Partitioning and Assembly of Metal Particles and Their Bioconjugates

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Partitioning and Assembly of Metal Particles and Their Bioconjugates in Aqueous Two-Phase Systems Marcus R. Helfrich, Mahnaz El-Kouedi,† Mark R. Etherton,‡ and Christine D. Keating* Department of Chemistry, The Pennsylvania State University, University Park, Pennsylvania 16802 Received May 7, 2005. In Final Form: June 20, 2005 The behavior of metal nanospheres and nanowires and their bioconjugates in aqueous two-phase systems (ATPS) is described. The ATPS used in this work comprised poly(ethylene glycol) (PEG), dextran, and water or aqueous buffer. Au and Ag nanospheres less than 100 nm in diameter partition between the PEG-rich and dextran-rich phases on the basis of their surface chemistry and can be separated on this basis. Larger Au nanospheres and wires accumulate at the interface between the two aqueous phases. The influence of polymer molecular weight and concentration on interfacial assembly of Au wires is described. DNA-derivatized nanowires at the aqueous/aqueous interface retain the ability to selectively bind to fluorescent complementary DNA. In addition, Au nanoparticles have been bound to Au wires via selective DNA hybridization at the ATPS interface. Transmission electron microscopy and thermal denaturation experiments confirm that DNA-driven assembly is responsible for the formation of the nanosphere/wire assemblies. These results demonstrate the biocompatibility of the two-phase interface and point to future use as scaffolding in biorecognition-driven assembly.

Introduction When two chemically dissimilar polymers are mixed in sufficiently high weight percent in water, distinct aqueous phases occur, each enriched in one of the polymers.1 Unlike organic/aqueous biphasic systems, aqueous two-phase systems (ATPS) are biocompatible and can be prepared in the presence of high ionic strength buffers. As a result, these systems have been used for the separation of a variety of different types of biomolecules, including proteins, nucleic acids, and cellular materials.1-4 The partitioning of biomolecules within an ATPS can be influenced by several factors, including the size of the polymers, the size of the material being partitioned, the polymer composition within the individual phases, and the enthalpalic interactions between the polymers and the molecule of interest.1-5 The presence and identity of additional solutes (e.g., salts) can also impact partitioning, as can the pH dependence of charged polymers or biomacromolecules within the ATPS. In general, larger macromolecules tend to partition more extensively due to increased contact with the solution. Larger objects, such as organelles, can partition between one of the phases and the aqueous/aqueous interface.1 * To whom correspondence should be addressed. E-mail: [email protected]. † Present address: Department of Chemistry, University of North Carolina, Charlotte, NC 28223. ‡ Present address: University of Texas Southwestern, Dallas, TX 75390. (1) Albertsson, P. A. Partition of Cell Particles and Macromolecules; 2nd ed.; Wiley-Interscience: New York, 1971. (2) Aqueous Two-Phase Systems: Methods and Protocols; Hatti-Kaul, R. Ed.; Methods in Biotechnology; Humana Press: Totowa, NJ, 2000; Vol. 11. (3) Aqueous Two-Phase Systems; Walter, H., Johansson, G., Eds.; Methods in Enzymology; Academic Press Inc.: San Diego, 1994; Vol. 228. (4) Partitioning in Aqueous Two-Phase Systems: Theory, Methods, Uses, and Applications to Biotechnology; Walter, H., Brooks, D. E., Fisher, D., Eds.; Academic Press: Orlando, 1985. (5) Johansson, H. O.; Karlstro¨m, G.; Tjerneld, F.; Haynes, C. A. J. Chromatogr., B 1998, 711, 3-17.

While the partitioning of biomolecules within ATPS has been examined extensively, little is known about the behavior of inorganic particles in these types of systems. The partitioning of latex and TiO2 particles was found to depend on both particle size and surface chemistry.6 Although no quantitative results were given, the authors reported excellent partitioning, “for all practical purposes completely into one of the phases.”6 More recently, the role of surface chemistry (surface wetability) on the particle-induced phase separation of poly(ethylene oxide)/ dextran solutions near the cloud-point has been investigated.7 In both of these studies, the inorganic particles partitioned into one of the two phases, with the preferential wetting of particle surface by one of the polymers directing partitioning.4 When inorganic nanoparticles are added to organic/ aqueous biphasic systems, particle assembly at the liquid/ liquid interface is often observed.8-14 There have been reports of millimeter-, micrometer-, and nanoscale particles at such interfaces, and a variety of interesting structures have been formed. In recent work by Wang, Russell, and co-workers, virus particles were assembled at an oil/water interface.9 These virus particles are quite stable and retained their structural integrity at the (6) Baxter, S. M.; Sperry, P. R.; Fu, Z. Langmuir 1997, 13, 39483952. (7) Olsson, M.; Joabsson, F.; Piculell, L. Langmuir 2005, 21, 15601567. (8) Dinsmore, A. D.; Hsu, M. F.; Nikolaides, M. G.; Marquez, M.; Bausch, A. R.; Weitz, D. A. Science 2002, 298, 1006-1009. (9) Russell, J. T.; Lin, Y.; Boker, A.; Su, Long, Carl, P.; Zettl, H.; He, J.; Sill, K.; Tangirala, R.; Emrick, T.; Littrell, K.; Thiyagarajan, P.; Cookson, D.; Fery, A.; Wang, Q.; Russell, T. P. Angew. Chem., Int. Ed. 2005, 44, 2420-2426. (10) Dai, L. L.; Sharma, R.; Wu, C.-y. Langmuir 2005, 21, 26412643. (11) Kumar, A.; Mandal, S.; Mathew, S. P.; Selvakannan, P. R.; Mandale, A. B.; Chaudhari, R. V.; Sastry, M. Langmuir 2002, 18, 64786483. (12) Zheng, L.; Li, J. J. Phys. Chem. B 2005, 109, 1108-1112. (13) Choi, I. S.; Weck, M.; Xu, B.; Jeon, N. L.; Whitesides, G. M. Langmuir 2000, 16, 2997-2999. (14) Lin, Y.; Boker, A.; Skaff, H.; Cookson, D.; Dinsmore, A. D.; Emrick, T.; Russell, T. P. Langmuir 2005, 21, 191-194.

