Supramolecular Assemblies of Amphiphilic Homopolymers - Langmuir

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Supramolecular Assemblies of Amphiphilic Homopolymers Tejaswini S. Kale, Akamol Klaikherd, Bhooshan Popere, and S. Thayumanavan* Department of Chemistry, University of Massachusetts, Amherst, Massachusetts 01003 Received March 2, 2009. Revised Manuscript Received April 18, 2009 Amphiphilic molecules self-assemble in solvents because of the differential solvation of the hydrophilic and lipophilic functionalities. Small-molecule surfactants have long been known to form micelles in water that can solubilize lipophilic guest molecules in their water-excluded interior. Polymeric surfactants based on block copolymers are also known to form several types of aggregates in water owing either to the mutual incompatibility of the blocks or better solvation of one of the blocks by the solvent. Incorporating amphiphilicity at smaller length scales in polymers would provide an avenue to capture the interesting properties of macromolecules and fine tune their supramolecular assemblies. To address this issue, we designed and synthesized amphiphilic homopolymers containing hydrophilic and lipophilic functionalities in the monomer. Such a polymer can be imagined to be a string of small-molecule surfactants tethered together such that the hydrophilic and lipophilic functionalities are located on opposite faces, rendering the assemblies facially amphiphilic. This feature article describes the self-assembly of our amphiphilic homopolymers in polar and apolar solvents. These homopolymers not only form micelles in water but also form inverse micelles in organic solvents. Subtle changes to the molecular structure have been demonstrated to yield vesicles in water and inverted micelles in organic solvents. The characterization of these assemblies and their applications in separations, catalysis, and sensing are described here.

Introduction The self-organization of surfactants is known to yield assemblies such as micelles, vesicles, fibers, and helical structures, to name a few, because of the differential interaction of the hydrophilic and lipophilic moieties with the solvent in which these are dissolved.1-5 Assemblies displaying container properties, such as micelles and vesicles, are interesting for applications in controlled and targeted drug delivery and protein sensing, among others.6-8 This is because micelles are capable of solubilizing lipophilic guest molecules, such as therapeutics, for instance, in an aqueous environment by localizing these into their waterexcluded lipophilic core. Tailoring the aggregation properties and the chemistries of the outer shell and the core of these structures, therefore, is of considerable interest. Both small molecules and polymers are known to undergo such self-assembly when containing functionalities that are incompatible with each other but can have low-energy interactions with *Corresponding author. E-mail: [email protected]. (1) Evans, D. F.; Wennerstrom, H. The Colloidal Domain, 2nd ed.; Wiley-VCH: New York, 1999. (2) Menger, F. M. Acc. Chem. Res. 1979, 12, 111–117. (3) Miyake, M.; Yamada, K.; Oyama, N. Langmuir 2008, 24, 8527–8532. (4) Tung, S. H.; Lee, H. Y.; Raghavan, S. R. J. Am. Chem. Soc. 2008, 130, 8813– 8817. (5) Vauthey, S.; Santoso, S.; Gong, H.; Watson, N. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 5355–5360. (6) Ganta, S.; Devalapally, H.; Shahiwala, A.; Amiji, M. J. Controlled Release 2008, 126, 187–204. (7) Ghosh, S.; Irvin, K.; Thayumanavan, S. Langmuir 2007, 23, 7916–7919. (8) Kataoka, K.; Harada, A.; Nagasaki, Y. Adv. Drug Delivery Rev. 2001, 47, 113–131. (9) Brunsveld, L.; Folmer, B. J. B.; Meijer, E. W.; Sijbesma, R. P. Chem. Rev. 2001, 101, 4071–4098. (10) Discher, B. M.; Won, Y. Y.; Ege, D. S.; Lee, J. C.-M.; Bates, F. S.; Discher, D. E.; A., H. D. Science 1999, 284, 1143–1146. (11) Fuhrhop, J.-H.; Wang, T. Chem. Rev. 2004, 104, 2901–2937. (12) Lee, M.; Cho, B. K.; Zin, W. C. Chem. Rev. 2001, 101, 3869–3892. (13) Moffitt, M.; Khougaz, K.; Eisenberg, A. Acc. Chem. Res. 1996, 29, 95–102. (14) Neiser, M. W.; Muth, S.; Kolb, U.; Harris, J. R.; Okuda, J.; Schmidt, M. Angew. Chem., Int. Ed. 2004, 43, 3192–3195. (15) Zhang, L.; Yu, K.; Eisenberg, A. Science 1996, 272, 1777–1779.

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the bulk solvent.1-5,9-16 For instance, when the relative degree of hydrophilicity and lipophilicity in a molecule are tuned, the hydrophilic groups can effectively shield the lipophilic functionalities in an aqueous medium and form assemblies with a water-excluded interior. Similarly, in an apolar solvent, the well-solvated lipophilic groups can encapsulate the hydrophilic functionalities. Such assemblies can in fact solubilize small numbers of water molecules in their interiors, the so-called “water pool”. Apart from possessing the required hydrophilic-lipophilic balance (HLB), these molecules must be present in sufficient concentration to form assemblies. The minimum concentration of a surfactant, at which the small molecule is in equilibrium with its amphiphilic assembly, is called its critical aggregation concentration, or cac. When referring to micelles, this is called the critical micellar concentration, or cmc.17-20 Small-molecule surfactants are popularly used as detergents, emulsifiers, stabilizers, and so forth. However, their large cmc values and the relatively low mechanical stabilities of their assemblies inherently limit their applications.1-5 Polymeric surfactants, however, form similar assemblies but with better stabilities and considerably lower cmc values. These have therefore received considerable attention.9-15 Polymeric surfactants may be broadly classified into homopolymers and block, random, alternate, and graft copolymers. Among these, block copolymers are of particular interest because these are known to form a variety of assemblies, the morphology and structure of which depend on each of the block lengths and their chemistries.17,20-23 The driving force for their self-assembly in solution is the (16) Okhapkin, I. M.; Makhaeva, E. E.; Khokhlov, A. R. Adv. Polym. Sci. 2006, 195, 177–210. (17) Bates, F. S. Science 1991, 251, 898–905. (18) Forster, S.; Plantenberg, T. Angew. Chem., Int. Ed. 2002, 41, 688–714. (19) Klok, H.-A.; Lecommandoux, S. Adv. Mater. 2001, 13, 1217–1229. (20) Ryu, D. Y.; Shin, K.; Drockenmuller, E.; Hawker, C. J.; Russell, T. P. Science 2005, 308, 236–239. (21) Zhang, L.; Eisenberg, A. J. Am. Chem. Soc. 1996, 118, 3168–3181. (22) Zhang, L.; Eisenberg, A. Macromolecules 1996, 29, 8805–8815. (23) Zhang, L.; Yu, K.; Eisenberg, A. Langmuir 1996, 12, 5980–5984.

