 Proteins readily adsorb on the surface of nanoparticles forming a “protein corona”, which hinders aggregation of Ag nanoparticles and results in weak SERS signals.3−6 We recorded SERS spectra of

Figure 5. SERS of filtered blood plasma on nonencapsulated Ag nanoparticles (black); SERS of filtered blood plasma spiked with 1,10phenanthroline on nonencapsulated Ag nanoparticles (red) and AgNCs (green). Blood plasma was filtered through a membrane with a 3 kDa cutoff. Spectra normalized with respect to the tallest peak. C (Phen) = 50 nM.

Figure 4. SERS spectrum of phenanthroline on Ag nanoparticles (black); SERS spectrum of blood plasma spiked with 1,10phenanthroline on Ag nanoparticles at 2× magnified intensity (red) and AgNCs (green). C (Phen) = 50 nM.

is fully consistent with the previously reported results.15,41,44 The SERS spectrum of filtered plasma spiked with Phen on nonencapsulated Ag nanoparticles appeared to be essentially the same (Figure 5, red line), showing no characteristic signals of Phen. Competitive adsorption of smaller molecules on the surface of Ag nanoparticles and peaks overlapping are likely to be the main reasons for the absence of a Phen signal in the spectrum. In contrast, when AgNCs were used as substrates, the Phen signal dominated the SERS spectrum of filtered plasma (Figure 5, green). We conclude that the ultrathin porous walls of nanocapsules limit competitive adsorption of interfering molecules from plasma and maintain high activity of entrapped Ag nanoparticles. These observations demonstrate the feasibility of size-selective SERS analysis of physiological fluids with minimal sample processing. The permeability of nanocapsules can be also controlled through electrostatic interactions of charged molecules.24,45,46 Recent studies showed that the charge on the surface of substrates allowed regulating the adsorption of charged molecules in SERS analysis.13,39,47−51 E


Research Article

ACS Applied Materials & Interfaces To expand the utility of nanocapsules-based SERS nanoprobes, we investigated the charge-selective permeability in SERS analysis. We synthesized nanocapsules with a positively charged surface by adding a polymerizable cationic surfactant methacryloyloxyundecyl trimethylammonium bromide (MUTB) to the suspension of vesicles. We expected MUTB to incorporate into an external leaflet of monomer-loaded bilayers of vesicles (Figure 6A,B) and copolymerize with the monomers and cross-linkers in the bilayer to become an integral part of the nanocapsule (Figure 6C). These newly prepared charged nanocapsules formed dispersions in water that remained stable for at least several months. We then synthesized positively charged AuNCs using encapsulated TA as sacrificial molecule (Figure S9). Zeta

potential measurements confirmed a positive charge of MUTBmodified AuNCs at pH 3.3 (Figure 6D). In order to test charge selectivity of Au substrates, we mixed them with 4-aminothiophenol (4-ATP) under acidic and neutral conditions. The pKa of protonated 4-ATP was reported to be 4.3.52 At pH 3.3, the amino group of 4-ATP is protonated and positively charged while it is neutral at pH 7. Results of SERS sample mapping at neutral pH are very similar for reference nanoparticles, nonmodified AuNCs, and positively charged AuNCs (Figure 7A−C), respectively. In contrast, at pH 3.3, only reference nanoparticles and neutral AuNCs strongly enhance the signal of 4-ATP (Figure 7D,E), whereas positively charged AuNCs show almost no SERS signal (Figure 7F). The drastic difference in signal enhancement by charged AuNCs under neutral and acidic conditions (Figure 7C,F, respectively) could be attributed to the very low concentration of 4-ATP in the interior of charged AuNCs. These data suggest that the electrostatic repulsion of positively charged protonated 4-ATP from the positively charged surface of nanocapsules prevented the diffusion of 4-ATP molecules into the core of AuNCs. The combination of charge selectivity with the size selectivity described above opens the opportunity to increase substrate selectivity or design other functional nanodevices for SERS analysis.48,53,54

