Optimization of Interactions between a Cationic Conjugated Polymer

J. Phys. Chem. B 2008, 112, 9295–9300

9295

Optimization of Interactions between a Cationic Conjugated Polymer and Chromophore-Labeled DNA for Optical Amplification of Fluorescent Sensors Kan-Yi Pu, Summer Yi-Hui Pan, and Bin Liu* Department of Chemical and Biomolecular Engineering, 4 Engineering DriVe 4, National UniVersity of Singapore, Singapore 117567, Singapore ReceiVed: March 5, 2008; ReVised Manuscript ReceiVed: May 5, 2008

Cationic conjugated polymers (CCPs) have been widely utilized as signal amplifiers in biosensors to improve the detection sensitivity through fluorescence resonance energy transfer (FRET) from CCPs to dye-labeled probes or targets. This paper investigates the effect of sodium dodecyl sulfate (SDS) on energy transfer between a cationic polyfluoreneethynylene copolymer (P1) and Texas Red labeled single-stranded DNA (ssDNATR). The presence of SDS in solution affects both the optical properties of P1 and TR emission within P1/ssDNA-TR complexes, which provides basic information on the role of SDS in FRET between P1 and ssDNA-TR. Although the quantum yield of P1 decreases in the presence of low concentrations of SDS, the presence of SDS reduces TR fluorescence quenching within P1/ssDNA-TR complexes and increases the number of optically active polymer repeat units within the proximity of TR, which are beneficial to P1-sensitized TR emission. In the absence of SDS, FRET from P1 to ssDNA-TR provides a 2.6-fold enhancement in TR emission intensity as compared to that upon direct excitation of TR at 595 nm. At the optimum SDS concentration (5 µM), P1-sensitized TR signal output increases to 11.3-fold relative to direct excitation of TR. This study highlights the importance of modulation of the CCP/ssDNA-dye interaction in improving the signal output of dye-labeled DNA by CCP through FRET. Introduction A unique platform for construction of chemical and biological sensors is provided by conjugated polyelectrolytes (CPEs), which are π-conjugated polymers with water-soluble charged side groups.1 Cationic conjugated polymers (CCPs) have been utilized for biomolecular detection based on the changes in their optoelectronic properties upon complexation with the target.2 Among these assays, CCP-based DNA biosensors involving fluorescence resonance energy transfer (FRET) protocols are under extensive investigation.3 These biosensors employ CCP as a light-harvesting molecule (energy donor) and the chromophore (C*) attached to PNA or DNA probes as a fluorescent signal output unit (energy acceptor).3 Amplification of C* emission signal by excitation of CCP, relative to direct excitation of C*, originates from the rapid intrachain and interchain exciton migration from CCP to C* via FRET.4 Electrostatic attractions between positively charged CCP and negatively charged phosphate groups in DNA-C* play an indispensable role in bringing CCP and DNA-C* into close proximity to satisfy the FRET distance requirement.5 Within the CCP/DNA-C* complexes, there are cross-interaction between CCP and C*, and selfinteraction among C*.6,7 The cross-interaction shortens the donor-acceptor distance and favors the FRET process; whereas self-interaction among C* causes C* fluorescence quenching, ultimately leading to a reduced signal output.6,7a It is hence of high importance to minimize acceptor fluorescence quenching upon CCP/DNA-C* complexation for improved signal output of C*.7 The signal output and the sensitivity of CCP-based DNA sensors could be improved by optimization of FRET conditions.8 Intensive efforts have been devoted to shorten the FRET * E-mail: [email protected].

distance,5b increase the spectral overlap,9a improve the conformational freedom between donor and acceptor,9b and enhance the donor quantum yield.9c However, strategies to improve signal output through reducing acceptor fluorescence quenching are less explored. Recently, we took advantage of silica-nanoparticle surface immobilized DNA-C* probes to minimize acceptor quenching upon CCP and DNA-C* complexation,9d which led to enhanced signal amplification as compared to assays conducted in homogeneous solutions. In addition, we found that a decrease in charge density of CCP could reduce acceptor fluorescence quenching within the CCP/DNA-C* complexes, which in turn improve the signal output.9e Although research activities on surfactant and CPE aqueous systems have been conducted, most of them only focused on examining how the CPE-surfactant interactions influence the optoelectronic properties of CPE.10–12 It has been pointed out that the CPE-surfactant interactions are dependent on the charge type (cationic, anionic, or nonionic) of surfactant, which in turn affect the chain conformation, aggregation, and fluorescence of CPE in different manners.10–12 Of particular interest is that introducing an oppositely charged surfactant into the CPE aqueous system can neutralize the charges of CPE.11f On the basis of our previous finding that acceptor quenching within CCP/DNA-C* complexes is dependent on the charge density of CCP, it occurs to us that utilization of an anionic surfactant to regulate the microenvironment of CCP and DNA-C* should have a positive impact on the FRET process between them. In this contribution, we demonstrate an effective and convenient strategy to improve the signal output of dye-labeled single-stranded DNA (ssDNA) sensitized by CCP. Poly[9,9′bis(6-N,N,N-trimethylammonium)-hexyl)-2,7-fluorenyleneethynylene-alt-1,4-phenylene dibromide] (P1) (Scheme 1) was synthesized and employed as the light-harvesting molecule in this study. An anionic surfactant (sodium dodecyl sulfate, SDS)