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Invited Feature Article Scheme 1. Schematic Representation of Micelle-Type and InverseMicelle-Type Assemblies

Figure 1. Structure of amphiphilic polymers based on styrene and acrylamide.

immiscibility of either one of the blocks in the solvent or their mutual incompatibility. Block copolymers constituted from a hydrophilic and a lipophilic block are well known in the literature.24-31 For example, poly(styrene-b-acrylic acid) forms selfassembled structures owing to the mutual incompatibility of the polystyrene block and the poly(acrylic acid) block along with its immiscibility in water. This has led to a number of interesting structures that have been exhaustively studied.22,32-36 It will be interesting if such incompatibility could be engineered on a smaller length scale (i.e., on a monomer scale) because this would allow great tunability in the supramolecular assemblies. We conceived of achieving this by incorporating amphiphilic properties within a monomer unit (i.e., by synthesizing amphiphilic homopolymers). In this feature article, we focus on the molecular design, properties, and applications of this unique class of molecules in catalysis, sensing, and separations, as developed in our group. We first describe the unique features of both the micellar and inverse micellar assemblies of the amphiphilic homopolymers. We also show that structural tuning at the molecular level in these polymers can afford either micelle-type assemblies or vesicle-type assemblies. Then, we describe the utility of these assemblies in various applications. The inverse micellar assemblies have been used to separate peptides on the basis of their isoelectric points (pI). In the case of micelles, we have utilized these as reaction vessels in the aqueous phase. These amphiphilic assemblies are unique in that the sequestered lipophilic substrates seem to be significantly more site-isolated than in block-copolymer-based and small-molecule-based amphiphilic assemblies. We have also utilized the charged exterior of these amphiphilic nanoassemblies to bind to proteins. The polymer-protein interaction has been utilized both to modulate the enzymatic activity as well as to introduce a new concept in protein sensing. In fact, these amphiphilic assemblies provide a rather unique opportunity to introduce a new concept for creating patterns in sensing. (24) Motala-Timol, S.; Jhurry, D.; Zhou, J. W.; Bhaw-Luximon, A.; Mohun, G.; Ritter, H. Macromolecules 2008, 41, 5571–5576. (25) Chan, S. C.; Kuo, S. W.; Lu, C. H.; Lee, H. F.; Chang, F. C. Polymer 2007, 48, 5059–5068. (26) Tung, P. H.; Kuo, S. W.; Chan, S. C.; Hsu, C. H.; Wang, C. F.; Chang, F. C. Macromol. Chem. Phys. 2007, 208, 1823–1831. (27) Colombani, O.; Ruppel, M.; Schubert, F.; Zettl, H.; Pergushov, D. V.; Muller, A. H. E. Macromolecules 2007, 40, 4338–4350. (28) Colombani, O.; Ruppel, M.; Burkhardt, M.; Drechsler, M.; Schumacher, M.; Gradzielski, M.; Schweins, R.; Muller, A. H. E. Macromolecules 2007, 40, 4351–4362. (29) Bhargava, P.; Tu, Y. F.; Zheng, J. X.; Xiong, H. M.; Quirk, R. P.; Cheng, S. Z. D. J. Am. Chem. Soc. 2007, 129, 1113–1121. (30) Liu, L. B.; Gao, X.; Cong, Y.; Li, B. Y.; Han, Y. C. Macromol. Rapid Commun. 2006, 27, 260–265. (31) Gohy, J. F.; Creutz, S.; Garcia, M.; Mahltig, B.; Stamm, M.; Jerome, R. Macromolecules 2000, 33, 6378–6387. (32) Astafieva, I.; Zhong, X. F.; Eisenberg, A. Macromolecules 1993, 26, 7339– 7352. (33) Burguiere, C.; Chassenieux, C.; Charleux, B. Polymer 2003, 44, 509–518. (34) Dire, C.; Charleux, B.; Magnet, S.; Couvreur, L. Macromolecules 2007, 40, 1897–1903. (35) Abraham, S.; Ha, C. S.; Kim, I. J. Polym. Sci., Part A: Polym. Chem. 2005, 43, 6367–6378. (36) Laruelle, G.; Francois, J.; Billon, L. Macromol. Rapid Commun. 2004, 25, 1839–1844.

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Highlights of each of these applications are discussed in the following sections.

Amphiphilic Homopolymers We have introduced amphiphilic homopolymers based on a polystyrene backbone in which the two meta positions are occupied by a hydrophilic carboxylic acid functionality and a lipophilic benzyl moiety as shown in Figure 1.37 Our design hypothesis is that this molecule facilitates the phase segregation of the amphiphilic moieties in the polymer assembly rendering these polymers facially amphiphilic because it allows for the two amphiphilic groups to be located on two faces of the assembly. Therefore, these polymers not only form micelles in aqueous medium but also form inverted micelles in apolar solvents such as toluene (Scheme 1). We also found that a hydrophilic functionality linked to a lipophilic one via a simple flexible linker attached to a polymerizable unit can also be converted into an amphiphilic homopolymer capable of forming both micelles and inverted micelles.38 The aggregation behavior of these polymers was studied using transmission electron microscopy (TEM), dynamic light scattering (DLS), and static light scattering (SLS) whereas the container properties were evaluated using dye-encapsulation studies. Pyrene is a useful spectroscopic probe for studying container properties because its emission spectrum bears signature characteristics based on its microenvironment. The ratio of the intensity of the first and third peaks (I1/I3) in the emission spectrum is indicative of the polarity of its microenvironment.32,39-41 Even though the solubility of pyrene in water is very low, a greater amount of pyrene is solubilized in water upon addition of the amphiphilic polymer. A plot of emission intensity against polymer concentration exhibits an inflection point at the critical aggregation concentration (cac). Similar experiments were carried out using water-soluble dyes such as rose bengal in toluene to obtain evidence of the formation of inverted micelles.42 The cac of these polymers was found to be in the submicromolar concentration range. The cac of small-molecule equivalents was found to be much higher than for their corresponding polymer, establishing the advantage of tethering these together. For instance, the cac of 6 and its small-molecule equivalent were found to be 5 mg/L (0.25 μM) and 450 mg/L (2.1 mM), (37) Basu, S.; Vutukuri, D.; Shyamroy, S.; Sandanaraj, B. S.; Thayumanavan, S. J. Am. Chem. Soc. 2004, 126, 9890–9891. (38) Savariar, E. N.; Aathimankandan, S. V.; Thayumanavan, S. J. Am. Chem. Soc. 2006, 128, 16224–16230. (39) Kalyanasundaram, K.; Thomas, J. K. J. Am. Chem. Soc. 1977, 99, 2039– 2044. (40) Kwon, G.; Naito, M.; Yokoyama, M.; Okano, T.; Sakurai, Y.; Kataoka, K. Langmuir 1993, 9, 945–949. (41) Wilhelm, M.; Zhao, C.-L.; Wang, Y.; Xu, R.; Winnik, M. A.; Mura, J.-L.; Riess, G.; Croucher, M. D. Macromolecules 1991, 24, 1033–1040. (42) Basu, S.; Vutukuri, D. R.; Thayumanavan, S. J. Am. Chem. Soc. 2005, 127, 16794–16795.