10.1021/la051220z CCC: $30.25 © 2005 American Chemical Society Published on Web 07/28/2005

Metal Particle/Bioconjugate Partitioning, Assembly

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Table 1. DNA Sequences Used in This Work sequence description

sequence number

sequence (5′ to 3′)

biotinylated sequence 1 biotinylated sequence 2 Au conjugate sequence 1 complementary linking strand noncomplementary linking strand complementary fluorescent strand noncomplementary fluorescent strand

I II III IV V VI VII

(biotin-TEG)-T6 CGC ATT CAG GAT (biotin-TEG)-T18 CGC ATT CAG GAT TCT CAA CTC GTA-(C3H6-SH) TAC GAG TTG AGA ATC CTG AAT GCG GCG TAA GTC CTA ACA GAT CTC CAT (6-FAM)-ATC CTG AAT GCG (6-FAM)-GCT ACG GCC TAG

interface. Despite these promising results, it is not generally possible to conduct biorecognition-driven assembly reactions at the organic/aqueous interface due to unfavorable interactions between the biomolecules and the organic phase. ATPS are a potentially attractive alternative to organic/aqueous interfaces due to their biocompatibity and low, tunable, interfacial tensions. Biorecognition-driven nanoparticle assembly is interesting from a materials perspective and has application in biosensing;15-19 it has not been explored in ATPS. Polymeric solutes can act as volume excluders as well as interact with the particles and solvent, altering assembly behavior. Recently, we reported the effect of polymer molecular weight and concentration on DNA-directed nanoparticle assembly in PEG and dextran solutions.20 We found significant increases in the thermodynamic stability of DNA/Au nanoparticle aggregates in PEG solutions and a smaller increase for dextran solutions. Herein, we report the behavior of Au and Ag nanospheres and Au nanowires in several PEG/dextran ATPS compositions. Nanospheres smaller than 80 nm in diameter partitioned between the PEG-rich and dextran-rich phases of the ATPS, whereas larger particles accumulated at the aqueous/aqueous interface. An increase in the molecular weight or concentration of the polymers within the ATPS results in a subsequent reduction in the mobility of particles trapped at the interface. We demonstrate that the interface between the polymer solutions is biocompatible and that DNA hybridization can be conducted on the surface of the trapped nanoparticles. Using this approach, the DNA-directed assembly of 14-nm gold nanoparticles to the surface of 3-µm × 320-nm Au nanowires is performed. Experimental Section Materials. All chemicals were used as received except where noted. Gold (Orotemp 24) and silver (Silver 1025 and Silver Cyless) plating solutions were purchased from Technic, Inc. HAuCl4‚3H2O was received from Acros Organics. Neutravidin protein, 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC), and N-hydroxysulfosuccinimide (sulfo-NHS) were purchased from Pierce. All oligonucleotides used in this work are listed in Table 1, were synthesized using an Applied Biosystems Expedite 8909 Oligonucleotide synthesizer, and were purified using established protocols with NAP-5 columns (Amersham Pharmacia). The poly(ethylene glycol)s used in this work include PEG MW 8000 (Sigma) and PEG 4600 (Sigma) and PEG 12 000 MW (Fluka). Dextran 10 000 MW, dextran 100 000 MW, dextran 505 000 MW, mercaptoethanesulfonic acid (MESA), 11-mercaptoundecanoic acid (MUA), mercaptoethylamine (MEA), trisodium citrate dihydrate, and DL-dithiothreitol (DTT) were purchased from Sigma-Aldrich. (3-Aminopropyl)trimethoxysilane (APTMS), tet(15) Rosi, N. L.; Mirkin, C. A. Chem. Rev. 2005, 105, 1547-1562. (16) Mirkin, C. A. Inorg. Chem. 2000, 39, 2258-2272. (17) Mann, S.; Shenton, W.; Li, M.; Connolly, S.; Fitzmaurice, D. Adv. Mater. 2000, 12, 147-150. (18) Niemeyer, C. M. Angew. Chem., Int. Ed. 2001, 40, 4128-4158. (19) Alivisatos, P. Nat. Biotechnol. 2004, 22, 47-52. (20) Goodrich, G. P.; Helfrich, M. R.; Overberg, J. J.; Keating, C. D. Langmuir, 2004, 20, 10246-10251.