10.1021/jp8019717 CCC: $40.75  2008 American Chemical Society Published on Web 07/16/2008

9296 J. Phys. Chem. B, Vol. 112, No. 31, 2008

Pu et al.

SCHEME 1: Chemical Structure of P1

was used to interact with P1. To eliminate the interference of photoinduced charge transfer process as well as to avoid the background noise induced by the emission spectral overlap between P1 and the acceptor dye, Texas Red (TR) labeled ssDNA (ssDNA-TR: 5′-TR-ATC TTG ACT ATG TGG GTG CT-3′) was used as the probe in this study.6,7 All experiments were conducted in water to exclude the effect of buffer ions on surfactant micelle structures as well as their interactions with P1 and DNA.13 Investigation of surfactant effect on the optical properties of P1 and TR fluorescence quenching was also performed, which served as a basis to understand how surfactant influenced the FRET process between CCP and dye-labeled ssDNA. Experimental Section Methods. The NMR spectra were collected on Bruker ACF300 (300 MHz). The Element analysis was performed on Perkin-Elmer 2400 CHN/CHNS and Eurovector EA3000 Elemental Analyzers. The mass spectra were obtained using the matrix assisted laser desorption ionizationstime-of-flightsmass spectrometer (MALDI-TOF-MS) system, Autoflex II Bruker Daltronics from Bruker Daltronics. GPC analysis was conducted with a Waters 2690 liquid chromatography system equipped with Waters 996 photodiode detector and Waters 2420 evaporative light scattering detector and with three Phenogel GPC columns in one loop. The molecular weight and polydispersity were obtained using polystyrenes as the standard and THF as the eluent at a flow rate of 1.0 mL/min at 35 °C. UV-vis spectra were recorded on a Shimadzu UV-1700 spectrometer. Fluorescence measurements were carried out on a Perkin-Elmer LS55 equipped with a xenon lamp excitation source and a Hamamatsu (Japan) 928 PMT, using 90 degree angle detection for solution samples. The excitation energy at different wavelength was automatically adjusted to the same level by an excitation correction file. Quantum yields were measured using quinine sulfate as the standard, with a quantum yield of 55% in H2SO4 (0.1 M). All these detection experiments were conducted at 25 ( 1 °C in Mill-Q water (18.2 MΩ). Materials. DNA oligonucleotide (5′-TR-ATC TTG ACT ATG TGG GTG CT-3′) was purchased from Research Biolabs, Singapore. Other chemical reagents were purchased from SigmaAldrich Chemical Co. Results and Discussion Synthesis and Characterization. The water-soluble CCP was synthesized according to Scheme S1 in the Supporting Information. 2,7-Dibromo-9,9′-bis(6-bromohexyl)fluorene was reacted with trimethylsilyl acetylene in a (Ph3P)2PdCl2/CuI catalyzed Sonagashira coupling reaction. It was followed by a trimethylsilyl deprotection process in a basic solution to yield 9,9′bis(6-bromohexyl)-2,7-diethynylfluorene (2). The correct structure and purity of 2 were affirmed by NMR, mass spectrum and elemental analysis. 2 was polymerized with 1,4-diiodobenzene Via the Pd(PPh3)4/CuI catalyzed Sonagashira coupling in a mixed solvent of diisopropylamine/toluene (1: 2) to give the neutral polymer, poly[9,9′-bis(6-bromohexyl)-2,7-fluorenyleneethynylene-alt-1,4-phenylene] (P0). The number-average mo-

Figure 1. The normalized absorption and emission spectra of P1 ([RU] ) 2 µM) in water (excitation at 385 nm), and the normalized absorption spectrum of TR (1 µM) in water.