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Chart 1. Structures of Amphiphilic Polymers Synthesized to Study the Effect of Structure on the Aggregation Behavior of These Polymers

Table 1. Aggregation Behavior and Aggregate Properties of Amphiphilic Polymers38 polymer

cac (μM)

I1/I3 (above cac)

3 4 5 6 7 8 9 10 11 12

0.58 0.39 0.33 0.25 0.24 0.033 NA NA 5.8 0.074

1.10 1.10 0.98 1.20 1.16 1.14 1.85 1.75 1.52 1.22

respectively (Chart 1 and Table 1). An even greater cac gain was observed in the case of 8 and its small-molecule equivalent estimated as 0.6 mg/L (0.33 nM) and 136 mg/L (0.5 mM), respectively. We attribute this difference to the relative packing of the lipophilic chains in the polymer and the small molecule. The relative distance between two adjacent amphiphilic functionalities is dictated by the covalent bonds in the polymer, unlike in the small-molecule micelle. With the container properties established, we proceeded to understand the aggregation behavior in further detail. We suggested earlier that the micelle vs inverted micelle container properties are due to a molecular-level change in conformation. These hypotheses were supported by TEM and light-scattering experiments. TEM images clearly indicate the formation of micelles and inverted micelles when samples were drawn from water and toluene, respectively (Figure 2). The location of both hydrophilic and lipophilic groups in these assemblies was determined by placing heavy atoms on each of these functionalities individually. Because these polymers do not dissolve in neutral water, 1 to 2 equiv of base must be added to achieve dissolution by means of breaking the extended intermolecular hydrogen bonding network formed between the carboxylic acid groups. In one case, we used potassium hydroxide as the base, where the K+ would be the counterion for the hydrophilic carboxylate functionalities. We compared the TEM images obtained from this amphiphilic assembly with that obtained when CsOH was used as the base. The contrast obtained due to the vastly different electron density of Cs+ as compared to that of the other atoms in the molecule distinctly indicates a dark corona in the case of micelles and a dark core in the case of inverted micelles (Figure 2). Similarly, bromination of the hydrocarbon chain aided the location of this chain in the aggregates, where the dark core was observed in micelles and a dark corona was obtained in the case of inverse micelles. The radius of the assemblies was determined to be 35 nm from these experiments for aggregates of 1. However, because the samples were dried in order to obtain the TEM image, the observed size may not correlate to the solution-phase behavior of the polymers by this technique. DLS and SLS experiments were 9662 DOI: 10.1021/la900734d

therefore carried out to establish the size and shape of the aggregates in solution. The hydrodynamic radius, RH, of the assemblies in aqueous solution was found to be 53 nm whereas the radius of gyration, RG, was 41 nm. The RG/RH ratio, known as F, is indicative of the shape of the aggregates. In the case of 1, the ratio was about 0.77, further supporting the formation of micelles. These sizes, however, appear to be large for the aggregates to be considered micelles, yet the TEM results indicating a dark interior and the container properties establishing the sequestration of lipophilic guest molecules along with the shape-sensitive F suggest that the assemblies are indeed micellar. Such larger than expected aggregates are known in the literature for small-molecule surfactants as well as for block copolymers and are simply understood to be aggregates of micelles.43-46 If we were indeed able to control the nature of the assemblies by tuning the functional groups at the molecular level, then we conceived that vesicle-type assemblies could be obtained from bolamphphile-based amphiphilic homopolymers. In this case, the monomer unit would contain two hydrophilic functionalities connected by a lipophilic alkyl chain. We synthesized polymer 11 as the possible vesicular analog for polymer 8. We were gratified to observe that 11 indeed formed vesicle-type assemblies (Figure 2) whereas polymer 8 exhibited micelle-type assemblies. We had also hypothesized that the vesicle-type assemblies require a critical lipophilic chain length. To test this, we synthesized and tested polymers 9 and 10 with shorter hydrocarbon chains. We found that neither of these exhibited any supramolecular assemblies. We also conceived that such requirements are much less stringent with micelle-forming polymers; we observed that even polymer 6 exhibits micelle-type assemblies. Vesicle assemblies can be formed from block copolymers and from polymers containing two lipophilic tails for every hydrophilic headgroup.47-52 The advantage of our system is that we could dial in the self-assembly by tuning the structure on the molecular level. Also, the processing requirements are much easier with our amphiphilic homopolymers. The aggregates formed by 11 analyzed using DLS and SLS were found to have a hydrodynamic radius of 28 nm and a radius (43) Erhardt, R.; Zhang, M.; Boker, A.; Zettl, H.; Abetzc, C.; Fredrik, P.; Krausch, G.; Abetz, V.; Muller, A. H. E. J. Am. Chem. Soc. 2003, 125, 3260–3267. (44) Kujawa, P.; Tanaka, F.; Winnik, F. M. Macromolecules 2006, 39, 3048– 3055. (45) Burkhardt, M.; Ruppel, M.; Sandrine, T.; Drechsler, M.; Schweins, R.; Pergushov, D. V.; Gradzielski, M.; Zezin, A. B.; Muller, A. H. E. Langmuir 2008, 24, 1769–1777. (46) Chakraborty, H.; Sarkar, M. Langmuir 2004, 20, 3551–3558. (47) Schillen, K.; Bryskhe, K.; Melnikova, Y. S. Macromolecules 1999, 32, 6885– 6888. (48) Liu, F.; Eisenberg, A. J. Am. Chem. Soc. 2003, 125, 15059–15064. (49) Nardin, C.; Thoeni, S.; Widmer, J.; Winterhalter, M.; Meier, W. Chem. Commun. 2000, 1433–1434. (50) Du, J.; Armes, S. P. J. Am. Chem. Soc. 2005, 127, 12800–12801. (51) Ringsdorf, H.; Schlarb, B.; Venzmer, J. Angew. Chem., Int. Ed. 1988, 27, 113–158. (52) Tsuchida, E.; Nishide, H.; Yuasa, M.; Babe, T.; Fukuzumi, M. Macromolecules 1989, 22, 66–72.

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Invited Feature Article

Figure 2. TEM images of the micelle-like and inverted-micelle-like structures formed by polymers 1c, 1d, and 1e. (A) Image of a normal micelle-like particle from an aqueous solution of polymer 1c. (B) Image from an aqueous polymer 1e. (C) Image of an inverted micelle-like particle formed in a toluene solution of polymer 1c. (D) Image from a toluene solution of polymer 1e. (E) Image from an aqueous solution of polymer 1d. (F) Image from a toluene solution of polymer 1d. (G) Image of vesicles of polymer 11 in water. (H) Image of inverted micelles from a toluene solution of polymer 11.