raethyl orthosilicate (TEOS), and carboxyethylsilanetriol (CEST) were received from Gelest. 11-Amino-1-undecanethiol hydrochloride was purchased from Dojindo Laboratories (Japan). All colloidal gold solutions with diameters larger than 14 nm were purchased from Ted Pella, Inc. All H2O used in these experiments was distilled and purified to a resistance of 18.2 MΩ using a Barnstead Nanopure system. Instrumentation. Ultraviolet-Visible Spectroscopy. All UVvis spectra were obtained on a Hewlett-Packard 8453A diodearray spectrophotometer using ChemStation UV-vis software. The melting temperature, Tm, for the bioconjugate aggregates was determined using the thermal denaturation mode of the software and a Hewlett-Packard 89090A Peltier temperature control unit, using a 160-µL quartz cuvette with a 1-cm path length (Starna Cells). Transmission Electron Microscopy. A JEOL 1200 EXII TEM was used to obtain TEM micrographs. Images were taken using a high-resolution Tietz F224 CCD camera using the associated software package. Viscosity. A TA Instruments CSL2 100 Rheometer was used to measure the viscosity of the polymers. Interfacial Tension. A home-built tensiometer was assembled on the basis of the previous design of Vonnegaut.21 The rotating cell was a borosilicate tube (10.16 cm in length, 0.394 cm i.d.) and was driven by an Ika Laboritechnik RW20DZM rotary mixer (240-2000 rpm) with integrated speed display. One end of the tube was fitted with a Teflon stopper that attached into a precision metal bearing with three Teflon screws to couple the tube to the mixer. The other end of the tube was fed through a second precision metal bearing and was sealed with a Teflon stopper with a fine bore through the middle. The top phase was introduced through this hole into the sample chamber and was sealed with a thin Teflon plug before rotation was started. Interfacial tension measurements were conducted using the procedure outlined by Princen et al.22 The length of the droplets during spinning was measured using a Kodak MDS 100 digital camera with provided software and were analyzed postcapture using Adobe Photoshop (version 7). Optical Microscopy. Optical images of nanowires at the ATPS interface were collected using a Nikon Eclipse 800 upright microscope equipped with a Hamamatsu ORCA 12-bit, highresolution, cooled, CCD camera with 6.7-µm pixel (1024 × 1024). Nanowires were imaged using a 60× dipping lens (NA ) 1.00) with a 100-W Hg lamp for illumination. Reflected light images were collected using a Nikon brightfield reflectance cube fitted with a 50:50 beam splitter. To determine nanowire mobility at the interface, a sequence of 20 sequential images was collected using the minimum exposure time for the camera (850 ms). These sequences were then converted to individual images (TIFF) and transferred to Adobe Photoshop, where the angle of the nanowire was measured as a function of time. The average angular rotation listed in Table 1 in the Supporting Information is the average of the total range of angles covered by the nanowires over a period of ∼20 s for each ATPS. Fluorescence Microscopy on Nanowires. DNA conjugates were performed using a Nikon TE-300 inverted microscope fitted with a Photometrics CoolSnap HQ cooled, CCD camera and a 175-W Xe arc lamp with filter wheel (Sutter Instruments). All microscope images were collected using Image-Pro Plus software (version 4.5). (21) Vonnegut, B. Rev. Sci. Instrum. 1942, 13, 6-9. (22) Princen, H. M.; Zia, I. Y. Z.; Mason, S. G. J. Colloid Int. Sci. 1967, 23, 99-107.

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Polymer Concentrations. Polymer concentrations were determined using a Leica Auto-Abbe refractometer with sample stage cooled to 25 °C using a VWR brand water bath. Dextran concentrations were measured using a Perkin-Elmer model 343 polarimeter with a 350-µL sample cell at room temperature. The composition of the top and bottom polymer phases in a given ATPS was determined using a combination of polarimetry and refractometry, as previously described.2 Nanowire Synthesis and Functionalization. Nanowires ∼320 nm in diameter and ∼3-µm in length were grown galvanostatically from commercially available aluminum oxide membranes (Whatman) using previously described methods.23,24 Nanowires were derivatized with a variety of thiol-bearing organic groups. To accomplish this, the wires were pelleted using a Biofuge Pico centrifuge (Heraeus) at ∼3400g for 4 min and resuspended in 1 mL of a 0.1 M ethanolic solution of the thiol of interest. The wires were vigorously shaken in this solution for 1 h, after which they were washed in ethanol four times and then in H2O four times before resuspending to a final volume of 1 mL in H2O. DNA/nanowire conjugates were prepared by first functionalizing the nanowires with Neutravidin. A 500-µL aliquot of a 0.25 mg/mL Neutravidin solution was reacted with 1 mL of wires in H2O for 2 h before excess protein was removed through centrifugation. A 2-fold excess of biotinylated oligonucleotide was then added, and the wires were vortexed a minimum of 4 h.25 Excess oligonucleotide was removed by centrifugation, and removal of the supernatant, a minimum of three times. Wires were then resuspended in 1 mL of the appropriate phosphatebuffered saline solution for the given experiment. Glass-coated nanowires were prepared using a modification of the procedure first described by Yin et al.26 To glass-coat wires, 300 µL of a 1-mL wire solution (in ethanol) was added to 500 µL of ethanol along with 160 µL of H2O and 15 µL of concentrated ammonium hydroxide. To this, 40 µL of freshly distilled TEOS was added, and the solution was immediately sealed and sonicated for 45 min. The wires were pelleted (∼6100 g for 2 min); washed six times with ethanol; and finally, resuspended to a final volume of 300 µL in ethanol. To functionalize the surface of these glass-coated wires, 1 mL of glass-coated nanowires was pelleted and resuspended in 1 mL of the desired silane (10% solution in ethanol). The sample was vortexed vigorously for 1 h. Excess silane was removed via centrifugation, and removal of the supernate, a minimum of four times. Wires were brought to a final volume of 1 mL in H2O. Nanoparticle Synthesis. Colloidal gold (14-nm diameter) was prepared through the reduction of HAuCl4 with sodium citrate, as previously described.27 Average particle diameters were 14.2 ( 1.3 nm, determined by TEM using NIH Image software. The same batch of 14-nm gold colloid was used for all of the experiments described, and the concentration of the nanoparticles was determined to be 1.58 × 10-8 M.20 Colloidal Ag nanoparticles were prepared by EDTA reduction as previously described.28,29 These particles are generally spherical but polydisperse (diameters ∼14 ( 8 nm).29 DNA/Colloidal Gold Conjugates. Au/DNA conjugates were prepared by assembly of thiolated oligonucleotides onto Au nanospheres using a slightly modified version of the procedure described by Mirkin and co-workers.30,31 Conjugates were spun (23) Nicewarner-Pen˜a, S. R.; Freeman, R. G.; Reiss, B. D.; He, L.; Pen˜a, D. J.; Walton, I. D.; Cromer, R.; Keating, C. D.; Natan, M. J. Science, 2001, 294, 137-141. (24) Nicewarner-Pen˜a, S. R.; Carado, A. J.; Shale, K. E.; Keating, C. D. J. Phys. Chem. B 2003, 107, 7360-7367. (25) The amount of biotinylated DNA to be added to a sample was determined on the basis of a surface coverage of 7.7 × 1011 oligo/cm2. This surface coverage was determined by removal of fluorescently labeled biotinylated DNA strands from the surface of a known number of 3-µm × 320-nm wires. (26) Yin, Y.; Lu, Y. Xia, Y. Nano Lett. 2002, 2, 427-430. (27) Grabar, K. C.; Freeman, R. G.; Hommer, M. B.; Natan, M. J. Anal. Chem. 1995, 67, 735-743. (28) Lee, N.-S.; Sheng, R.-s.; Morris, M. D.; Schopfer, L. M. J. Am. Chem. Soc. 1986, 108, 6179-6183. (29) Keating, C. D.; Kovaleski, K. M.; Natan, M. J. J. Phys. Chem. B 1998, 102, 9414-9425. (30) Mirkin, C. A.; Letsinger, R. L.; Mucic, R. C.; Storhoff, J. C. Nature 1996, 382, 607-609.