lecular weight and polydispersity of P0 are 14 000 and 2.4 respectively, determined by GPC using THF as the solvent and polystyrene as the standard. It is noteworthy that the bromohexyl side chain of 2 remained intact in the presence of diisopropylamine during polymerization. However, using triethylamine instead of diisopropylamine resulted in insoluble precipitates during the reaction, which was due to amination of the alky bromide.14 This is consistent with the previous reports, where poly(p-phenyleneethynylene)s with similar alky bromide side groups were also synthesized using diisopropylamine as the organic base.15 Treatment of P0 with trimethylamine in THF/ water yielded the water-soluble CCP, poly[9,9′-bis(6-N,N,Ntrimethylammonium)-hexyl)-2,7-fluorenyleneethynylene-alt-1,4phenylene dibromide] (P1). The chemical structure of P1 was also affirmed by NMR spectra (Figure S1 in the Supporting Inofrmation). The ratio of the integrated areas for -CH2CH2X (X ) Br, N(CH3)3) and -CH2CH2N(CH3)3 in the 1H NMR spectrum illustrates the high degree of quaternization (>95%) for P1. The absorption and emission spectra of P1 in water are shown in Figure 1. The concentration of P1 based on repeat unit (RU) is 2 µM. The absorption and emission maxima of P1 in water are located at 389 and 425 nm, respectively. The quantum yield of P1 in water is 0.22, which is comparable to poly[9,9′-bis(6N,N,N-trimethylammonium)-hexyl)-2,7-fluorene-alt-1,4-phenylene dibromide] (0.23).7a The absorption spectrum of TR is also shown in Figure 1. The overlap between P1 emission and TR absorption proves the feasibility of FRET process between P1 and TR. Surfactant Effect on Optical Properties of Donor. Changes in the absorption and emission spectra of P1 in water at [RU] ) 2 µM upon addition of SDS are shown in Figure 2. In these experiments, the SDS concentration varies from 0 to 8 µM, which is well below the critical micelle concentration (CMC) of SDS (8 mM).16 In the presence of SDS, a new absorption peak appears at 418 nm, while the emission spectra show slight red-shifts relative to that in the absence of SDS. With increased [SDS], there are intensity decreases in both the absorption and emission spectra of P1. At [SDS] ) 8 µM, the emission intensity of P1 is quenched to 28% of that in the absence of SDS. The quantum yields of P1 at different [SDS] are shown in Figure S2 in the Supporting Information. At [SDS] ) 8 µM, the quantum yield of P1 is 0.06. The spectral change and the decreased quantum yield reflect the aggregation of P1 in the presence of SDS,17,18 which stems from the reduced watersolubility of P1/SDS complexes associated with the charge neutralization process.11f The charge neutralization is caused by electrostatic attractions between oppositely charged CPE and surfactant.11f For anionic poly{1,4-phenylene[9,9-bis-(4-phe-

Optical Amplification of Fluorescent Sensors

J. Phys. Chem. B, Vol. 112, No. 31, 2008 9297

Figure 4. Emission spectra of ssDNA-TR upon addition of P1 (a) in the absence of surfactant (b) in the presence of SDS. [SDS] ) 5 µM, [ssDNA-TR] ) 20 nM (excitation at 595 nm). Figure 2. Absorption (a) and emission (b) spectra of P1 in water at [RU] ) 2 µM upon addition of SDS.

Figure 3. Stern-Volmer plot of P1 quenched by SDS (inset: the linear range of the Stern-Volmer plot).

noxybutylsulfonate)]fluorene-2,7-diyl}, upon addition of a cationic surfactant, cetyltrimethylammonium bromide, the charge neutralization process in water not only induced a decrease in spectral intensity, but also triggered the fluorenone-defect emission as a result of aggregation.11f Considering that the emission from a fluorenone defect is likely to be detected only when polymer chains are closely packed,19 this observation indicates that charge-neutralization enables compact chain aggregation. To quantify a fluorescence quenching process, the SternVolmer equation can be applied:8

I0/I ) 1 + KSV[Q]

