Metal Nanoparticle-Loaded Microgels with Selective Permeability for

Feb 5, 2016 - Gayoung Eom , Hongki Kim , Ahreum Hwang , Hye-Young Son , Yuna Choi , Jeong Moon , Donghyeong Kim , Miyeon Lee , Eun-Kyung Lim ...
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Metal Nanoparticle-Loaded Microgels with Selective Permeability for Direct Detection of Small Molecules in Biological Fluids Dong Jae Kim,† Tae Yoon Jeon,† Youn-Kyoung Baek,‡ Sung-Gyu Park,‡ Dong-Ho Kim,*,‡ and Shin-Hyun Kim*,† †

Department of Chemical and Biomolecular Engineering, Korea Advanced Institute of Science and Technology (KAIST), Daejeon 305-701, Republic of Korea ‡ Advanced Functional Thin Films Department, Korea Institute of Materials Science (KIMS), Changwon, Gyeongnam 641-831, Republic of Korea S Supporting Information *

ABSTRACT: We report a microfluidic strategy for creating semipermeable microgels containing metal nanoparticles to directly detect small molecules included in the solution of large adhesive proteins using surface-enhanced Raman scattering. With a capillary microfluidic device, gold nanoparticle-laden microgels are prepared to have uniform size. The microgels allow diffusion of smaller molecules than mesh size of their gel network while excluding larger molecules. This enables the selective infusion of small analytes onto the surface of gold nanoparticles from the solutions of adhesive proteins, thereby providing high Raman intensity by metal-surface enhancement; otherwise, proteins adsorb the surface, which significantly reduces the intensity. Therefore, this microgel platform enables the direct detection of analytes from biological fluids and obviates complicated pre- or post-treatment of samples. In addition, the microgels are able to be injected into target volume such as vessels or living organisms, which are then either recovered for analysis or potentially analyzed in situ. This simple but pragmatic method will provide new opportunity in a wide range of molecular detection applications based on Raman spectrum.

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INTRODUCTION The molecules are characterized by discrete vibrational energy states. Photons that are inelastically scattered by molecules lose or gain certain levels of energy due to transfer among the energy states during the scattering event in a process referred to as Raman scattering.1 Raman scattering provides a moleculespecific spectrum and serves as a fingerprint for molecules. A sufficient Raman scattering intensity must be acquired for molecular characterization, necessitating high concentrations of molecules and a high intensity of monochromic light. Technical limitations on achieving such conditions may be overcome by employing metal nanostructures that strongly localize an electromagnetic field at their surface through the collective oscillations of electrons,2 thereby providing a highly enhanced Raman scattering signal that is measurable using common spectrometers. This technique is referred to as surfaceenhanced Raman scattering (SERS).3,4 The magnitude of the intensity enhancement may reach values as high as O(1012) through the design of metal nanostructures with nanogaps,5−7 sharp tips,8−10 and rough surfaces;11−13 these structures strongly couple electromagnetic waves at the surface. A variety of SERS-active substrates and nanoparticles have been developed to provide higher enhancement factors, and some of these systems are commercially available for molecular detection. However, complicated sample pre- and sometimes post-treatments are required, thereby restricting the use of © XXXX American Chemical Society

SERS. Pretreatment usually involves a dialysis process using a semipermeable membrane to exclude large adhesive molecules from samples. Post-treatment typically involves a process of concentrating nanoparticles to obtain high-density metal structures. In particular, the selective removal of proteins is a prerequisite for biological fluids extracted from living organisms to prevent irreversible and rapid adhesion on the metal surface and to allow noncompetitive access by small analyte molecules. Pretreatments frequently involve significant dilution or loss of analytes, thereby reducing the signal-to-noise ratio.14−18 Although metal nanoparticles, encapsulated with silica shell, are used to exclude adhesive protein, dense shells only allow diffusion of very small ions, achieving restricted applications.19 Here, we have embedded gold nanoparticles in a matrix of hydrogel microparticles that have consistent mesh size, thereby allowing the infusion of small molecules to the gold nanoparticles while excluding large molecules. A microfluidic device was used to form a uniform emulsified aqueous suspension of gel precursors containing gold nanoparticles in a continuous oil phase, which was then photocross-linked to immobilize the gold nanoparticles in the gel. The microgels provided a clear suspended molecule size cutoff threshold for Received: January 11, 2016 Revised: February 5, 2016

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DOI: 10.1021/acs.chemmater.6b00115 Chem. Mater. XXXX, XXX, XXX−XXX

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Chemistry of Materials permeation through the gel network, thereby enabling the selective exclusion of molecules larger than the mesh size. This method allowed the Raman spectra of small molecules to be obtained directly from mixtures containing large proteins while avoiding any deterioration in the intensity, thereby obviating pretreatment. The intensity could be further improved simply by removing water from the microgels (drying), which concentrated the gold nanoparticles in each microparticle. Microgels could be injected using a needle and could be recovered using a magnet by embedding magnetic particles in the microgels.