of gyration of 29 nm. Shape-sensitive F was computed to be 1.04 in this case, indicating vesicle formation. TEM also confirms vesicle formation in water as the water-filled interior is enclosed in a lipophilic membrane that appears dark in the image. It is also interesting that these types of polymers are also soluble in apolar solvents such as toluene and form inverted micelles in these solvents. On the basis of the sizes obtained from TEM, these also appear to be compound micelles, or aggregates of inverted micelles. Another tool to establish the kind of interior of the aggregates is to analyze the container properties. As mentioned earlier, the emission spectrum of pyrene is indicative of its microenvironment. A detailed structure-property study was carried out on different polymers containing varying degrees of lipophilicity on the basis of varying hydrocarbon chain length, as shown in Chart 1, and the properties of their aggregates are as listed in Table 1. In all of the sets of polymers, the cac was found to decrease, as expected, with an increase in the lipophilic chain length. The lipophilicity of the interior also increases as seen from the ratio I1/I3, which decreases in magnitude with increasing chain length. It is interesting that polymers 9 and 10 do not provide a sufficiently different environment for pyrene compared to that for bulk water, as seen from the I1/I3 values. In fact, as mentioned earlier, these do not aggregate in water as confirmed by the fact that no cac could be determined for these polymers. The value of I1/I3 in the case of 11 was found to be 1.52, which is sufficiently different from that of bulk water but indicates higher polarity than that obtained from micellar interiors. This is because the lipophilic membrane in the vesicles is bound by water on both sides and thus is unable to exclude water as effectively as the micelles can. Polymer 12 provides a comparatively more lipophilic environment as the length of the lipophilic chain has increased as compared to that of 11. Our amphiphilic homopolymers exhibit micelle-type assemblies in the aqueous phase and inverse-micelle-type assemblies in apolar solvents. It is interesting to ask about the nature of the assembly when these homopolymers are dispersed in an immiscible mixture of aqueous and apolar solvents. To test this, we prepared a solution of the polymer in water and equilibrated it with an apolar solvent such as dichloromethane, in which the Langmuir 2009, 25(17), 9660–9670

polymer is known to dissolve. The same experiment was also carried out by first dissolving the polymer in an apolar solvent and equilibrating this with water. In both cases, we found that these polymers are kinetically trapped in the solvent in which they are initially assembled (i.e., the polymers do not cross over from one phase to another (Figure 3a)).42 We took advantage of this observation for separations, where we conceived the possibility of separating two different molecules dispersed in the aqueous phase by extracting one of the molecules into the inverse micelle interiors. To test this possibility, we dissolved a cationic dye (rhodamine 6G) and an anionic dye (rose bengal) in water. The carboxylic acid-based inverse micelle was able to extract the cationic dye selectively into the organic phase (Figure 3b). We utilized this principle to extract peptides selectively on the basis of their isoelectric points (pI).

Peptide Extraction Using Inverted Micelles Significant information from cell digests, multiprotein extracts, and peptides can be obtained using mass spectrometry, especially matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS).53-55 However complex samples, available as dilute solutions, pose significant challenges in this form of analysis. Although a number of sample cleanup methods have been developed, methods that also provide the selective extraction of analytes are less well known and are often based on conventional chromatography techniques.56 Liquid-liquid extractions can be used for this purpose for a combined effect of extraction, purification, and potential concentration in a single step. However, these are most commonly useful in the case of lipophilic analytes that can be solubilized in organic solvents. Biomolecules such as proteins, peptides, and fractions thereof are water-soluble molecules. If, however, one could solubilize these hydrophilic molecules into the organic phase, then it would be a good leap (53) Karas, M.; Hillenkamp, F. Anal. Chem. 1988, 60, 2299–2301. (54) Stults, J. T. Curr. Opin. Struct. Biol. 1995, 5, 691–698. (55) Tanaka, K.; Waki, H.; Ido, Y.; Akita, S.; Yoshida, Y.; Yoshida, T. Rapid Commun. Mass Spectrom. 1988, 2, 151–153. (56) Xu, Y.; Bruening, L. M.; Watson, J. T. Anal. Chem. 2004, 76, 3106–3111.

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Figure 4. Schematic representation of the selective extraction of complementary peptides from their aqueous solution by using inverse micelles of an amphiphilic homopolymer. Table 2. pH-Based Extraction of a Peptide Mixture Using Polymer 5 ion abundance ratio (K ( SM)a peptide (pI)

Figure 3. (a) Polymer micelles are kinetically trapped in the medium in which they are initially assembled. (b) Charge-based separation of dyes obtained by using reverse micelles of 5.

forward in their analysis. The amphiphilic polymers developed in our group form inverted micelles in organic solvents, and these assemblies do not migrate to the aqueous phase. Water can be solubilized in the hydrophilic interior of these micelles, rendering these as potential nanocontainers for hydrophilic guest molecules.38 Another aspect of these polymers is the charge-based specificity that can be introduced in the extraction of the guest molecules on the basis of the fact that our hydrophilic functionality is an anionic carboxylic acid. Therefore, positively charged guests can be selectively sequestered into the inverted micelles, as discussed earlier. We have applied this advantage to analyze peptide mixtures with the aim of eventually impacting protein identification and thus proteomics.57 Peptides are short amino acid sequences with a defined isoelectric point (pI). When in a solution of pH lower than its pI, the peptide is in the protonated state (i.e., it is positively charged). This can therefore be sequestered into the inverted micelles of amphiphilic polymer in an organic phase because it is reasonable to assume that the pH of the aqueous phase would also dictate the pH of the water pool inside the inverted micelle. The organic phase can eventually be analyzed using MALDI-MS. Such pH-based separations can lead to selective extraction based on pI and, when adjusted accurately, can greatly reduce the number of variables in the mixture analysis (Figure 4). We used a solution of 5 in toluene (100 μM) as the source of inverted micelles. This solution was mixed with an aqueous solution of peptides adjusted to suitable pH. This mixture was vortex mixed, followed by centrifugation to yield the phaseseparated mixture of which the organic phase was analyzed. Sequential treatment of the aqueous phase adjusted to appropriately varied pH led to fractionation of the peptide mixture into (57) Combariza, Y.; Savariar, E. N.; Vutukuri, D. R.; Thayumanavan, S.; Vachet, R. W. Anal. Chem. 2007, 79, 7124–7130.

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pH 4.0

pH 7.1

pH 9.0

bradykinin (12.5) 680 ( 138 1162 ( 73 842 ( 151 kinetensin (11.1) 337 ( 79 882 ( 212 616 ( 198 ACTH human (9.3) 221 ( 37 422 ( 87 128 ( 41 angiotensin (7.7) 237 ( 56 536 ( 116 34 ( 14 spinorphin (6.1) 36 ( 12 2.0 ( 1.0 1.0 ( 0.5 β-amyloid (4.1) 17 ( 6 0.4 ( 0.2 0.10 ( 0.07 preproenkephalin (3.6) 2.2 ( 0.9 0.4 ( 0.1 0.12 ( 0.04 a The ion abundance ratio is defined as the ratio of the MALDI ion abundance of the peptide in the organic phase that contains inverted micelles to that in the aqueous phase after the extraction. The ion abundance ratio (K) and standard errors of the mean (SM) are for the [M + H]+ signal except for spinorphin and β-amyloid, in which case the values were calculated using the [M + K]+ signal. The SM values were calculated from at least four replicate measurements in each case.