Helfrich et al. down twice at ∼10 930g for 35 min, the supernatant was removed, and the soft pellet containing the Au particles was resuspended in 0.3 M NaCl/10 mM phosphate buffer pH 7. Conjugate concentration was determined by UV-vis using the experimentally determined extinction coefficient (1.94 × 108 M-1 cm-1) for the Au particles (based on Au concentration from atomic absorption spectroscopy and particle size from TEM). Preparation of Aqueous Two-Phase Systems. To determine the compositions of the polymer phases that would be used to prepare the ATPS in these experiments, the phase diagram for each polymer combination was first defined through cloudpoint titration.2 The phase behavior of PEG/dextran ATPS is somewhat sensitive to changes in temperature, particularly at low polymer concentrations.32,33 The concentrations of PEG and dextran used in this work were chosen such that two phases would be maintained, even at elevated temperatures often used in DNA hybridization experiments. To prepare an ATPS, the two polymers were weighed into a glass vial along with the appropriate amount of buffer to bring the resulting ATPS to the polymer concentration desired. The ATPS was then vigorously stirred until the polymers fully dissolved. The stir bar was removed, and the polymer solutions were allowed to phase-separate overnight. The use of phosphatebuffered saline solutions did not significantly alter the position of the binodal curve when compared to the phase diagram in H2O alone. The composition of the top and bottom polymer phases in a given ATPS was determined using a combination of polarimetry and refractometry as previously described.2 Polymer solutions were prepared as described above and allowed to phase-separate for a period of 2 days before removal of the two phases for analysis. For polarimetry, samples were first diluted 1:4 in buffer prior to evaluation to reduce their viscosity and aid in loading the sample cell. Deposition of Nanowires at the ATPS Interface. For a typical assembly experiment, 5.0 mL of the ATPS solution was transferred to a 35-mm plastic Petri dish (VWR Scientific) and allowed to phase-separate overnight. To this interface, 100 µL of nanowires was added through the top phase and allowed to settle via gravity overnight. The microscope objective (a 60× dipping lens) was slowly lowered through the top phase to minimize influencing wire packing at the interface. Images were collected following a 5-10-min equilibration time. Interfacial Hybridization Experiments. During preparation of the ATPS, 2 µL of fluorescently labeled DNA (DNA sequence VI or VII) was added, and the solution was allowed to phase-separate for 16 h in the dark. To this, 125 µL of gold nanowires functionalized with Neutravidin and DNA sequence I was pipetted in through the top phase and allowed to settle to the interface and hybridize in the dark for 36 h. Following this period, the wires were removed from the interface via pipet, and excess DNA was removed with successive washes of hybridization buffer. Samples were kept in the dark prior to observation. Both the complementary and noncomplementary DNA oligonucleotides were labeled with 6-carboxyfluorescein (6-FAM), and fluorescence images were acquired using a Nikon FITC cube (λex ) 465-495 nm, dichroic @ 505 nm, and λem ) 515-555 nm). After capturing the fluorescence image, the reflectance image of the wires was captured using a 520-nm, short-pass, excitation filter in combination with a Nikon brightfield reflectance cube fitted with a 50:50 beam splitter. Nanowire-Colloid Assembly Experiments. Au/DNA conjugates functionalized with sequence III were dispersed in the initial ATPS during formation. This was accomplished by substituting 1 mL of the hybridization buffer for 1 mL of the bioconjugates. Upon phase separation, the bioconjugates were freely dispersed in the lower dextran phase, a result of the singlestranded DNA on the particle exterior. To this ATPS, 100 µL of DNA-coated nanowires (DNA sequence II) was added through the top phase and allowed to settle at the interface overnight. (31) Nicewarner-Pen˜a, S. R.; Raina, S.; Goodrich, G. P.; Fedoroff, N. V.; Keating, C. D. J. Am. Chem. Soc. 2002, 24, 7314-7323. (32) Long, M. S.; Jones, C.; Helfrich, M. R.; Mangeney-Slavin, L. K.; Keating, C. D. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 5920-5925. (33) Helfrich, M. R.; Mangeney-Slavin, L. K.; Long, M. S.; Djoko, K. Y.; Keating, C. D. J. Am. Chem. Soc. 2002, 124, 13374-13375.

Metal Particle/Bioconjugate Partitioning, Assembly

Figure 1. Nanosphere partitioning. Digital photograph (A) and optical extinction spectra (B) for Au nanospheres and digital photograph (C) and optical extinction spectra (D) for Ag nanospheres in an ATPS. ATPS composition was 9% PEG 8 kDa and 9% dextran 10 kDa. In both B and D, solid lines indicate spectra for the upper, PEG-rich phase, and dotted lines indicate spectra for the lower, dextran-rich phase. The particles were allowed to hybridize for a period of 48-96 h prior to the wires’ being removed from the interface via pipet. Excess Au/DNA conjugates were removed by repeated centrifugation (∼3400g for 4 min) and subsequent washing with hybridization buffer (a minimum of five times). Nanowire/ bioconjugate assemblies were imaged by TEM using 200-mesh Cu grids (Ted Pella) backed with Formvar. To reduce solution evaporation during the thermal denaturation, a small amount of mineral oil was added to the top of the solution, and the cuvette was capped. All plots were taken using 1 °C intervals with a hold time of 2 min (temperature range ) 40-80 °C). The Tm of the system was determined from the first derivative of the resulting melt curve.