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RESULTS AND DISCUSSION Microfluidic Production of Gold Nanoparticle-Embedded Microgels. The essential strategy of our approach relies on the formation of a water-saturated hydrogel network having a uniform mesh size to provide size-selective permeability through the gel network, as illustrated in Figure 1, panel a. At the same time, the molecules comprising the gel matrix should display minimal inelastic scattering to provide unperturbed Raman spectra of the small molecules from the gel matrix. These two requirements were satisfied by preparing microgels composed of a poly(ethylene glycol) (PEG) backbone by photopolymerizing precursors in water-in-oil emulsion drops. Gold nanoparticles were captured in the matrix of the uniform microgels by emulsifying a mixture of poly(ethylene glycol)diacrylate (PEGDA) and water containing 1 w/w% photoinitiator and 0.8 w/w% gold nanoparticles in hexadecane containing 5 w/w% surfactant in a capillary microfluidic device to produce monodisperse emulsion drops, as shown in Figure 1, panels b and c. The gold nanoparticles had an average diameter of 38 nm, as shown in Figure S1 of the Supporting Information, sufficiently large for capture in the gel matrix. The emulsion drops flowed through the capillary channel and were irradiated with ultraviolet (UV) light during collection in a vial. This process produced homogeneous cross-linking among the PEGDA molecules to form a microgel in drops. The microgel was then transferred from hexadecane to isopropyl alcohol and finally to water. The microgel was highly uniform and transparent, as shown in Figure 1, panel d. Gold nanoparticles were homogeneously embedded in the microgels without significant aggregation, as confirmed in the absorption spectra presented in Figure 1, panel e. Gold nanoparticles embedded in the microgel exhibited an absorption peak at 531 nm that coincided with the peaks obtained from a uniform aqueous dispersion of the nanoparticles. No broadening or second peak formation was observed, suggesting negligible aggregation. Size-Selective Permeability of Microgels. Homogeneous cross-linking provides a consistent mesh size and sizeselective permeability. The cutoff permeation value depended on the mesh size, which could be controlled by varying the mixing ratio of PEGDA and water in the drops. Because PEGDA gel fills whole volume of water drop, as the fraction of PEGDA in the drop varies, so does the number of cross-links in the gel volume (cross-linking density). Higher cross-linking density results in tighter mesh and smaller cutoff value. Microgels prepared from a PEGDA/water weight ratio of 1:9 possessed a relatively low cross-linking density that permitted the infusion of fluorescein isothiocyanate (FITC)-tagged dextran with a molecular weight (Mw) of 40 000 g mol−1 and a hydrodynamic diameter of 9 nm while excluding dextran with a Mw of 70 000 g mol−1 and a hydrodynamic diameter of 12 nm, as shown in Figure S2a; the microgels were incubated in

Figure 1. (a) Schematic illustration of a semipermeable microgel containing gold nanoparticles (Au NPs) that allow for the sizeselective infusion of materials from the surrounding medium. (b, c) Schematic diagram and still-shot optical microscopy (OM) image of a capillary microfluidic device, showing the generation of monodisperse water-in-oil emulsion droplets containing Au NPs and PEGDA. In situ UV irradiation cross-links the PEGDA molecules in the droplet. (d) OM images of monodisperse microgels dispersed in water. (e) Normalized absorption spectra of aqueous suspensions of Au NPs and microgels containing Au NPs. (f) Time dependence of the normalized fluorescence intensity, (I − I0)/(Imax − I0), within the microgels dispersed in an aqueous solution of dye molecules of three different sizes: sulforhodamine B (Mw 558.67), rhodamine B isothiocyanate (RITC)-tagged dextran (Mw 10 000), and fluorescein isothiocyanate (FITC)-tagged dextran (Mw 20 000), where I0 and Imax are the intensities at the beginning of the measurement (t = 0) and the maximum intensity, respectively. Confocal microscopy images of microgels dispersed in aqueous solutions of sulforhodamine B (top) and RITC-dextran (bottom), collected at the denoted times.

a solution for 1 day prior to imaging. Therefore, the cutoff value was estimated to fall between 9 and 12 nm. A PEGDA/water weight ratio of 3:7 yielded a microgel cutoff value of 7−9 nm, as shown in Figure S2b. Microgels prepared using a PEGDA/ water weight ratio of 7:3 yielded a cutoff value of 5−7 nm, as shown in Figure S2c. For all three microgels, fluorescent intensity of permeable FITC-tagged dextran was uniform across the microgels, indicating that the microgels were homogeneously cross-linked. Selective infusion for microgels with the ratio of 7:3 was further confirmed using a mixture of rhodamine B isothiocyanate (RITC)-dextran with a Mw of 10 000 g mol−1 and FITC-dextran with a Mw of 20 000 g mol−1, as shown in the Figure S3. The size of microgels in hexadecane is the same as the size of emulsion drops, and that in water is slightly influenced by the mixing ratio because higher cross-linking density yields lower degree of swelling. B