small fractions based on their pI identified by the ion abundance in MALDI-MS using R-cyano-hydroxycinnamic acid as the matrix. The results of one such peptide extraction and analysis experiment are shown in Table 2. Here, a mixture of peptides with pI values ranging from 12.5 to 3.6 was buffered to either pH 9.0, 7.1, or 4.0 and an extraction was performed using polymer solution in toluene. In each case, it was observed that peptides bearing a pI value greater than the pH of the solution were extracted efficiently whereas those with lower pI values were left behind in the aqueous phase. An interesting observation was made while performing these peptide extraction experiments. We found that because peptides are inherently insoluble in the organic phase no signal was obtained when the organic phase was analyzed using MALDIMS in the absence of the amphiphilic homopolymer. However, the inverse micelles of the polymer provide a favorable microenvironment for solubilizing the peptide, and manifold signal amplification was observed in the MALDI-MS of the organic phase upon addition of the polymer. This method not only provides a selective extraction technique but also yields samples in a suitable concentration range to be analyzed by MALDI-MS. The pI cutoff for the selectivity in the extraction can be tuned by simply adjusting the pH of the solution. We also find that the increased sensitivity in ion signals is due to not only the concentration effect but also the fact that the Langmuir 2009, 25(17), 9660–9670

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Invited Feature Article Scheme 2. Photo-Fries Rearrangement of 1-Naphthyl Benzoate

polymeric material provides an enhancement in MALDI ion yields. This method has further been applied to the analysis of protein digests with promising results. We are currently investigating the possibility of changing the hydrophilic functionality to tune the extraction capability further. The simplicity of this method along with its applicability in analyzing dilute solutions makes it immensely useful in analyzing complex mixtures of biomolecules currently limited by the extraction and purification techniques.

Nanoreactors for Photochemical Organic Reactions in Water Owing to the water-excluded microenvironment present in the interior of the polymer micelles, organic reactants can easily be solubilized in these micelles. These reactants can then be subjected to appropriate reaction conditions to carry out organic reactions in water. We utilized the micellar solution of 1 prepared in basic water (pH 8.5) to carry out photochemical reactions in order to understand further the nature of the microenvironment provided by the polymer.58 In one case, we added 1-naphthyl benzoate to a solution of 1 to encapsulate it into the interior of the micelles. This molecule, when dissolved in hexane, is known to undergo photo-Fries rearrangement upon irradiation and produce four products as shown in Scheme 2. The reactant can, however, also undergo hydrolysis to form R-napthol and benzoic acid if it were exposed to the bulk basic aqueous medium because we need basic water to dissolve the polymer. We carried out this reaction in solutions of different surfactants and were gratified to find that 1 provided the micellar environment, which completely suppressed the base hydrolysis and yielded 2-benzoyl-1-naphthol (compound a) as the only product (Scheme 2). This confirms the existence of a waterexcluded interior in the micelles of this amphiphilic homopolymer. In contrast, the micellar environment provided by amphiphilic block copolymer poly(styrene-co-sodium acrylate) (PS-b-PSA) prepared at pH 9.5 led to considerable hydrolysis products being generated along with the ortho and para rearrangement products in a ratio of 1:8. The product distribution obtained using smallmolecule surfactants such as cetyl trimethylammonium chloride (CTAC), sodium dodecyl sulfate (SDS), and sodium dodecanoate (SDO) as shown in Table 3 also consisted of a significant amount of para rearrangement product. Further investigation into the nature of micellar interiors was carried out using other reactants.59 The product profile obtained from these led us to the conclusion that the lipophilic reaction cavities of 1 confine the sequestered guest molecules to a larger extent than do smallmolecule-based and block-copolymer-based surfactants.

adapt their conformations upon binding to surfaces of biomolecules.60-62 Moreover, multiple contacts between the polymer and the protein surfaces can provide a significant enhancement in binding efficiency. Such interactions with proteins render polymers attractive candidates for protein surface binding implications in protein-protein interactions, protein-nucleic acid interactions, and the development of new enzyme inhibitors. Although the nature of the interaction between polymer and protein can be covalent or noncovalent, covalent modification of a protein with a polymer offers the possibility of irreversibly modifying its biological activity. Noncovalent interactions of proteins with synthetic macromolecules, however, offer the possibility of reversible binding and modulation of their function. Our amphiphilic polymers provide a unique opportunity because of their ability to form spherical micellar nanoassemblies with a negatively charged exterior providing a platform for noncovalent interaction with the positive motif of the protein via electrostatic interaction.63 To study the polymeric micelle-protein complexes, we chose R-chymotrypsin (ChT) with a pI of 8.8 as a model protein. Because the cationic patch of this protein is located at the active site, it provides an opportunity to examine the polymer-protein interaction through enzyme inhibition assays. (See Figure 5 for a schematic representation.) The interaction between polymeric micelle and ChT was first quantified by using nondenaturing gel electrophoresis and enzyme assay. The results indicate that the binding ratio of polymer/protein was 1:10 on the basis of the maximal intensity in the neutral region compared to that on the anodic or cathodic side of the gel. The inhibition of ChT activity by the polymer was monitored by the hydrolysis of a chromogenic substrate, N-succinyl-L-phenylalanine-p-nitroanilide (SPNA). The studies were carried out with different concentrations of polymer 1 ranging from 10-6 to 10-8 M while maintaining the concentration of ChT constant. The binding constant and ratio of polymer 1/ChT were obtained by plotting the activity of ChT against polymer concentration. The dissociation constant was determined to be 7  10-7 M with a binding ratio of 1:10 polymer/ChT. The binding ratio estimated here is consistent with the results obtained from the gel electrophoresis study. The possibility of denaturing ChT was investigated using fluorescence and circular dichroism. Both techniques indicated that there was no significant change in the protein structure although the lipophilic polymer backbone and the benzyl substituent are intimately associated with the anionic carboxylate moiety. We attribute this to the flexibility of polymer 1 that most likely adapts to the ChT surface rather than forcing the protein to adapt to the anionic surface of the nanoassembly.64

Polymer-Protein Interaction Artificial polymers are of great interest for their interactions with biological polymers as a result of their potential to (58) Arumugam, S.; Vutukuri, D.; Thayumanavan, S.; Ramamurthy, V. J. Am. Chem. Soc. 2005, 127, 13200–13206. (59) Arumugam, S.; Vutukuri, D.; Thayumanavan, S.; Ramamurthy, V. J. Photochem. Photobiol., A 2007, 185, 168–171.

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(60) Gestwicki, J. E.; Strong, L. E.; Kiessling, L. L. Chem. Biol. 2000, 7, 583–591. (61) Mammen, M.; Choi, S. K.; Whitesides, G. M. Angew. Chem., Int. Ed. 1998, 37, 2755–2794. (62) Mourez, M.; Kane, R. S.; Mogridge, J.; Metallo, S.; Deschatelets, P.; Sellman, B. R.; Whitesides, G. M.; J., C. R. Nat. Biotechnol. 2001, 19, 958–961. (63) Sandanaraj, B. S.; Vutukuri, D. R.; Simard, J. M.; Klaikherd, A.; Hong, R.; Rotello, V. M.; Thayumanavan, S. J. Am. Chem. Soc. 2005, 127, 10693–10698. (64) Fischer, N. O.; Verma, A.; Goodman, C. M.; Simard, J. M.; Rotello, V. M. J. Am. Chem. Soc. 2003, 125, 13387–13391.