(1)

where I0 and I are defined as the emission intensities in the absence and presence of the quencher, respectively; while [Q] is the concentration of the quencher. KSV is the Stern-Volmer constant, which provides a quantitative measure of quenching efficiency. Figure 3 shows the Stern-Volmer plot of P1 quenched by SDS. At the low [SDS] range (0 - 4 µM), a linear plot with KSV ) 8.4 × 104 M-1 is obtained. This KSV value is approximate to 2.8 × 104 M-1 for the quenching of an anionic poly(p-phenylenevinylene) by a cationic surfactant, dodecylt-

rimethylammonium bromide (DTAB).20 Since the fluorescence lifetime of P1 is ∼0.5 ns,21 the quenching rate (KSV/τ) for P1/ SDS is 1.68 × 1014 M-1s-1, which is well above the up-limit value (1010 M-1s-1) for diffusion-controlled dynamic quenching.8 As a result, the static quenching mechanism dominates the quenching process. This is in line with the fact that P1/ SDS complexation is the origin of the fluorescence quenching. For static quenching, the KSV value can be used to represent the association constant (Ka).8b The Ka between P1 and SDS is thus estimated to be 8.4 × 104 M-1. Surfactant Effect on Acceptor Quenching. To study the effect of SDS on TR fluorescence quenching, two parallel experiments were conducted by adding P1 to the ssDNA-TR aqueous solution at [ssDNA-TR] ) 20 nM in the absence and presence of SDS, respectively. With the addition of P1, changes in the intrinsic emission properties of TR in the absence and presence of SDS were monitored by direct excitation of TR at its absorption maximum (595 nm). It is noteworthy that introduction of SDS into ssDNA-TR solution prior to P1 addition cannot alter the initial PL intensity and emission maximum of ssDNA-TR. Addition of P1 to the ssDNA-TR solution in the absence of SDS results in TR fluorescence quenching, concomitant with a gradual red-shift of TR emission maximum (Figure 4a). At [RU] ) 594 nM, the TR emission intensity decreases by 86%; meanwhile the TR emission maximum is red-shifted by 6 nm from 618 to 624 nm. The red-shifted emission maximum of TR indicates that TR fluorescence quenching is caused by TR aggregation upon complexation between P1 and ssDNA-TR.7 On the contrary, the presence of [SDS] ) 5 µM in the ssDNATR solution leads to significant suppression of TR fluorescence quenching (Figure 4b). At [RU] ) 594 nM, the loss of TR emission intensity is only 13% as compared to that in the absence of SDS (86%). In addition, the TR emission maximum remains at 618 nm, indicating no significant change in the environment of TR upon P1/ssDNA-TR complexation.7 The effect of SDS concentration on the fluorescence quenching of TR by P1 is investigated using the Stern-Volmer analysis. Figure 5 shows the Stern-Volmer plots of ssDNATR quenched by P1 within the linear range, in the presence of

9298 J. Phys. Chem. B, Vol. 112, No. 31, 2008

Figure 5. Stern-Volmer plots of ssDNA-TR quenched by P1 in the presence of different SDS concentrations.

different SDS concentrations. The corresponding Stern-Volmer plots with [RU] ranging from 0 to 594 nM are shown in Figure S3 in the Supporting Information. The KSV values are calculated to be 3.0 × 106, 1.8 × 106, 6.2 × 105, 3.0 × 105, and 1.5 × 105 M-1 for [SDS] ) 0, 0.5, 1, 5, and 8 µM, respectively. The decreased quenching constant with increased SDS concentration indicates that TR fluorescence quenching by P1 is reduced at elevated SDS concentrations. Since the fluorescence lifetime of TR in water is ∼4.2 ns,22 the quenching rates (Ksv/τ) of TR in the absence of SDS and in the presence of [SDS] ) 8 µM are 7.1 × 1015 and 3.5 × 1014 M-1 s-1, respectively. These values are far above the up-limit value of a diffusion-controlled dynamic quenching.8 Hence, the static quenching mechanism is favorable for the fluorescence quenching of ssDNA-TR by P1 both in the absence and presence of SDS. Moreover, the association constant for ssDNA-TR and P1 could be roughly estimated from the quenching constant, which is ∼3.0 × 106 M-1. Acceptor fluorescence quenching is generally observed for CCP/ssDNA-C* systems as a result of the increased local dye (C*) concentration upon CCP/DNA-C* complexation.6,7,9d For the P1/ssDNA-TR system in the absence of SDS, TR fluorescence quenching is witnessed by the red-shifted TR emission maximum and the decreased TR emission intensity upon P1/ ssDNA-TR complexation. However, the presence of SDS interferes in the P1/ssDNA-TR complexation process due to the electrostatic attraction between P1 and SDS, giving rise to a reduced TR fluorescence quenching. Considering the charge neutralization process between SDS and P1 discussed in the previous section together with the reported obsevations,11f,12e the suggested mechanism is that the anionic SDS molecules interact with P1 to neutralize some of the cationic charges, resulting in a reduced charge density of P1. Previous studies on polycations and DNA interactions have shown that a reduction in polycation charge density weakens the electrostatic attraction between the polymers and DNA, which simultaneously attenuates the DNA compaction within the complexes.23,9e Accordingly, the reduced charge density of P1 by SDS could weaken the electrostatic attraction between P1 and ssDNA-TR, mitigating the ssDNA-TR compaction. In addition, the steric hindrance of SDS along the polymer chain of P1 should also contribute to more loosened P1/ssDNA-TR complex structures. The steric hindrance and the weakened electrostatic attraction could lead to an increased TR-TR distance and the hampered TR-TR contact within the complexes in comparison with that in the absence of SDS.23,7a As such, the fluorescence quenching of TR upon ssDNA-TR/P1 complexation is progressively reduced with increased [SDS] (Figure 5). On the other hand, when comparing the association constant of P1/ssDNA-TR (∼ 3.0 × 106 M-1) with that of P1/SDS (∼ 8.4 × 104 M-1), it