DOI: 10.1021/acs.chemmater.6b00115 Chem. Mater. XXXX, XXX, XXX−XXX

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Chemistry of Materials Microgels prepared from a PEGDA/water weight ratio of 7:3 were selected to exclude proteins with a diameter exceeding 5 nm. The microgel permeation rates of three molecules having different sizes, sulforhodamine B (Mw 558.67 g mol−1), RITCdextran with a Mw of 10 000 g mol−1, and FITC-dextran with a Mw of 20 000 g mol−1, were investigated, as shown in Figure 1, panel f and Figure S4. The fluorescence intensity in the microgel, I(t), increased from a small initial value, I0, as the molecules diffused into the microgels from surrounding fluid, reaching a plateau, Imax. The time dependence of I(t) followed the diffusion equation:20 I − I0 6 =1− 2 Imax − I0 π

∞

∑n= 1

1 −n2π 2Dt / R2 e n2

(1)

where R is microgel radius, and the only fitting parameter is D, the diffusivity of molecules in the gel. The fits yielded D values of 7.34 μm2 s−1 for sulforhodamine B, 0.836 μm2 s−1 for RITCdextran, and 0 for the impermeable FITC-dextran. The diffusivity of a molecules in a dilute solution may be estimated using the Stokes−Einstein equation, D = kBT(3πηd)−1, and this value was 218 μm2 s−1 for sulforhodamine B and 95 μm2 s−1 for FITC-dextran, where kB is the Boltzmann constant, T is the absolute temperature, η is the viscosity, and d is the hydrodynamic diameter of the molecule. The diffusivity in the microgel, obtained from the fits, was 3.37% of the Stokes− Einstein diffusivity for sulforhodamine B and 0.88% for RITCdextran, indicating that the diffusion of molecules with a diameter comparable to the mesh size was highly suppressed in the gel network. The fluorescence intensity from sulforhodamine B reached a plateau within about 1 min, a time scale that is sufficiently short to permit the rapid detection of small molecules. Wet Microgels and Dried Particles for SERS Analysis. The microgels were formed in hexadecane prior to transferal into water. During the transfer, the microgels swelled further. Microgels made from 70 w/w% PEGDA had an average diameter of 70.2 μm in hexadecane. After transfer, the microgels had an average diameter of 76.6 μm, with a coefficient of variation as small as 1.14%. Therefore, the microgels were composed of 45.2 w/w% water when immersed in an aqueous environment, as estimated from material balance calculations. The water could be completely removed from the microgels to form spherical solid particles composed of a collapsed network with an average diameter of 61.7 μm, slightly larger than the diameter calculated from the mass balance, 60.4 μm, assuming complete collapse of the network. The images of the microgels in hexadecane, water, and air are shown in Figure 2, panel a and Figure S5, and the corresponding size distributions of the samples are shown in Figure 2, panel b. Both the microgels and the dried particles contain homogeneously embedded gold nanoparticles and provided a SERS substrate for adsorbed molecules. To ensure that the gel material provided negligible Raman scattering signals, we measured the Raman spectra of both the microgels and the dried particles without any additives using a dispersive Raman system equipped with a 633 nm laser at a power of 6.5 mW, as shown in Figure S6. A focal volume diameter of 1 μm, depth of 1.8 μm, and an integration time of 10 s were used throughout this work. The microgels did not display any meaningful peaks over the Raman shift range of 200−1700 cm−1. Although dried particles displayed several peaks in the same range, the intensities of these peaks were low and did not influence the

Figure 2. (a, b) OM images and size distribution of microgels in hexadecane and water and dried particles in the air. (c) Raman spectra of rhodamine 6G (R6G), measured using the wet microgel, dried particle, or surrounding solution of 1 μM R6G. (d) Concentration dependence of the Raman peak intensity at 613 cm−1, measured from the wet microgels and dried particles.

Raman spectra of the analytes. The SERS activity of the microgel was evaluated using rhodamine 6G (R6G, Mw 479.02 g mol−1) as an analyte molecule. The microgels were dispersed in an aqueous solution of R6G at a concentration of 1 μM. The Raman spectrum measured from the microgel exhibited peaks characteristic of R6G at 613, 774, 1187, 1310, 1363, 1510, and 1649 cm−1, as expected. Several weak peaks were attributed to the formation of a complex between the positively charged R6G and the partially negative gel matrix. Use of the benzenethiol (BT, Mw 110.19 g mol−1) analyte did not yield these peaks, as shown in Figure S7 of the Supporting Information. Five arbitrarily selected microgel samples revealed indistinguishable Raman spectral intensities, as shown in Figures S7b and S8, indicating a high uniformity in the microgels; Raman spectra were measured from the centers of microgels. Coefficients of variation of peak intensities at 613 cm−1 for R6G and at 999 cm−1 for BT were 4.07% and 3.72%, respectively. In addition, Raman intensity weakly depends on position of measurement spot within single microgel except the edge, as shown in Figure S9a,b, indicating homogeneity in the nanoparticle dispersion; a focal volume for measurement of Raman spectra is much larger than nanoparticle diameter of 38 nm and much smaller than microgel diameter of 76.6 μm. No peaks in the Raman spectrum were derived from the surrounding solution due to the absence of gold nanoparticles. When the concentration of gold nanoparticles in the PEGDA solution was increased from 0.8 w/w% to 2 w/w%, the nanoparticles formed aggregates, which yielded variation of Raman intensity depending on measurement spot position, as shown in Figure S10; the use of concentration 0.4 w/w% reduced Raman intensity. The peak intensity could be further improved by drying the microgel. Dried particles exhibited a 3.6-fold greater peak intensity than the wet microgels. In this measurement, the surrounding solution was carefully removed using wipes prior to drying to prevent further infusion of R6G. A 1.91-fold C