■

CONCLUSIONS Yolk−shell structures, or nanorattles, based on porous polymer nanocapsules with encapsulated Au or Ag nanoparticles acted as size- and charge-selective SERS nanoprobes. The selectivity was achieved by controlled permeability through the nanometer-thin shells of nanocapsules. Molecules that are smaller than the pore size or lacking electrostatic repulsion from the shells were able to enter the nanocapsules and exhibit Raman signals. Conversely, molecules that were not able to enter the nanocapsules due to their size or charge were not detected. Selective permeability was based entirely on size or charge of analytes and did not involve any specific interactions with the shells of nanocapsules. Furthermore, polymer nanocapsules alleviated biofouling of the surface of substrates in the analysis of physiological fluids, eliminating otherwise necessary complex pretreatment steps. These findings establish yolk−shell nanostructures described here as a viable platform technology for the targeted creation of selective SERS nanoprobes. As shown previously, the permeability of vesicle-templated polymer nanocapsules can be further controlled by imprinting of pores with different sizes or chemical environment and likely by subsequent functionalization of pores and surface of shells; therefore, we expect that this method can be readily adaptable to a broad range of analytes and complex mixtures.24 An important attribute of the nanorattle or yolk−shell nanoprobes described here is that they are fully compatible with existing methods aiming at achieving selective SERS analysis by selective coatings of metal nanoparticles.21,55 One can envision future devices where the shell of the nanocapsule would provide initial separation of analytes in a mixture based on their size and charge with additional selectivity achieved by a permselective layer on encapsulated metal nanoparticles. Because of the versatility of this approach and potential for tuning the functionalization of the nanocapsules, we anticipate the ability to achieve highly selective SERS analysis in complex mixtures with minimal or no preprocessing of samples.

Figure 6. Scheme of synthesis of surface-charged polymer nanocapsules and corresponding Cryo-TEM micrographs of monomerloaded surfactant vesicles before (A) and after addition of MUTB (B). Surface-charged polymer nanocapsule after polymerization (C) and zeta potential of surface-charged AuNCs (D) along with a structure of cationic surfactant methacryloyloxyundecyl trimethylammonium bromide (MUTB). F


Research Article


Figure 7. Nonselective and size-selective transport of 4-ATP to substrates at different pH and correspondent signal intensity (counts) of 4-ATP on the SERS maps at pH = 7 (A, B, C) and pH = 3.3 (D, E, F). Substrates are Au nanoparticles (A, D), not charged AuNCs (B, E), and surface (+) charged AuNCs (C, F). Pixel size is 1 μm2.

■

■

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsami.6b05522.

■

REFERENCES

(1) Alvarez-Puebla, R. A.; Liz-Marzán, L. M. SERS-Based Diagnosis and Biodetection. Small 2010, 6, 604−610. (2) Wang, Y.; Yan, B.; Chen, L. SERS Tags: Novel Optical Nanoprobes for Bioanalysis. Chem. Rev. 2013, 113, 1391−1428. (3) Zhang, D.; Ansar, S. M.; Vangala, K.; Jiang, D. Protein Adsorption Drastically Reduces Surface-Enhanced Raman Signal of Dye Molecules. J. Raman Spectrosc. 2010, 41, 952−957. (4) Bonifacio, A.; Cervo, S.; Sergo, V. Label-Free Surface-Enhanced Raman Spectroscopy of Biofluids: Fundamental Aspects and Diagnostic Applications. Anal. Bioanal. Chem. 2015, 407, 8265−8277. (5) Walkey, C. D.; Chan, W. C. W. Understanding and Controlling the Interaction of Nanomaterials with Proteins in a Physiological Environment. Chem. Soc. Rev. 2012, 41, 2780−2799. (6) Gebauer, J. S.; Malissek, M.; Simon, S.; Knauer, S. K.; Maskos, M.; Stauber, R. H.; Peukert, W.; Treuel, L. Impact of the Nanoparticle−Protein Corona on Colloidal Stability and Protein Structure. Langmuir 2012, 28, 9673−9679. (7) Shiohara, A.; Wang, Y.; Liz-Marzán, L. M. Recent Approaches toward Creation of Hot Spots for SERS Detection. J. Photochem. Photobiol., C 2014, 21, 2−25. (8) Chen, S.; Li, X.; Guo, Y.; Qi, J. A Ag-Molecularly Imprinted Polymer Composite for Efficient Surface-Enhanced Raman Scattering Activities under a Low-Energy Laser. Analyst 2015, 140, 3239−3243. (9) Guo, Y.; Kang, L.; Chen, S.; Li, X. High Performance SurfaceEnhanced Raman Scattering from Molecular Imprinting Polymer Capsulated Silver Spheres. Phys. Chem. Chem. Phys. 2015, 17, 21343− 21347. (10) Xue, J.-Q.; Li, D.-W.; Qu, L.-L.; Long, Y.-T. Surface-Imprinted Core−Shell Au Nanoparticles for Selective Detection of Bisphenol a Based on Surface-Enhanced Raman Scattering. Anal. Chim. Acta 2013, 777, 57−62. (11) Ahmad, R.; Griffete, N.; Lamouri, A.; Felidj, N.; Chehimi, M. M.; Mangeney, C. Nanocomposites of Gold Nanoparticles@