Results and Discussion In this section, we first describe the partitioning behavior of metal nanospheres in a PEG/dextran ATPS as a function of surface chemistry and particle size. We then describe assembly of nanowires to the aqueous/ aqueous interface and factors that impact the motion of wires at this interface. Finally, we demonstrate biorecognition-driven nanoparticle assembly at the ATPS interface. Nanosphere Partitioning between the Phases. Addition of 14-nm colloidal Au nanospheres to a PEG/ dextran ATPS leads to preferential accumulation of the nanospheres in the upper, PEG-rich phase (Figure 1 A and B). Partitioning is very efficient; the top phase is an intense red/orange color due to the Au nanospheres, whereas the bottom phase appears colorless. The strong absorbance at 520 nm for the Au nanospheres can be seen in the extinction spectrum for the PEG-rich phase, whereas the dextran-rich phase is essentially colorless (note that extinction arises from both plasmon absorbance and

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scattering; for these 12-nm-diameter particles, absorbance dominates). We estimate a partitioning coefficient, K, where K is equal to the concentration of Au nanospheres in the top phase divided by that in the lower phase,1 of g39 for the Au nanospheres in this ATPS (this is a lower limit because we did a simple division of the extinction values at 520 nm for the two spectra shown in Figure 1B; a portion of the extinction in the dextan-rich phase is due to scattering from the polymer solution itself). In contrast, addition of Ag nanospheres to an ATPS of the same composition results in Ag nanosphere accumulation in the bottom, dextran-rich phase (Figure 1C and D). The distinctive yellow color and absorbance band, ∼400 nm for the Ag nanospheres, is observed almost exclusively in the dextran-rich phase. Since a small absorbance feature due to Ag particles can be seen in the PEG-rich phase, for this sample, K ) 0.0378, indicating at least a 26-fold greater concentration in the dextran-rich phase. Interestingly, the position of the absorbance maximum, which occurs at 404 nm in the dextran-rich phase, is shifted to ∼420 nm in the PEG-rich phase. This red shift could arise from differences in adsorbed layers or in bulk refractive index between the two phases, from preferential accumulation of particles having different size or shape (and therefore different optical properties) in the two phases, or from both. Since we have not observed different phase preferences for different sizes of Au spheres, we interpret this red shift as the result of microscopic (i.e., adsorbed) or bulk refractive index differences between the two phases.34 Nanosphere partitioning, like molecule partitioning, is understood to result from several factors, including chemical interactions between the polymers and the particle surface, as well as interactions with the different solvent environments.35 The two phases of a PEG/dextran ATPS display differences in polarity, water mobility, and proton activity.35 Preferential adsorption of one polymer onto the particle surface reduces the surface free energy and can be used to control the partitioning of the material.4,6,7 The different phase preferences observed for the Au and Ag nanospheres can be understood on the basis of their surface chemistries. The Ag nanospheres were prepared by EDTA reduction of AgNO3, whereas the Au nanospheres were formed by citrate reduction of [AuCl4]-. Although both types of particles are negatively charged, they have different adsorbates on their surfaces. This can be illustrated by the greatly reduced partitioning of Ag particles after exposure to the citrate-rich solution in which the Au spheres are suspended, either by mixing the two particle types together (Figure 2A and B) or by addition of the solution from the Au nanosphere suspension after removal of the Au nanospheres by centrifugation (not shown). Partitioning for the adsorbates themselves does not parallel the nanosphere partitioning; e.g., trisodium citrate has been shown to partition weakly into the dextran-rich phase of PEG 4 kDa/dextran 500 kDa ATPS.1 This is unsurprising, since the orientation of citrate on the particles (i.e., bound via carboxylate groups)28 will result in different interactions with the ATPS. In addition, surface adsorbates may include not only the citrate or EDTA molecules but also oxidation products from the nanoparticle synthesis reaction (e.g., homocitrate), Cl- in the case of Au particles, as well as the PEG or dextran (34) (a) Mulvaney, P. Langmuir 1996, 12, 788-800. (b) Yonzon, C. R.; Jeoung, E.; Zou, S.; Schatz, G. C.; Mrksich, M.; Van Duyne, R. P. J. Am. Chem. Soc. 2004, 39, 12669-12676. (35) Zaslavsky, B. Y. Aqueous Two-Phase Partitioning; Marcel Dekker: New York, 1994.

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Figure 3. Digital photograph showing the position of the gold nanoparticles in the top phase of a PEG 8K/dextran 10K ATPS after 96 h. The dark reddish color in the bottom phase of the 100-nm sample as well as the gold ring in the bottom of the 250-nm sample are the result of nanoparticles sticking to the walls of the test tube during the initial phase separation.

Figure 2. Digital photograph (A) and optical extinction spectra (B) for Au and Ag nanospheres in an ATPS. Digital photograph (C) and optical extinction spectra (D) for DNA/Au nanosphere bioconjugates (DNA sequence III) in an ATPS. ATPS composition was 9% PEG 8 kDa and 9% dextran 10 kDa. Solid lines indicate spectra for the upper, PEG-rich phase, and dotted lines indicate spectra for the lower, dextran-rich phase.