DOI: 10.1021/acs.chemmater.6b00115 Chem. Mater. XXXX, XXX, XXX−XXX

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Chemistry of Materials increase in the gold nanoparticle density accompanied the drying process, estimated based on the particle size reduction. The additional 1.88-fold intensity enhancement was attributed to the formation of SERS-active nanogaps between neighboring nanoparticles despite very low density. We further investigated the limit of detection (LOD) for both the wet microgels and the dried microparticles. By reducing the concentration of R6G from 1 mM to 10 nM, the Raman intensity obtained from the wet microgels monotonically decreased, as shown in Figure 2, panel d and Figure S11. No meaningful signals were obtained from a 1 nM R6G solution, indicating that the LOD was roughly 10 nM. Dried microparticles showed higher Raman signal intensities at all concentrations and a very small but detectable intensity even at a concentration of 1 nM. The value of the LOD is usually affected by the enhancement factor of the metal nanostructures. Therefore, the LOD may be significantly reduced simply by replacing the spherical gold nanoparticles with anisotropic metal nanoparticles such as polyhedral and star-shaped nanoparticles.21,22 The drying and swelling of the microgels are highly reversible. Therefore, the microgels can be stored in dried state and used in wet state. To prove this, we completely dried the microgels and stored them for 2 days. Afterward, the dried microparticles were redispersed in water. Upon the hydration, the microgels recovered the original size, as shown in Figures S12a−c. In addition, hydrated microgels showed comparable Raman intensity with that of the original microgels, as shown in Figure S12d. Detection of Small Molecules from Protein Solution. The microgels with a clear permeation cutoff value can prevent the diffusion of large protein molecules into the gel matrix, thereby selectively allowing for the infusion of small molecules. This effect was demonstrated by dispersing microgels made using a 7:3 PEGDA/water weight ratio into a binary mixture of 10 μM R6G and 1 mM bovine serum albumin (BSA, Mw 66 463 g mol−1). Albumin is a major protein component of blood plasma and is widely used for surface passivation in biological immunoassays due to its nonspecific adhesive properties.23 The Stokes diameter of BSA is 7 nm.24 BSA binds strongly to the surfaces of citrate-coated gold nanoparticles.25,26 Gold nanoparticles directly dispersed in water were not protected against a BSA and therefore became covered with BSA in the binary mixture, as illustrated in Figure 3, panel a. Therefore, the R6G molecules could not effectively access the gold nanoparticle surface, which considerably reduced the Raman intensity relative to that measured from the gold nanoparticles dispersed in a solution of R6G only, as shown in Figure 3, panel b. By contrast, the microgels with a permeation cutoff value of 5−7 nm effectively excluded BSA from the gel matrix while allowing for the infusion of R6G, as illustrated in Figure 3, panel c. The Raman spectrum intensity measured from the microgels dispersed in the binary mixture was comparable to that measured using the microgels dispersed in the aqueous solution of R6G, at a fixed R6G concentration of 10 μM, as shown in Figure 3, panel d. Microgels can provide Raman signal of multiple small molecules ingredients while excluding large protein molecules. For example, when the microgels were dispersed in ternary mixture of 0.1 μM Malachite green (MG, Mw 364.91 g mol−1), 1 μM R6G, and 1 mM BSA, they selectively allowed the infusion of R6G and MG, as illustrated in Figure 3, panel e; MG is a synthetic chemical used as gemicide in aquarium and fabric dye but prohibited by the FDA because of its toxicity. As a result, the

Figure 3. (a) Schematic diagram showing the Au NPs dispersed in an aqueous solution of small molecules and large adhesive proteins. (b) Raman spectra measured from the Au NPs directly dispersed in an aqueous solution of 10 μM R6G and 1 mM BSA, and an aqueous solution of 10 μM R6G. The Raman intensity of the R6G was significantly reduced in the BSA solution. (c) Schematic diagram showing the semipermeable microgels, which permit the diffusion of small molecules while excluding large adhesive proteins. (d) Raman spectra measured from the wet microgels dispersed in an aqueous solution containing 10 μM R6G and 1 mM BSA, or in an aqueous solution containing 10 μM R6G only. The two spectra were comparable. (e) Schematic diagram showing the selective diffusion of two different small molecules while excluding large adhesive proteins. (f) Raman spectra measured from the wet microgels dispersed in an aqueous solution containing 0.1 μM Malachite green (MG), 1 μM R6G, and 1 mM BSA, in an aqueous solution containing 0.1 μM MG and 1 μM R6G, and in an aqueous solution containing 0.1 μM MG only, respectively. Characteristic peak positions of MG are denoted with vertical lines.