SERS spectra and distribution of nanoparticles by the size (PDF)

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Author Contributions

The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Funding

This work was supported by the National Science Foundation (CHE-1522525 and DMR-1338021). Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS We thank Dr. Jessica Rouge for help with the zeta-potential measurements. Cryo-TEM studies were performed at the Center for Nanoscale Systems (CNS) at Harvard University, a member of the National Nanotechnology Coordinated Infrastructure Network (NNCI), which is supported by the National Science Foundation under NSF award no. 1541959. G


Research Article

ACS Applied Materials & Interfaces Molecularly Imprinted Polymers: Chemistry, Processing, and Applications in Sensors. Chem. Mater. 2015, 27, 5464−5478. (12) Kamra, T.; Zhou, T.; Montelius, L.; Schnadt, J.; Ye, L. Implementation of Molecularly Imprinted Polymer Beads for Surface Enhanced Raman Detection. Anal. Chem. 2015, 87, 5056−5061. (13) Marsich, L.; Bonifacio, A.; Mandal, S.; Krol, S.; Beleites, C.; Sergo, V. Poly-L-Lysine-Coated Silver Nanoparticles as Positively Charged Substrates for Surface-Enhanced Raman Scattering. Langmuir 2012, 28, 13166−13171. (14) Vicario, A.; Sergo, V.; Toffoli, G.; Bonifacio, A. SurfaceEnhanced Raman Spectroscopy of the Anti-Cancer Drug Irinotecan in Presence of Human Serum Albumin. Colloids Surf., B 2015, 127, 41− 46. (15) Hsu, P.-H.; Chiang, H. K. Surface-Enhanced Raman Spectroscopy for Quantitative Measurement of Lactic Acid at Physiological Concentration in Human Serum. J. Raman Spectrosc. 2010, 41, 1610− 1614. (16) Han, Z.; Liu, H.; Wang, B.; Weng, S.; Yang, L.; Liu, J. ThreeDimensional Surface-Enhanced Raman Scattering Hotspots in Spherical Colloidal Superstructure for Identification and Detection of Drugs in Human Urine. Anal. Chem. 2015, 87, 4821−4828. (17) Sivanesan, A.; Kalaivani, G.; Fischer, A.; Stiba, K.; Leimkühler, S.; Weidinger, I. M. Complementary Surface-Enhanced Resonance Raman Spectroscopic Biodetection of Mixed Protein Solutions by Chitosan- and Silica-Coated Plasmon-Tuned Silver Nanoparticles. Anal. Chem. 2012, 84, 5759−5764. (18) Van Gough, D.; Wolosiuk, A.; Braun, P. V. Mesoporous ZnS Nanorattles: Programmed Size Selected Access to Encapsulated Enzymes. Nano Lett. 2009, 9, 1994−1998. (19) Roca, M.; Haes, A. J. Silica−Void−Gold Nanoparticles: Temporally Stable Surface-Enhanced Raman Scattering Substrates. J. Am. Chem. Soc. 2008, 130, 14273−14279. (20) López-Puente, V.; Abalde-Cela, S.; Angelomé, P. C.; AlvarezPuebla, R. A.; Liz-Marzán, L. M. Plasmonic Mesoporous Composites as Molecular Sieves for SERS Detection. J. Phys. Chem. Lett. 2013, 4, 2715−2720. (21) López-Puente, V.; Angelomé, P. C.; Soler-Illia, G. J. A. A.; LizMarzán, L. M. Selective SERS Sensing Modulated by Functionalized Mesoporous Films. ACS Appl. Mater. Interfaces 2015, 7, 25633−25640. (22) Kim, M. D.; Dergunov, S. A.; Richter, A. G.; Durbin, J.; Shmakov, S. N.; Jia, Y.; Kenbeilova, S.; Orazbekuly, Y.; Kengpeiil, A.; Lindner, E.; Pingali, S. V.; Urban, V. S.; Weigand, S.; Pinkhassik, E. Facile Directed Assembly of Hollow Polymer Nanocapsules within Spontaneously Formed Catanionic Surfactant Vesicles. Langmuir 2014, 30, 7061−7069. (23) Dergunov, S. A.; Richter, A. G.; Kim, M. D.; Pingali, S. V.; Urban, V. S.; Pinkhassik, E. Synergistic Self-Assembly of Scaffolds and Building Blocks for Directed Synthesis of Organic Nanomaterials. Chem. Commun. 2013, 49, 11026−11028. (24) Dergunov, S. A.; Durbin, J.; Pattanaik, S.; Pinkhassik, E. pHMediated Catch and Release of Charged Molecules with Porous Hollow Nanocapsules. J. Am. Chem. Soc. 2014, 136, 2212−2215. (25) Dergunov, S. A.; Kesterson, K.; Li, W.; Wang, Z.; Pinkhassik, E. Synthesis, Characterization, and Long-Term Stability of Hollow Polymer Nanocapsules with Nanometer-Thin Walls. Macromolecules 2010, 43, 7785−7792. (26) Danila, D. C.; Banner, L. T.; Karimova, E. J.; Tsurkan, L.; Wang, X. Y.; Pinkhassik, E. Directed Assembly of Sub-Nanometer Thin Organic Materials with Programmed-Size Nanopores. Angew. Chem., Int. Ed. 2008, 47, 7036−7039. (27) Zhang, Y.; Tekobo, S.; Tu, Y.; Zhou, Q.; Jin, X.; Dergunov, S. A.; Pinkhassik, E.; Yan, B. Permission to Enter Cell by Shape: Nanodisk Vs Nanosphere. ACS Appl. Mater. Interfaces 2012, 4, 4099− 4105. (28) Dergunov, S. A.; Pinkhassik, E. Functionalization of Imprinted Nanopores in Nanometer-Thin Organic Materials. Angew. Chem., Int. Ed. 2008, 47, 8264−8267.