polymers. The presence of ions in solution can also impact partitioning, an effect that has been used to modulate protein separations in ATPS.1 We note that the position of the Ag surface plasmon absorbance is blue-shifted in the PEG-rich phase, as compared with the dextran-rich phase in Figure 2B, indicating the different chemical environments of the particles in the two phases.34 Differences in particle surface chemistry can be used to control the location of materials in the ATPS. Figure 2C and D shows the partitioning of DNA/Au nanosphere bioconjugates, in which each 14-nm Au sphere has been coated with many thiolated oligonucleotides, in a 9% PEG 8 kDa/9% dextran 10 kDa ATPS. The partitioning of the DNA/Au conjugates to the lower, dextran-rich phase is consistent with the behavior of free, single-stranded DNA in ATPS prepared in high ionic strength solutions.1,36,37 The derivatization of colloidal gold particles with biomolecules offers a new route for controlling the partitioning behavior of these materials within an ATPS. In some cases, separations can be performed. Although the partitioning of the unfunctionalized Ag nanospheres used here was less impressive after these particles were exposed to the citrate solution, the partial separation can be seen in Figure 2A. With modified surface chemistry, it should be possible to get a more impressive separation. Nanosphere Partitioning between the Top Phase and the Aqueous/Aqueous Interface. Albertsson has suggested that partitioning between the two phases of an ATPS can be increased by increasing the size of the molecule or particle to be partitioned, as this will increase the area of surface interactions.1 For metal nanoparticles, (36) Albertsson, P. A. Biochim. Biophys. Acta 1965, 103, 1-12. (37) Pettijohn, D. E. Eur. J. Biochem. 1967, 3, 25-32.

this trend cannot be expected to continue indefinitely, due to gravity and the high density of the particles (19.3 g/cm3 for Au). To determine the influence of particle size on the partitioning of citrate-stabilized Au nanoparticles, monodisperse samples of 12-, 30-, 50-, 80-, 100-, and 250-nmdiameter Au spheres were added to a 9% PEG 8 kDa/9% dextran 10 kDa ATPS prior to phase separation. Upon phase separation, the citrate-reduced Au particles of all sizes had accumulated in the upper, PEG-rich phase. Very large or dense particles can accumulate at the aqueous/ aqueous interface. In the experiments described above, ATPS phase separation was complete within 12 h. Within the first 56 h, the 250- and 100-nm diameter colloidal Au particles began to settle to the interface (Figure 3). These particles remain at the interface indefinitely, indicating that the interfacial tension is sufficient to overcome the force of gravity on the particles. After 96 h, significant partitioning to the interface was not observed for the nanoparticles with diameters smaller than 100-nm. Nanowire Assembly at the Aqueous/Aqueous Interface. When Au nanowires (70, 220, and 320 nm in diameter and 3 µm in length) were mixed into an ATPS, rapid sedimentation resulted in less than one-half of the wires collecting at the interface, with the remainder settling to the bottom of the Petri dish (i.e., any nanowire initially located below the forming aqueous/aqueous interface ended up on the bottom of the dish, while those above the forming interface sedimented to the interface). This had not been observed for the spheres described above, due to their initial accumulation in the PEG phase and their slower sedimentation as compared with wires. To increase the efficiency of wire capture at the interface, nanowires were added through the upper PEG phase without mixing the phases. This approach results in essentially all of the wires residing at the interface within 12 h, Figure 4. The rate at which particles settle to the interface can be increased by centrifugation or by reducing the depth of the top phase, for example, by moving to a shallow container. The interface between the polymer phases of an ATPS is characterized by a low interfacial tension, typically on the order of 0.0001-0.1 dyn/cm.1 Altering the polymer molecular weights or concentrations in an ATPS provides tunability in interfacial tension and could be expected to impact particle assembly at the interface. To determine how these changes would influence the ability to collect wires at the interface, the interfacial tensions for each ATPS were determined using spinning drop tensiometry.1,21,22 The interfacial tensions for various PEG/dextran

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Figure 4. Photograph (left) and microscope image (right) of DNA-derivatized Au nanowires at the interface of an ATPS: (a) aqueous/air interface and (b) aqueous/aqueous interface. Nanowires are 320 nm in diameter and 2 µm in length and are only present at the interface (b). Table 2. Effect of ATPS Composition on Interfacial Tension aqueous two-phase system

polymer concns (% PEG - % dex)

interfacial tension (dyn/cm)

PEG 4.6K/dex 100K

7-7 9-9 11-11 9-9 11-11 13-13 7-7 9-9 11-11 7-7 9-9 11-11 7-7 9-9 11-11

0.013 ( 0.002 0.073 ( 0.003 0.198 ( 0.023 0.030 ( 0.002 0.142 ( 0.045 0.309 ( 0.045 0.053 ( 0.002 0.163 ( 0.033 0.314 ( 0.017 0.112 ( 0.021 0.207 ( 0.019 0.345 ( 0.061 0.077 ( 0.005 0.170 ( 0.011 0.254 ( 0.009

PEG 8K/dex 10K PEG 8K/dex 100K PEG 8K/dex 505K PEG 12K/dex 100K

ATPS compositions ranged from 0.013 to 0.35 dyn/cm and are shown in Table 2. As expected, increased polymer concentration and increased molecular weights led to increased interfacial tension. Au nanowires (320 nm × 3 µm) collected at the interface of any of these ATPS (the force of gravity on a single Au wire is on the order of 5 × 10-9 dyne). For comparison, typical interfacial tensions for organic/aqueous interfaces are generally in the 1-20 dyn/cm range.1 To determine what influence surface chemistry has on the placement of metal nanowires at the polymer interface, wires were deriviatized with functional groups prior to their addition. The gold surface of the nanowire was coated with several different alkanethiols to yield both a negative surface charge (using MESA and MUA) as well as a positive charge (using 11-amino-1-undecanethiol and MEA). Nanowires were also encapsulated in a thin (∼50 nm thick) layer of SiO2 prior to similar surface derivatization with the organosilanes CEST (terminal carboxylate) and APTMS (terminal amine). The incorporation of charged terminal functionalities on the surface of the nanowires did not influence wire placement at the polymer-polymer interface. Nanowire derivatization with biotin, Neutravidin, biotinylated ss-DNA (18-56 base pairs in length), human IgG and anti-human IgG also yielded similar results. These data indicate that the nanowires can be derivatized with a variety of different chemistries without compromising their placement at the interface. The phase behavior of aqueous two-phase systems can be altered by changes in the pH, temperature, or salt