Raman spectrum obtained from the microgel possesses characteristic peaks from both small moelcules of MG and R6G, of which intensity is comparable to that measured in a binary mixture of MG and R6G in absence of BSA, as shown in Figure 3, panel f; the Raman spectrum measured from microgel in aqueous solution of MG is also included to identify the characteristic peaks of MG. The above three sets of Raman measurements revealed that the microgels could exclude large protein molecules while permitting the diffusion of small analytes, thereby providing a method for directly detecting small molecules in biological fluids containing large adhesive molecules. D

DOI: 10.1021/acs.chemmater.6b00115 Chem. Mater. XXXX, XXX, XXX−XXX

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Detection of Analytes in Milk Using Magnetoresponsive Microgels. The microgels could be further functionalized to be magnetoresponsive by embedding magnetic particles into the gel matrix. To this end, we dispersed magnetic particles with a diameter of 100 nm at a concentration of 0.2 w/w% in the mixture of PEGDA and water containing 0.8 w/w% gold nanoparticles. This suspension was used as a dispersed phase. The resulting microgels were found to be magnetoresponsive, as shown in Figure S13. The microgels could be injected using a needle into any target volume. For example, we injected them into whole milk containing 0.1 mM BT using a needle, as shown in the left panel of Figure 4, panel a. The microgels could be recovered

CONCLUSION Infinitesimal amounts of small molecules may be crucial to the viability and healthcare of living organisms so that detection of such molecules in biological fluids is important. Although SERS can provide a useful means for identifying molecules without the need for tagging, virgin metal surfaces are prone to contamination by the preferential adsorption of large proteins that interrupt the detection. Microgels containing metal nanoparticles obviate this contaminating adsorption by selectively allowing the infusion of small molecules, thereby providing a strong SERS spectrum without the need for pretreatments. To further enhance SERS activity and reduce LOD, metal nanoparticles with nanogaps or sharp tips can be used. We believe our microgel platform benefits point-of-care analysis of biomarkers or toxic chemicals that may be present in biological fluids and foods as complicate pretreatment of samples is evitable. In the future aspect, the microgels may be hypodermically injected into a body and used for in vivo detection by embedding metal nanoparticles with strong SERS activity in near-infrared (NIR) region.27−30

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EXPERIMENTAL SECTION

Preparation of a Capillary Microfluidic Device. Monodisperse water-in-oil emulsion droplets were prepared using a glass capillary microfluidic device composed of two tapered cylindrical capillaries nested in a square capillary.31 One of the cylindrical capillaries (1B100F-6, World Precision Instruments, Inc.) was tapered by a capillary puller (P97, Sutter Instrument) and sanded to yield a 40 μm diameter orifice, while the other capillary was prepared to have a 100 μm diameter orifice. These capillaries were treated with trimethoxy(octadecyl)silane (Sigma-Aldrich) to render them hydrophobic and were then coaxially aligned to have a separation distance of 60 μm within the square capillary (Atlantic International Technologies, Inc.). Fluids were injected through the square capillary by cutting the plastic bottom part of a dispensing needle (75165A677, McMaster-Carr) to fit the entrance of the square capillary, followed by sealing with epoxy resin. Synthesis of the Gold Nanoparticles. Gold nanoparticles with an average diameter of 38 nm were synthesized using the sodium citrate reduction method.32 To this end, an aqueous solution containing 0.01 w/w% gold(III) chloride hydrate (HAuCl4 (aq), 99.99%, Aldrich) was boiled with stirring and refluxing in a roundbottom flask, and 1.5 mL of an aqueous solution containing 1 w/w% sodium citrate tribasic dehydrate (Sigma-Aldrich) was quickly added to the boiling solution. After 30 min, the solution was cooled to room temperature. The shapes and sizes of the nanoparticles were characterized by transmission electron microscopy (JEM-2100 F, JEOL). Fabrication of the Microgels. Gold nanoparticles were dispersed to an 0.8 w/w% concentration in a mixture of PEGDA and water containing 1 w/w% photoinitiator, 1-[4-(2-hydroxyethoxy)-phenyl]-2hydroxy-2-methyl-1-propane-1-one (Irgacure 2959, BASF). The cutoff permeation value was adjusted by varying the weight ratio of PEGDA to water, 1:9, 3:7, and 7:3. The suspension was injected through a cylindrical capillary with a 40 μm diameter orifice at a volumetric flow rate of 100 μL h−1 using a syringe pump (Legato 100, KD Scientific) to form emulsion drops. A hexadecane continuous phase containing 5 w/w% ABIL EM90 surfactant (Evonik Industries) was injected through the interstices between the cylindrical and square capillaries at a volumetric flow rate of 2200 μL h−1. The suspension at the tip of the cylindrical capillary was emulsified in a dripping mode to form monodisperse drops, whereas the continuous phase was focused into the 100 μm diameter orifice of the other cylindrical capillary, which exerted a high drag force on the hanging drop on the tip of the injection capillary. Drop generation was observed using a high-speed camera (Phantom v7.3, Vision Research Inc.) mounted on an inverted optical microscope (Eclipse TS100, Nikon). The drops flowed through