(29) Dergunov, S. A.; Miksa, B.; Ganus, B.; Lindner, E.; Pinkhassik, E. Nanocapsules with ″Invisible″ Walls. Chem. Commun. 2010, 46, 1485−1487. (30) Kim, M. D.; Dergunov, S. A.; Lindner, E.; Pinkhassik, E. DyeLoaded Porous Nanocapsules Immobilized in a Permeable Polyvinyl Alcohol Matrix: A Versatile Optical Sensor Platform. Anal. Chem. 2012, 84, 2695−2701. (31) Shmakov, S. N.; Dergunov, S. A.; Pinkhassik, E. Ship-in-a-Bottle Entrapment of Molecules in Porous Nanocapsules. Chem. Commun. 2011, 47, 8223−8225. (32) Shmakov, S. N.; Pinkhassik, E. Simultaneous Templating of Polymer Nanocapsules and Entrapped Silver Nanoparticles. Chem. Commun. 2010, 46, 7346−7348. (33) Shmakov, S. N.; Jia, Y.; Pinkhassik, E. Selectively Initiated Shipin-a-Bottle Assembly of Yolk−Shell Nanostructures. Chem. Mater. 2014, 26, 1126−1132. (34) Jia, Y.; Shmakov, S. N.; Register, P.; Pinkhassik, E. Size-Selective Yolk-Shell Nanoreactors with Nanometer-Thin Porous Polymer Shells. Chem.Eur. J. 2015, 21, 12709−12714. (35) Michas, J.; Paleos, C. M.; Dais, P. Polymerization of Head and Tail Methacrylate Micelle-Forming Surfactants and Thermotropic Liquid-Crystalline Character of These Monomers and Their Polymers. Liq. Cryst. 1989, 5, 1737−1745. (36) Hamid, S. M.; Sherrington, D. C. Novel Quaternary Ammonium Amphiphilic (Meth)Acrylates: 1. Synthesis, Melting and Interfacial Behaviour. Polymer 1987, 28, 325−331. (37) Liu, S.; González, Y. I.; Kaler, E. W. Structural Fixation of Spontaneous Vesicles in Aqueous Mixtures of Polymerizable Anionic and Cationic Surfactants. Langmuir 2003, 19, 10732−10738. (38) Sanz-Ortiz, M. N.; Sentosun, K.; Bals, S.; Liz-Marzán, L. M. Templated Growth of Surface Enhanced Raman Scattering-Active Branched Gold Nanoparticles within Radial Mesoporous Silica Shells. ACS Nano 2015, 9, 10489−10497. (39) Baik, S. Y.; Cho, Y. J.; Lim, Y. R.; Im, H. S.; Jang, D. M.; Myung, Y.; Park, J.; Kang, H. S. Charge-Selective Surface-Enhanced Raman Scattering Using Silver and Gold Nanoparticles Deposited on Silicon− Carbon Core−Shell Nanowires. ACS Nano 2012, 6, 2459−2470. (40) Polavarapu, L.; Porta, A. L.; Novikov, S. M.; Coronado-Puchau, M.; Liz-Marzán, L. M. Pen-on-Paper Approach toward the Design of Universal Surface Enhanced Raman Scattering Substrates. Small 2014, 10, 3065−3071. (41) Feng, S.; Lin, D.; Lin, J.; Li, B.; Huang, Z.; Chen, G.; Zhang, W.; Wang, L.; Pan, J.; Chen, R.; Zeng, H. Blood Plasma Surface-Enhanced Raman Spectroscopy for Non-Invasive Optical Detection of Cervical Cancer. Analyst 2013, 138, 3967−3974. (42) Song, H. M.; Deng, L.; Khashab, N. M. Intracellular SurfaceEnhanced Raman Scattering (SERS) with Thermally Stable Gold Nanoflowers Grown from Pt and Pd Seeds. Nanoscale 2013, 5, 4321− 4329. (43) Muniz-Miranda, M. Surface Enhanced Raman Scattering and Normal Coordinate Analysis of 1,10-Phenanthroline Adsorbed on Silver Sols. J. Phys. Chem. A 2000, 104, 7803−7810. (44) Bonifacio, A.; Dalla Marta, S.; Spizzo, R.; Cervo, S.; Steffan, A.; Colombatti, A.; Sergo, V. Surface-Enhanced Raman Spectroscopy of Blood Plasma and Serum Using Ag and Au Nanoparticles: A Systematic Study. Anal. Bioanal. Chem. 2014, 406, 2355−2365. (45) Antipov, A. A.; Sukhorukov, G. B.; Leporatti, S.; Radtchenko, I. L.; Donath, E.; Möhwald, H. Polyelectrolyte Multilayer Capsule Permeability Control. Colloids Surf., A 2002, 198−200, 535−541. (46) Antipov, A. A.; Sukhorukov, G. B. Polyelectrolyte Multilayer Capsules as Vehicles with Tunable Permeability. Adv. Colloid Interface Sci. 2004, 111, 49−61. (47) Alvarez-Puebla, R. A.; Arceo, E.; Goulet, P. J. G.; Garrido, J. J.; Aroca, R. F. Role of Nanoparticle Surface Charge in Surface-Enhanced Raman Scattering. J. Phys. Chem. B 2005, 109, 3787−3792. (48) Stewart, A.; Murray, S.; Bell, S. E. J. Simple Preparation of Positively Charged Silver Nanoparticles for Detection of Anions by Surface-Enhanced Raman Spectroscopy. Analyst 2015, 140, 2988− 2994. H