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concentration.1 This effect is most noticeable for polymer concentrations in the region on the phase diagram where the ATPS changes from one phase to two phases. At these compositions, the interfacial tension is quite small, resulting in a poorly defined interface. The weight percent of the polymers used in these experiments was selected to ensure that a reproducible interface was formed that was insensitive to experimental variations in temperature and salt concentration. As a result, the two-phase systems studied here are far removed from the region where the ATPS transitions between one and two phases, with high concentrations (>12 wt/wt %) of each polymer in its respective phase. This, combined with the high molecular weights of the polymers, results in the formation of two viscous polymer phases. Since controlled nanowire assembly at the ATPS interface will require not only that the interfacial tension is sufficient to overcome gravity, but also that the wires retain some mobility at the interface, we were interested in the effect of the ATPS composition on wire mobility. Changing the ATPS composition impacts not only interfacial tension, but also viscosity. Figure 5 shows the relationship between mobility and viscosity for MESAcoated nanowires for the five ATPS shown in Table 2. Increases in the polymer concentration within a specific ATPS resulted in a corresponding decrease in mobility of the wires at the interface. By varying the molecular weights of the dextrans used (10, 100, and 505 kDa) versus those of the PEGs used (4.6, 8, and 12 kDa), the viscosity of the phases can be modulated (note that interfacial tension is also changed in these experiments). The effect on the two phases is not the same, however. At constant PEG molecular weight, increasing the dextran molecular weight resulted in lower viscosity in the top phase and much higher viscosity in the bottom phase, as compared with lower molecular weight dextrans (Figure 5A and B). This result is consistent with the expected greater partioning of the polymers between the phases as molecular weight is increased; therefore, the increased viscosity associated with this high-molecular-weight polymer increases the viscosity of the lower (dextran-rich) phase, while the reduced partitioning of dextran into the upper (PEG-rich) phase reduces its viscosity. The same trends can be seen when the dextran concentration is held constant and the PEG molecular weight is varied (Figure 5C and D). Nanowire mobility at the interface is significantly impacted by changes in the ATPS composition. We report mobilities as the maximum angle of rotation over a 20-s video collected of the wires at the interface; rotation was selected over lateral displacement to reduce the effect of sample drift on the measurements. Figure 5 shows that mobility is proportional to viscosity within an ATPS as the polymer concentration is increased but that viscosity alone cannot explain the observed differences in mobility. For example, in Figure 5A, while for each polymer pair, mobility increases substantially as the viscosity is decreased, different polymer pairs (e.g. PEG 8k with dextran 10 vs 100 kDa) can have different mobilities, even when their viscosities are quite similar. This behavior can be rationalized by differences in the position of the wires at the polymer interface in these different ATPS. If the nanowires are preferentially wetted by one of the polymers, as were the nanospheres, then changing the ATPS composition could influence the relative position of the wires at the interface (i.e., largely in the PEG phase vs largely in the dextran phase). As a result, the mobility of the wires could vary significantly between two or more samples, although the viscosity of the individual phases

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Figure 5. Plot of wire mobility (as rotation in a 20-s time frame) versus viscosity in the upper and lower phases for a series of ATPS compositions. Data are plotted for both the top (A) and bottom (B) phases of ATPS comprising PEG 8 kDa and dextrans of various molecular weights and for the top (C) and bottom (D) phases of ATPS comprising dextran 100 kDa and PEGs of various molecular weights. For each ATPS, three points are shown, corresponding to the polymer weight percents shown in Table 2 (higher weight percents correspond to higher viscosities).

is similar. For example, in Figure 5B, the viscosity difference in wire mobility between 7 and 13 wt % of PEG 8 kDa/dextran 10 kDa is quite large, but the change in viscosity for the more viscous, dextran-rich phase is relatively small. This could be the result of greater wetting by the more viscous dextran-rich phase at the higher polymer weight percents. It should be noted that, although the difference in surface tension between these two samples is an order of magnitude, interfacial tension does not explain the observed differences in wire mobility (i.e., we observe very different mobilities for some ATPS with very similar interfacial tensions). This supports the interpretation that differences in nanowire wettability are largely responsible for the observed mobility results. For nanowire assembly purposes, although all of the ATPS compositions investigated here had sufficient interfacial tension to support Au nanowires against the force of gravity, the composition of the ATPS had a significant effect on wire mobility at the interface. By changing the polymer concentration and molecular weight, the interfacial tension, viscosity, and wire mobility at the interface could be tuned over a large range. The highest mobilities were observed for ATPS compositions with lower polymer molecular weights and concentrations. Biorecognition-Driven Nanoparticle Assembly at the Aqueous/Aqueous Interface. The biocompatibility of the polymers along with the ability to control the conditions within the ATPS (e.g., salt concentration, temperature, and pH) have long made these systems attractive for the separation of biomolecules;1,35 however,

no biomolecule-directed assembly experiments have been conducted at this polymer-polymer interface. It should be noted that volume exclusion effects due to high concentrations of macromolecules (e.g., PEG, dextran) can dramatically influence association equilibria38,39 and have been found to improve hybridization efficiencies through increasing effective DNA concentrations.40 However, under some conditions, these effects can also lead to an increase in the stability of mismatched or noncomplementary duplexes. We have previously shown that when the concentration of PEG is high (e.g., >10 wt %, as it would be in the top phase of an ATPS), macromolecular crowding effects can, under some conditions, result in the nonspecific association of DNA/Au nanoparticle bioconjugates.20 Thus, we were interested not only in determining whether the nanowire-bound DNA would still bind to its complementary strand while at the interface, but also in whether this process would occur selectively. DNA-coated Au wires were added to an ATPS interface where a fluorescently labeled complementary oligonucleotide had already been dispersed in the dextran phase during preparation. After 36 h, the wires were removed from the interface, washed a minimum of six times in hybridization buffer, and imaged using fluorescence microscopy. Figure 6 compares fluorescence images for DNA/Au nanowire bioconjugates after 36 h of hybridization to complementary or noncomplementary fluorescent (38) Minton, A. P. J. Biol. Chem. 2001, 276, 10577-10580. (39) Ellis, R. J. Trends Biochem. Sci. 2001, 26, 597-604. (40) Wooley, P.; Wills, P. R. Biophys. Chem. 1985, 22, 89-94.