Figure 4. (a) A set of images showing the injection of magnetoresponsive microgels into whole milk containing benzenethiol (BT) (left panel) and the recovery of the microgels using a magnet (middle and right panels). (b) Raman spectra of BT measured using wet microgels immersed in milk or aqueous solutions, where both solutions include 0.1 mM BT. (c) Optical microscopy image of the microgels dispersed in whole milk.

from the milk by gently stirring the solution with a magnet connected to a stick, as shown in the middle and right panels. The microgels attached to the surface of the magnet and were recovered for SERS measurements. The Raman spectrum of the BT molecules could be obtained from the recovered microgel, as shown in Figure 4, panel b. The microgels were injected and recovered from the aqueous solution of BT at the given concentration to yield a nearly identical Raman spectral intensity. By contrast, gold nanoparticles dispersed directly in the milk containing BT exhibited a Raman signal intensity that was a factor of 1/3.3-times lower than the corresponding intensity obtained from the BT dispersed in water at the same concentration, as shown in Figure S14. This set of experiments indicated that the microgels could effectively exclude large water-soluble proteins, casein micelles, and colloids of butterfat globules from the microgels while also permitting the infusion of small BT molecules. Optical microscopy images of the microgels dispersed in milk revealed the exclusion of micelles and colloids, as shown in Figure 4, panel c. E

DOI: 10.1021/acs.chemmater.6b00115 Chem. Mater. XXXX, XXX, XXX−XXX

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Chemistry of Materials the collection capillary and were collected in a glass vial that was continuously UV-irradiated using a fiber-coupled spot UV system (Inocure 100N, Lichtzen Co., Ltd.) to polymerize the PEGDA in the drops. The resultant microgels were washed away using isopropyl alcohol several times prior to being transferred to water. The microgels were rendered magnetoresponsive by dispersing magnetic particles 100 nm in diameter (MTX-INK(H), Nanobrick) in the mixture of PEGDA and water at a concentration of 0.2 w/w%. Characterization of the Microgels. The size distributions of the microgels in three different surrounding media (hexadecane, water, and air) were evaluated based on the optical microscopy images collected from more than 100 microgels. Dried microparticles were observed using scanning electron microscopy (S-4800, Hitachi) after application of an OsO4 coating. The absorption spectra were measured by deposing the microgels onto a glass substrate to form a hexagonally packed monolayer. The spectra were measured using a UV−vis spectrometer (Infinite M200 Pro, Tecan). The absorption spectrum of the gold nanoparticles dispersed in water was measured using the same equipment. The cutoff permeation value and diffusivity of the molecules in the gel matrix were measured using a variety of fluorescent molecules: sulforhodamine B (Mw 558.67 g mol−1, SigmaAldrich), rhodamine B isothiocyanate-tagged dextran (Mw 10 000 g mol−1, Sigma-Aldrich), fluorescein isothiocyanate-tagged dextran (Mw 10 000, 20 000, 40 000, and 70 000 g mol−1, Sigma-Aldrich). The cutoff values were measured by dispersing the microgels in an aqueous solution containing the single or binary fluorescent molecules for 1 day and then imaging them using a confocal laser scanning microscope (LSM 510, Carl Zeiss). The diffusivity of each microgel was measured by continuously imaging the samples as soon as the microgel medium had been replaced with the dye solution. The time-dependent variations in the fluorescence intensities measured from the microgels were then fit to the diffusion equation to estimate the diffusivity. Measurements of the Raman Spectra. Three different small target molecules were used to obtain the Raman spectra from the microgels: Rhodamine 6G (R6G, Sigma-Aldrich), benzenethiol (BT, Sigma-Aldrich), and Malachite green (MG, Sigma-Aldrich). The microgels were dispersed in an aqueous solution of the molecules for 1 h, from which Raman spectra were measured using a dispersive Raman system (Horiba Jobin Yvon) with a laser wavelength of 633 nm, a spot diameter of 1 μm, and a power of 6.5 mW. The signal was acquired for 10 s. In addition, Raman spectra were measured using the microparticles in air after blot-drying the excess medium and drying the microparticles. To demonstrate the selective permeation of small molecules and their detection, we dispersed either the microgels or the gold nanoparticles (the gold nanoparticles were dispersed at a concentration of 0.8 w/w%) in a binary mixture of 10 μM R6G and 1 mM bovine serum albumin (BSA, Sigma-Aldrich) for 1 h. The Raman spectra of the suspensions were then measured. Either the microgels or the gold nanoparticles were dispersed in a solution of R6G to compare the Raman intensities with their counterparts. In addition, we measure Raman spectra from microgels dispersed in a ternary mixture of 0.1 μM MG, 1 μM R6G, and 1 mM BSA, a binary mixture of 0.1 μM MG and 1 μM R6G, and a solution of 0.1 μM MG. The selective permeability of the microgels was further examined using a suspension of BT dissolved in whole milk (Seoul Milk Co.) or in water at the same concentration of 0.1 mM. Either magnetoresponsive microgels or gold nanoparticles were injected into the solutions using a needle (22G, Korea Vaccine Co.). After 1 h of incubation, the microgels were recovered by stirring the solution with a magnet, and the Raman spectra of the microgels were measured. A thin film of the gold nanoparticle samples was used for the Raman measurements, but otherwise the signal was very low due to scattering by the milk.