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Table 3. Product Distribution Obtained from a Reaction Carried out in Nanocontainers Formed Using Various Surfactants58 product distribution (%) medium hexane NaOH(aq) (pH 8.5) 1 (pH 8.5) PS-b-PSA (pH 9.5) CTAC SDS SDO

conc (mM)

a

b

c

d

30

7 60

3

0.11 0.11 10 10 100

60 2 >99 50 87 83 75

6 12 14 21

23 1 2 3

benzoic acid 38 21

1

Figure 6. Normalized activity of ChT (3.2 μM) with polymer 1 (0.8 μM) against three different substrates A, B, and C.

Figure 5. Interaction of anionic amphiphilic homopolymer micelle with the positive patch of protein.

The interaction of the polymer and protein is based on electrostatics. (The negatively charged carboxylate groups at the periphery of the micelles can have coulombic interactions with positively charged regions of the protein.) Fluorescence and CD studies show that the protein is not denatured in the polymerprotein complex. It should therefore be possible to release the bound ChT from the polymer surface by increasing the ionic strength of the medium and thus recover its enzymatic activity.65 We increased the ionic strength to 200 nM and found this indeed to be the case, which also suggests that the binding of ChT to polymer 1 is reversible. This provides additional evidence to suggest that electrostatics is a major driving force in this polymer-protein interaction. It is therefore interesting to examine the activity of the protein by changing the charge of the substrates. Such a comparison of substrates should provide information on the effect of charge within the substrate upon the activity of the polymer-enzyme complex (Figure 6). The polymer-ChT complex was studied for anionic, neutral, and cationic substrates. We found that the negative charge of SPNA can influence the inhibition efficiency through the steric effect and electrostatic interaction. Inhibition with the neutral substrate shows the extent of steric inhibition of binding. However, the hyperactivity of ChT with the cationic substrates suggests that electrostatic attraction between the substrate and the polymer-protein complex dominates the steric hindrance offered by the polymer. It is also interesting to be able to utilize the polymer--protein interactions to disrupt protein-protein interactions; the latter has been implicated in several diseases.66-68 Cytochrome c (Cc) is a cationic protein with a pI of 10.3; the heme edge of Cc is surrounded by an array of lysine and arginine residues, and cytochrome c peroxidase (CcP) is an anionic protein with a pI of 5.25, with 45 glutamate and aspartate residues.65 The interaction between Cc and CcP is primarily driven by electrostatics, and (65) Scott, R. A.; Mauk, A. G., Cytochrome c: A Multidisciplinary Approach: Univeristy Science Books: Sausalito, CA, 1996. (66) Guo, Z.; Zhou, D.; Schultz, P. G. Science 2000, 288, 2042–2045. (67) Lin, H.; Cornish, V. W. Angew. Chem., Int. Ed. 2001, 40, 871–874. (68) Rain, J. C.; Selig, L.; Reuse, H. D.; Battaglia, V.; Reverdy, C.; Simon, S.; Lenzen, G.; Petel, F.; Wojcik, J.; Schachter, V.; Chemama, Y.; Labigne, A.; Legrain, P. Nature 2001, 409, 211–215.

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Figure 7. Interaction of the anionic amphiphilic homopolymer assembly with cytochrome c, a positively charged protein, to interrupt the cytochrome c (Cc)-cytochrome c peroxidase (CcP) interaction.

the binding affinity understandably decreases with increasing ionic strength.69 Similar to ChT-polymer interaction, because polymer 1 (Chart 1) is negatively charged at 10 mM sodium phosphate buffer pH 6.0, we thought that this polymer would bind to the cationic protein, Cc (Figure 7).70 To investigate the binding of polymer 1 with Cc, native gel electrophoresis was performed, which shows that the binding ratio of polymer to Cc is approximately 1:8. Absorption spectroscopy and circular dichroism (CD) were used to investigate further the interaction between polymer 1 and Cc. Upon polymer binding with Cc, the heme moiety in the protein is solvent-exposed, and a water molecule replaces the Met80 ligand.66 Because the heme group of native Cc is deeply buried inside the protein and Met80 prevents the binding of H2O2 to iron, the rate of reaction with peroxide is very low.71 However, when the Met80 ligand is replaced by the more labile H2O and the heme group is solvent-exposed, the rate of reaction with H2O2 increases significantly as seen with unfolded and chemically modified Cc. Binding Fe(II)-Cc to polymer 1 increases the rate of oxidation to a maximal value of kobs = 0.0137 s-1, and exposure of the heme group of Cc to bulk solvent could be achieved by increasing the concentration of polymer 1. The exposure of the heme group is also further supported by cyclic voltammetry. Considering the strong interaction between polymer 1 and the Cc complex, this result encouraged us to determine whether this polymer can be used as a scaffold to disrupt Cc-CcP interactions. (69) Kresheck, G. C.; Vitello, L. B.; Erman, J. E. Biochemistry 1995, 34, 8398– 8405. (70) Sandanaraj, B. S.; Bayraktar, H.; Krishnamoorthy, K.; Knapp, M.; Thayumanavan, S. Langmuir 2007, 23, 3891–3897. (71) Moore, G. R.; Pettigrew, G. W. Cytochrome c: Evolution, Structural and Physiochemical Aspects. Springer: New York, 1990; pp 115-159.

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Figure 8. Synthesis of polymer 13.

Electrochemistry is a useful technique in this case.72 The complexation of Cc and CcP is driven by charged residues present on the exterior surface of two proteins, not by the metal ion in the heme group, which is present in the interior of the protein. Because CcP binds near the solvent-exposed face of the heme in Cc, such binding is anticipated to decrease the ability of the heme group to interact with the electrode surface, thereby decreasing the rate of electron transfer. The electron-transfer rate (k0) of Cc in the presence of Zn-CcP was smaller (6.4  10-5 cm/s) than in the absence of Zn-CcP. After polymer 1 was added to the preformed Cc/CcP complex, the electron-transfer reactivity of Cc was measured. The k0 of the Cc/CcP complex (6.4  10-5 cm/s) increased by nearly 3 orders of magnitude in the presence of polymer 1 (1.5  10-2 cm/s), which is very close to the value of the Cc/polymer 1 complex (2.25  10-2 cm/s). The observed results confirm our model that polymer 1 has the ability to disrupt the interaction of Cc with CcP by selectively binding to Cc. Using an artificial polymeric scaffold to modulate the protein functions noncovalently and reversibly is of interest. Such interactions also offer a new avenue for the use of amphiphilic polymer assemblies for protein sensing, which is our next topic of discussion.