Research Article

ACS Applied Materials & Interfaces (49) Tian, L.; Tadepalli, S.; Farrell, M. E.; Liu, K.-K.; Gandra, N.; Pellegrino, P. M.; Singamaneni, S. Multiplexed Charge-Selective Surface Enhanced Raman Scattering Based on Plasmonic Calligraphy. J. Mater. Chem. C 2014, 2, 5438−5446. (50) Alvarez-Puebla, R. A.; Aroca, R. F. Synthesis of Silver Nanoparticles with Controllable Surface Charge and Their Application to Surface-Enhanced Raman Scattering. Anal. Chem. 2009, 81, 2280− 2285. (51) Sun, Z.; Du, J.; Yan, L.; Chen, S.; Yang, Z.; Jing, C. Multifunctional Fe3O4@SiO2−Au Satellite Structured SERS Probe for Charge Selective Detection of Food Dyes. ACS Appl. Mater. Interfaces 2016, 8, 3056−3062. (52) Bryant, M. A.; Crooks, R. M. Determination of Surface pKa Values of Surface-Confined Molecules Derivatized with pH-Sensitive Pendant Groups. Langmuir 1993, 9, 385−387. (53) Sun, F.; Zhang, P.; Bai, T.; David Galvan, D.; Hung, H.-C.; Zhou, N.; Jiang, S.; Yu, Q. Functionalized Plasmonic Nanostructure Arrays for Direct and Accurate Mapping Extracellular pH of Living Cells in Complex Media Using SERS. Biosens. Bioelectron. 2015, 73, 202−207. (54) Sun, F.; Ella-Menye, J.-R.; Galvan, D. D.; Bai, T.; Hung, H.-C.; Chou, Y.-N.; Zhang, P.; Jiang, S.; Yu, Q. Stealth Surface Modification of Surface-Enhanced Raman Scattering Substrates for Sensitive and Accurate Detection in Protein Solutions. ACS Nano 2015, 9, 2668− 2676. (55) Bordoni, A. V.; Lombardo, M. V.; Regazzoni, A. E.; Soler-Illia, G. J. A. A.; Wolosiuk, A. Simple Thiol-Ene Click Chemistry Modification of SBA-15 Silica Pores with Carboxylic Acids. J. Colloid Interface Sci. 2015, 450, 316−324.

I


Controlled Permeability in Porous Polymer Nanocapsules Enabling

Recommend Documents