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Confocal, optical microscope images of microgel and additional SERS spectra (PDF)

AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by the Fundamental Research Program (PNK 4150) of the Korean Institute of Materials Science (KIMS) and the Midcareer Researcher Program (2014R1A2A2A01005813) and Global Research Laboratory (NRF-2015K1A1A2033054) through the National Research Foundation (NRF) grant funded by the Ministry of Science, ICT, and Future Planning (MSIP).

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REFERENCES

(1) Kneipp, K.; Kneipp, H.; Itzkan, I.; Dasari, R. R.; Feld, M. S. Ultrasensitive chemical analysis by raman spectroscopy. Chem. Rev. 1999, 99, 2957−2975. (2) Barnes, W. L.; Dereux, A.; Ebbesen, T. W. Surface plasmon subwavelength optics. Nature 2003, 424, 824−830. (3) Nie, S.; Emory, S. R. Probing single molecules and single nanoparticles by surface-enhanced raman scattering. Science 1997, 275, 1102−1106. (4) Kneipp, K.; Wang, Y.; Kneipp, H.; Perelman, L. T.; Itzkan, I.; Dasari, R. R.; Feld, M. S. Single molecule detection using surfaceenhanced raman scattering (SERS). Phys. Rev. Lett. 1997, 78, 1667− 1670. (5) Hatab, N. A.; Hsueh, C.-H.; Gaddis, A. L.; Retterer, S. T.; Li, J.H.; Eres, G.; Zhang, Z.; Gu, B. Free-standing optical gold bowtie nanoantenna with variable gap size for enhanced raman spectroscopy. Nano Lett. 2010, 10, 4952−4955. (6) Wang, H.; Levin, C. S.; Halas, N. J. Nanosphere Arrays with Controlled Sub-10-nm Gaps as Surface-Enhanced Raman Spectroscopy Substrates. J. Am. Chem. Soc. 2005, 127, 14992−14993. (7) Lim, D. K.; Jeon, K.-S.; Hwang, J.-H.; Kim, H.; Kwon, S.; Suh, Y. D.; Nam, J.-M. Highly uniform and reproducible surface-enhanced Raman scattering from DNA-tailorable nanoparticles with 1-nm interior gap. Nat. Nanotechnol. 2011, 6, 452−460. (8) Wang, P.; Liang, O.; Zhang, W.; Schroeder, T.; Xie, Y. H. Ultrasensitive graphene-plasmonic hybrid platform for label-free detection. Adv. Mater. 2013, 25, 4918−4924. (9) Lu, Y.; Liu, G. L.; Kim, J.; Mejia, Y. X.; Lee, L. P. Nanophotonic Crescent Moon Structures with Sharp Edge for Ultrasensitive Biomolecular Detection by Local Electromagnetic Field Enhancement Effect. Nano Lett. 2005, 5, 119−124. (10) Kelly, K. L.; Coronado, E.; Zhao, L. L.; Schatz, G. C. The optical properties of metal nanoparticle: the influence of size, shape, and dielectric environment. J. Phys. Chem. B 2003, 107, 668−677. (11) Zhang, L.; Lang, X. Y.; Hirata, A.; Chen, M. W. Wrinkled nanoporous gold films with ultrahigh surface-enhanced raman scattering enhancement. ACS Nano 2011, 5, 4407−4413. (12) Liang, H. Y.; Li, Z. P.; Wang, W. Z.; Wu, Y. S.; Xu, H. X. Highly surface-roughened “Flower-like” silver nanoparticles for extremely sensitive substrates of surface-enhanced raman scattering. Adv. Mater. 2009, 21, 4614−4618. (13) Wang, H.; Halas, N. J. Mesoscopic Au “Meatball” particles. Adv. Mater. 2008, 20, 820−825. (14) Zhang, X.-F.; Zou, M.-Q.; Qi, X.-H.; Liu, F.; Zhu, X.-H.; Zhao, B.-H. Detection of melamine in liquid milk using surface-enhanced raman scattering spectroscopy. J. Raman Spectrosc. 2010, 41, 1655− 1660.