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Figure 7. TEM micrograph of the hybridization of 12-nm Au colloids to a 3-µm × 320-nm Au wire in the absence (A) and presence (B) of a complementary linking oligonucleotide. The ATPS used in this experiment contained 9 wt % of PEG 8 kDa, 9 wt % of dextran 10 kDa, and 0.3 M hybridization buffer. The scale bars are 500 nm.

Figure 6. Reflectance (left panel) and fluorescence (right panel) optical microscopy images from 2-µm × 320-nm Au wires after hybridization for 36 h at the ATPS interface in the presence of a fluorescent complementary (A) or noncomplementary (B) oligonucleotide. Samples were removed from the ATPS interface and rinsed prior to imaging. Scheme 1 . DNA-Directed Assembly of Au Nanospheres onto Au Nanowires

Figure 8. First derivative plot of extinction during thermal denaturation of the DNA-directed Au nanosphere-nanowire assemblies shown in Figure 7. Extinction was measured at 520 nm, the characteristic absorbance of the nanospheres, as they were released from the nanowire assemblies at the bottom of the cuvette by thermal denaturation of the linking oligonucleotides.

oligonucleotides at the ATPS interface. These data indicate that DNA hybridization occurs at the interface with minimal nonspecific binding. We then performed biomolecule-directed particle assembly at the ATPS. Single-stranded DNA was attached to the surface of 320 nm × 3 µm nanowires via NeutrAvidin/biotin chemistry and bound to DNA/Au nanosphere bioconjugates via a complementary linker strand of DNA (Scheme 1). To avoid nonspecific crowding effects in the PEG-rich phase, DNA/Au nanosphere bioconjugates, having already been exposed to the complementary linker DNA strand, were added to the ATPS prior to phase separation. These bioconjugates partitioned into the lower dextran-rich phase upon phase separation (as in Figure 2C and D). Nanowires were then applied to the interface through the top phase for hybridization to the DNA/Au bioconjugates over a period of 48 h. Control experiments were performed in the absence of the linking strand. Following their removal from the interface and subsequent washing in buffer, the nanowires were imaged using TEM. Representative TEM images are shown in Figure 7. Although very few nanospheres are visible in the control experiment, the sample to which the complementary linking strand was added shows a high coverage of 14-nm spheres over the surfaces of the wires. These

data indicate that the nanoparticles are linked to the Au wires at the interface through hybridization and not as the result of nonspecific interactions. Thermal denaturation experiments were conducted to verify that the particle assemblies imaged using TEM were the result of DNA hybridization. Figure 8 shows the first derivative of the melting curve obtained for both the complementary and noncomplementary samples. Melting curves were determined by monitoring the increase in extinction at 520 nm due to the plasmon absorbance of the Au nanospheres as they are removed from the surface of the nanowire due to thermal denaturation of the DNA duplexes. An increase in this absorbance was recorded for the nanowire/nanosphere assemblies around 54 °C in the presence of the complementary linking oligonucleotide. No similar transition was recorded for the control. These data confirm that nanosphere binding to the surface of the Au wire is the direct result of DNA hybridization and not the result of nonspecific binding due to polymer crowding or other effects. In these experiments, we have taken advantage of partitioning to control the initial location of the DNA-coated 14-nm nanoparticles to prevent nonspecific aggregation due to macromolecular crowding by the PEG-rich phase, which is significant at these salt concentrations (0.3 M NaCl). Importantly, these data show

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that selective biorecognition-directed particle assembly can be performed at the aqueous/aqueous interface of an ATPS. Conclusion The behavior of Au and Ag nanospheres and Au wires and their bioconjugates has been investigated in a variety of PEG/dextran ATPS. Surface chemistry was found to be important for controlling the partitioning of Au and Ag nanospheres between the PEG-rich and dextran-rich phases. ATPS are a promising means of separation for nanoparticles having different adsorbed molecules and biomolecules and might, for example, be useful in separating protein/particle conjugates with different proteins or different protein conformations (i.e., native vs denatured). These types of separation experiments are routine for biomolecule separations, and we have observed for both proteins and DNA oligonucleotides that conjugation to Au nanospheres increases the partitioning substantially.41 Larger metal spheres and wires assemble at the aqueous/aqueous interface between the phases. The (41) Long, M. S.; Helfrich, M. R.; Keating, C. D. Polym. Mater. Sci. Eng. 2005, 92, 554.

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mobility of the wires is influenced by the composition of the ATPS, which controls polymer partitioning, phase viscosity, and interfacial tension. DNA oligonucleotides on the wire surface at the interface were capable of selective hybridization to complementary DNA strands that had partitioned into the dextran-rich phase. Using this approach, 14-nm Au nanospheres were selectively bound to the surface of 3-µm × 320-nm diameter Au wires via DNA-directed assembly. Although polymer solution viscosity and macromolecular crowding pose challenges for particle assembly at the ATPS, the tunability and biocompatibility of the polymer-polymer interface, coupled with the ability to control the initial positions of the reactants, makes ATPS attractive for biorecognitiondirected assembly of nanomaterials. Acknowledgment. This work was supported by the NSF (CHE-0239629), DARPA/ONR (Contract No. ONRN00014-98-1-00846), NSF-NIRT (CCR-0303976), and Penn State University. We thank the PSU Materials Research Institute for use of the rheometer. C.D.K. also acknowledges support from a Beckman Foundation Young Investigator Award and a Sloan Fellowship. LA051220Z