Protein Sensing Polyelectrolytes have been shown to interact with proteins on account of polyvalent interactions of an electrostatic nature, as exemplified above.73 These are water-soluble molecules that display several pendant charges, either cationic or anionic or both. Polymers in this class have been the subject of extensive research because of their potential applications in fields such as controlled drug release, surface coatings, and chemical and biological sensing, among several others. Conjugated polyelectrolytes have been the favorite candidates for fluorescence-based sensing of proteins on account of the inherent fluorescence of the polymer backbone and their water solubility.73-76 The likelihood of energy or electron transfer or both from the polymer to the biological analyte resulting in a change in the fluorescence pattern of the polymer renders them potent for sensing metalloproteins. In this case, protein cofactors such as the metalloporphyrins can accept energy or electrons from the excited state of the conjugated polymer. However, it has been demonstrated that such a change in fluorescence could also be a result of the polymer binding to nonmetalloproteins.77 A change in the polymer conformation upon binding to a target analyte results in a change in the

(72) Chen, L.; McBranch, D. W.; Wang, H.-L.; Helgeson, R.; Wudl, F.; Whitten, D. G. Proc. Natl. Acad. Sci. U.S.A. 1999, 96, 12287–12292. (73) Seyrek, E.; Dubin, P. L.; Tribet, C.; Gamble, E. A. Biomacromolecules 2003, 4, 273–282. (74) Sandanaraj, B. S.; Demont, R.; Aathimanikandan, S. V.; Savariar, E. N.; Thayumanavan, S. J. Am. Chem. Soc. 2006, 128, 10686–10687. (75) Wilson, J. N.; Wang, Y.; Lavigne, J. J.; Bunz, U. H. F. Chem. Commun. 2003, 1626–1627. (76) Fan, C.; Plaxco, K. W.; Heeger, A. J. J. Am. Chem. Soc. 2002, 124, 5642– 5643. (77) Kim, I.-B.; Dunkhorst, A.; Bunz, U. H. F. Langmuir 2005, 21, 7985–7989.

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Figure 9. Interaction of polymeric micelles that contain an anthracene unit (a) with nonmetalloproteins and (b) with metalloproteins.

fluorescence signature of the polymer. Thus, there is no guaranteed selectivity for metalloproteins. To address the issue of selectively sensing metalloproteins, we designed and synthesized a new class of nonconjugated fluorogenic polymers derived from polystyrene-based amphiphilic homopolymers78 (Figure 8). These polymers were shown to form optically clear nanoscale micellar assemblies in water.38 It was envisioned that the proximity of the fluorophore to the carboxylate moieties that bind to the protein would aid efficient energy or electron transfer. Furthermore, because the fluorophore is not a part of the polymer backbone, we hypothesized that any conformational change in the polymer due to binding would not affect the fluorescence response of the fluorophore. Thus, an appropriate energy or electron donating/accepting functionality (i.e., metalloprotein cofactor) must be contained within the protein to invoke the fluorescence response of the polymer (Figure 9) and therefore should be selective in sensing metalloproteins. The fluorescent anthracene pendant unit acts as the transducer for sensing, whereas the solvent-exposed carboxylate groups act as ligands for protein binding. The absorption and emission spectrum of 13 suggests that anthracene is efficiently incorporated into the polymer backbone and that the incorporation and aggregation in water do not alter the electronic properties of anthracene. The possibility of either energy or electron transfer from the excited state of anthracene to the metal cofactor in the protein should effect a change in the emission spectrum of anthracene upon polymer-protein binding. We tested our hypothesis of selective binding to metalloproteins by comparing the fluorescence response of polymer 13 to that of Cc (a metalloprotein) and lysozyme (a nonmetalloprotein). We observed that the fluorescence response from the anthracene moiety decreased as the Cc concentration was increased, whereas a similar increase in lysozyme concentration had no discernible effect on anthracene fluorescence. Because both proteins have (78) Sandanaraj, S. B.; Demont, R.; Thayumanavan, S. J. Am. Chem. Soc. 2007, 129, 3506–3507.

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similar pI values, the extent of their binding with the polymer can be assumed to be close to each other. The fluorescence decrease can be attributed to the fact that the porphyrin functionality in Cc is able to quench the excited state of anthracene by electron or energy transfer while the lysozyme is unable to do so because of the lack of a photoactive or electroactive functionality to access excited state of anthracene. We observed a concentration-dependent Stern-Volmer plot indicating a high KSV for Cc (2.0  105) whereas no change in fluorescence was observed for the lysozyme (KSV = 0). We also tested the fluorescence response of the same polymer (13) to 12 different proteins, 4 of which were metalloproteins, viz., ferritin, Cc, CcP, and myoglobin. We observed that all of the metalloproteins exhibited KSV values between 104 and 106. Irrespective of their pI values, the eight nonmetalloproteins did not affect the fluorescence response of the polymer, thereby indicating the selective response of our polymer to the metalloproteins. The sensing systems shown above are based on nonspecific (electrostatic) interactions. For sensing systems to be practical, one needs to incorporate greater selectivity. Selectivity can be introduced into a system through lock-and-key-type receptorligand interactions. An alternative to this approach involves the design of several nonspecific receptors, each of which responds differently to an analyte. From these, one can develop analytespecific patterns. For this approach to be effective in a complex set of analytes, it is essential that we have a large set of data points. If one could assemble the components of the receptor and/or the transducer noncovalently, then obtaining a large set of data points could become simpler. Although varying the receptor functionality to obtain patterns has been a topic of study,79 we have introduced the concept of utilizing variations in transducer structure to obtain analyte-specific patterns.80 The introduction of this orthogonal dimension into pattern generation could greatly simplify the pattern-based approach in sensing. The fact that the transducers are incorporated into the system by noncovalent interactions makes this approach even more attractive. We developed a new approach in that a single receptor binds nonspecifically to proteins and is capable of generating analytedependent patterns.81 In this approach, we harness the advantages offered by these amphiphilic homopolymers to (i) bind to proteins, (ii) form micelles in aqueous solutions, and (iii) encapsulate a lipophilic guest molecule. Thus, if the encapsulated molecule is able to transfer its excited-state energy or electrons to the bound protein, then one could generate patterns by simply varying the encapsulated fluorescent, lipophilic guest molecule (Figure 10). The concentration-dependent fluorescence quenching of the encapsulated pyrene was used to extract the Stern-Volmer quenching constant. The KSV for cytochrome c was observed to be 1.1  105, in good agreement with the value obtained by covalently attaching the dye molecule to the polymer backbone. The KSV values for three other metalloproteins;ferritin, hemoglobin, and myoglobin;were observed to be 1.0  106, 4.8  105, and 5.2  105, respectively, thereby indicating that a differential response toward metalloproteins is possible. We tested the viability of this pattern-generation strategy for recognition by varying the encapsulated lipophilic fluorophore (Figure 11). We found that the highest KSV value was observed for ferritin when D3 was used (2.0  106) whereas the lowest value (9.0  103) (79) Wright, A. T.; Griffin, M. J.; Zhong, Z.; McCleskey, S. C.; Anslyn, E. V.; McDevitt, J. T. Angew. Chem., Int. Ed. 2005, 44, 6375. (80) Wright, A. T.; Anslyn, E. V. Chem. Soc. Rev. 2006, 35, 14–28. (81) Ghosh, S.; Yesilyurt, V.; Savariar, E. N.; Ervine, K.; Thayumanavan, S. J. Polym. Sci., Part A: Polym. Chem. 2009, 47, 1052–1060.

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Figure 10. (a) Schematic of the differential transducer approach using amphiphilic homopolymer micelles. (b) Expected fluorescence quenching response for a hypothetical analyte.