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DOI: 10.1021/acs.chemmater.6b00115 Chem. Mater. XXXX, XXX, XXX−XXX

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Chemistry of Materials (15) Kim, A.; Barcelo, S. J.; Williams, R. S.; Li, Z. Melamine sensing in milk products by using surface enhanced raman scattering. Anal. Chem. 2012, 84, 9303−9309. (16) Qu, J. Y.; Wilson, B. C.; Suria, D. Concentration measurements of multiple analytes in human sera by near-infrared laser raman spectroscopy. Appl. Opt. 1999, 38, 5491−5498. (17) Smith, W. E. Practical understanding and use of surface enhanced raman scattering/surface enhanced resonance raman scattering in chemical and biological analysis. Chem. Soc. Rev. 2008, 37, 955−964. (18) Li, X.; Lee, H. K.; Phang, I. Y.; Lee, C. K.; Ling, X. Y. Superhydrophobic-oleophobic Ag nanowire platform: an analyteconcentrating and quantitative aqueous and organic toxin surfaceenhanced raman scattering sensor. Anal. Chem. 2014, 86, 10437− 10444. (19) Wang, F.; Widejko, R. G.; Yang, Z.; Nguyen, K. T.; Chen, H.; Fernando, L. P.; Christensen, K. A.; Anker, J. N. Surface-Enhanced Raman Scattering Detection of pH with Silica-Encapsulated 4Mercaptobenzoic Acid-Functionalized Silver Nanoparticles. Anal. Chem. 2012, 84, 8013−8019. (20) Crank, J. The Mathematics of Diffusion; Oxford University Press: Oxford, UK, 1975. (21) Nehl, C. L.; Liao, H. W.; Hafner, J. H. Optical properties of starshaped gold nanoparticles. Nano Lett. 2006, 6, 683−688. (22) Kwon, K.; Lee, K. Y.; Lee, Y. W.; Kim, M.; Heo, J.; Ahn, S. J.; Han, S. W. Controlled synthesis of icosahedral gold nanoparticles and their surface-enhanced raman scattering property. J. Phys. Chem. C 2007, 111, 1161−1165. (23) Kim, S. H.; Shim, J. W.; Yang, S. M. Microfluidic multicolor encoding of microspheres with nanoscopic surface complexity for multiplex immunoassays. Angew. Chem., Int. Ed. 2011, 50, 1171−1174. (24) Axelsson, I. Characterization of proteins and other macromolecules by agarose gel chromatography. J. Chromatogr. A 1978, 152, 21−32. (25) Brewer, S. H.; Glomm, W. R.; Johnson, M. C.; Knag, M. K.; Franzen, S. Probing BSA binding to citrate-coated gold nanoparticles and surfaces. Langmuir 2005, 21, 9303−9307. (26) Silin, V.; Weetall, H.; Vanderah, D. J. SPR studies of the nonspecific adsorption kinetics of human lgG and BSA on gold surfaces modified by self-assembled monolayers (SAMs). J. Colloid Interface Sci. 1997, 185, 94−103. (27) Indrasekara, A. S. D. S.; Meyers, S.; Shubeita, S.; Feldman, L. C.; Gustafsson, T.; Fabris, L. Gold nanostar substrates for SERS-based chemical sensing in the femtomolar regime. Nanoscale 2014, 6, 8891− 8899. (28) Zijlstra, P.; Paulo, P. M. R.; Orrit, M. Optical detection of single non-absorbing molecules using the surface plasmon resonance of a gold nanorod. Nat. Nanotechnol. 2012, 7, 379−382. (29) Chen, J.; Saeki, F.; Wiley, B. J.; Cang, H.; Cobb, M. J.; Li, Z.-Y.; Au, L.; Zhang, H.; Kimmey, M. B.; Li, X.; Xia, Y. Gold nanocages: Bioconjugation and their potential use as optical imaging contrast agents. Nano Lett. 2005, 5, 473−477. (30) Xie, J.; Zhang, Q.; Lee, J. Y.; Wang, D. I. C. The synthesis of SERS-active gold nanoflower tags for in vivo applications. ACS Nano 2008, 2, 2473−2480. (31) Utada, A. S.; Lorenceau, E.; Link, D. R.; Kaplan, P. D.; Stone, H. A.; Weitz, D. A. Monodisperse double emulsions generated from a microcapillary device. Science 2005, 308, 537−541. (32) Frens, G. Controlled nucleation for the regulation of the particle size in monodisperse gold suspensions. Nature, Phys. Sci. 1973, 241, 20−22.

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DOI: 10.1021/acs.chemmater.6b00115 Chem. Mater. XXXX, XXX, XXX−XXX