Figure 11. (a) Structure of the different fluorescent dyes used for pattern generation (b) Plot of KSV values for different transducerprotein combinations.

was observed when myoglobin was used to quench the fluorescence of D6. It is worth noting that the observed differential fluorescence quenching may result from a mixture of variables such as the excited-state lifetime of the fluorophore, the relative energy difference between the frontier orbitals of the fluorophore and the protein cofactor responsible for fluorescence quenching, and the spectral overlap and the physical distance between the fluorophore and the protein cofactor. In addition, it should be noted that polymer-protein binding might cause a change in the protein conformation, thereby introducing additional avenues for differential fluorescence quenching responses. A complementary and attractive approach to the noncovalent fluorophore variations above involves the exploration of the utility of noncovalently assembled receptors. For instance, combining a polyelectrolyte with a complementary small-molecule surfactant provides micellar assemblies by forming a Langmuir 2009, 25(17), 9660–9670

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Figure 13. TEM images of PPA-CTAB (a) before and (b) after the addition of β-Glu. Figure 12. (a) Schematic of the assembly and disassembly upon protein binding. (b) Structures of polymers and surfactants used in the study.

supramolecular polymeric surfactant, which could be considered to be a noncovalent analog of amphiphilic homopolymers discussed earlier.81,82 These assemblies are formed at concentrations lower than the cac of the small-molecule surfactant and display container properties. Thus, at concentrations of the polymer and small-molecule surfactant between the cac of the small molecule and the polymer-small molecule complex, the guest molecule can be released upon disruption of the polymer-surfactant assembly (Figure 12). Because polyelectrolyte can competitively bind with proteins, this interaction can disrupt the assembly based on the polymer-surfactant complex at the intermediate concentrations. Thus, by encapsulating a fluorescent transducer in these micelles, polymer-protein interaction can be studied and protein sensors based on such supramolecular polymeric surfactants can be demonstrated. To test our hypothesis, we monitored the environment-dependent fluorescence emission of encapsulated pyrene in poly(potassium acrylate) (PPA) and the cetyltrimethylammonium bromide complex micelle upon the addition of five different proteins, viz., bovine serum albumin (BSA), avidin, lysozyme, chymotrypsin (ChT), and β-glucosidase ( β-Glu). We observed that the emission intensity of pyrene decreased corresponding to a 78% release upon adding β-Glu; β-Glu by itself is incapable of quenching pyrene emission. TEM images of the polymer-surfactant complex revealed that the spherical structures from the polymer-surfactant complex of about 40 nm in radius disappeared after the addition of the protein (Figure 13). Additional evidence of micelle disruption was obtained using a dynamic light scattering (DLS) experiment. The average size of the micelles decreased from 50 nm (before protein addition) to about 3 nm after the addition of protein. Some level of predictability in terms of response to various analytes is indicative of a successful design for pattern generation. However, it is also necessary to have a certain degree of unpredictability for the analyte-dependent generation of fingerprints. In our design, we observe a balance of the required predictability and unpredictability in terms of response of the entire assembly to the added protein and the analyte-dependent response of the individual components of the assembly, respectively. These analyte-dependent responses, such as the higher dye release in the case of β-Glu in spite of it being a negatively charged protein (pI = 4.4), are counterintuitive, especially when the polymer binding to the protein is also negatively charged. This can be explained by the fact that proteins are polyampholeytes and hence a negatively charged polymer is as likely to bind to a negatively charged protein as is a positively charged polymer.83 (82) Savariar, E. N.; Ghosh, S.; Gonzalez, D.; Thayumanavan, S. J. Am. Chem. Soc. 2008, 130, 5416–5417. (83) Roy, R.; Sandanaraj, B. S.; Klaikherd, A.; Thayumanavan, S. Langmuir 2006, 22, 2695–2700.

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An array of receptor combinations was achieved by varying the structure of the polymer, the structure of the surfactant, and the charge of the polymer and the surfactant. These variations were utilized to combine the analyte-dependent patterns. It is interesting that these assemblies are indeed capable of generating protein-dependent patterns for metalloproteins as well as nonmetalloproteins.

Conclusions Amphiphilic polymers are of interest in several applications in the fields of nanotechnology and biotechnology. Self-assembly of the amphiphilic block copolymers has been demonstrated in both the solid phase and solution phase. In particular, amphiphilic block copolymers have been used to form micellar assemblies that encapsulate drug molecules for drug delivery applications. External stimuli such as light, pH, temperature, and redox potential have been studied to control the release of drug molecules.6,8,84-91 However, amphiphilic homopolymers present an opportunity for polyvalent interactions arising from the preorganization of amphiphilic functionalities within a small-molecule scaffold. These polymers assemble into micelle-like structures in polar solvents and inverse-micelle-like structures in apolar solvents. The interiors of these micellar assemblies seem unique in that they provide greater site isolation and rigidity for selectivity in unimolecular photochemical reactions as compared to the interiors of small-molecule-based and block-copolymer-based surfactants.58 This property provides a unique handle for developing a new scaffold to recognize proteins by their commensurate size and ability to adapt their conformations upon binding to a surface without denaturating the protein. This reversible interaction of the binding between the polymer and the protein with structure retention offers new prospects for protein stabilization and delivery. This protein-polymer complex could be used to control protein-substrate interactions and protein-protein interactions. Similarly, polymeric nanocontainers can be used to generate a protein sensor array through pattern sensing from a single receptor using noncovalent guest molecules. The fact that this approach is orthogonal to those reported in the literature using receptor variations significantly enhances the capabilities of multianalyte recognition using pattern sensing in general. This is because the combination of the multiple transducers approach with the multiple receptors approach provides two orthogonal (84) Jiang, J.; Tong, X.; Zhao, Y. J. Am. Chem. Soc. 2005, 127, 8290–8291. (85) Zhang, Q.; Clark, C. G.Jr; Wang, M.; Remsen, E. E.; Wooley, K. L. Nano Lett. 2002, 2, 1051–1054. (86) Sundararaman, A.; Stephan, T.; Grubbs, R. B. J. Am. Chem. Soc. 2008, 130, 12264–12265. (87) Gillies, E. R.; Frechet, J. M. J. Chem. Commun. 2003, 1640–1641. (88) Apostolovic, B.; Klok, H. A. Biomacromolecules 2008, 9, 3173–3180. (89) Du, J.; Tang, Y.; Lewis, A. L.; Armes, S. P. J. Am. Chem. Soc. 2005, 127, 17982–17983. (90) Takae, S.; Miyata, K.; Oba, M.; Ishii, T.; Nishiyama, N.; Itaka, K.; Yamasaki, Y.; Koyama, H.; Kataoka, K. J. Am. Chem. Soc. 2008, 130, 6001–6009. (91) Cerritelli, S.; Velluto, D.; A., H. J. Biomacromolecules 2007, 8, 1966–1972.

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dimensions for variations in obtaining the volume of data necessary for reliable analyte-specific patterns relatively easily. Moreover, the charged hydrophilic moiety present in these assemblies can be used for the separation and detection of peptides. This opens new avenues in the areas of peptide sequencing and analysis of complex mixtures of biomolecules.

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Acknowledgment. We thank the NIGMS of the National Institutes of Health, NSF, Office of Naval Research, and the Army Research Office for supporting various aspects of the work. We are also grateful for fruitful collaborations with colleagues, mentioned in the references, which were useful in bringing about some of the utility of our amphiphilic homopolymers.

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