Pu et al.

Figure 6. TR emission intensity as a function of [RU] upon excitation of P1 at 385 nm in the absence and presence of SDS. [SDS] ) 5 µM; [ssDNA-TR] ) 20 nM. Arrow indicates Imax.

is clear that P1/ssDNA-TR complexation is favorable among ssDNA-TR, SDS and P1. This could be attributed to the higher charge density of ssDNA-TR (20 charges per molecule) than that of SDS (one charge per molecule).13 As a consequence, only when the SDS concentration (g5 µM) is far in excess of the ssDNA-TR concentration (20 nM) can the TR fluorescence quenching be significantly minimized (as shown in Figure S3b in the Supporting Information). Furthermore, the favorable interaction between P1 and ssDNA-TR in the presence of SDS ensures FRET between P1 and ssDNA-TR. Surfactant Effect on FRET. Experiments of energy transfer between P1 and ssDNA-TR were conducted in water at fixed [ssDNA-TR] ) 20 nM in the absence and presence of SDS, respectively. The ssDNA-TR emission was collected upon excitation of P1 at 385 nm where there was no significant TR absorption as shown in Figure 1. As a result, the observed TR emission is mainly due to FRET from P1. Figure 6 shows the P1-sensitized TR emission intensity as a function of [RU] in the absence of SDS and in the presence of [SDS] ) 5 µM. In the absence of SDS, initial addition of P1 leads to a continuous increase in the TR emission intensity, which is followed by saturation when [P1] is higher than 264 nM. The acceptor saturation is a frequently observed phenomenon, after which a slight decrease in the dye emission intensity appears.3,6,7,9 At [RU] ) 264 nM and [ssDNA-TR] ) 20 nM, the concentrations of positive charges and negative charges in the solution are 528 nM and 400 nM, respectively. This is consistent with previous studies that acceptor saturation usually emerges when the amount of CCP positive charges is slightly more than that of ssDNA-C* negative charges.3b,9c In the presence of [SDS] ) 5 µM, addition of P1 to the ssDNA-TR solution causes a continuous increase in TR emission intensity when [RU] is in the range of 0 to 1716 nM. At [RU] ) 1716 nM, the saturated TR emission intensity (Imax) upon excitation of P1 at 385 nm is 1141 au in the presence of 5 µM SDS, which is significantly higher than that in the absence of SDS (266 au at [RU] ) 264 nM). The delayed acceptor saturation in the presence of SDS results from the charge neutralization of P1 by SDS, which provides the feasibility for more active polymer repeat units to be brought into close proximity of TR to benefit the FRET and signal output. The emission spectra corresponding to Imax are depicted in Figure 7. The intrinsic emission spectrum of the ssDNA-TR solution (20 nM) upon direct excitation of TR at its absorption maximum (595 nm) is also shown for comparison. TR has a maximum emission peak at 618 nm with an intrinsic emission intensity of 101 au upon direct excitation. In the absence of SDS, the maximum emission peak of TR upon excitation of P1 is red-shifted by 6 to 624 nm relative to the intrinsic TR

Optical Amplification of Fluorescent Sensors

Figure 7. Emission spectra of P1/ssDNA-TR corresponding to Imax. Excitation of P1 at 385 nm. The intrinsic emission spectrum of ssDNATR at [ssDNA-TR] ) 20 nM upon direct excitation of ssDNA-TR at 595 nm is also shown.

emission (618 nm), which is indicative of the substantial change in TR environment upon P1/ssDNA-TR complexation. On the contrary, in the presence of SDS, the TR emission maximum remains at 618 nm upon excitation of P1. This indicates that the presence of SDS causes no significant change in TR environment upon P1/ssDNA-TR complexation. On the basis of the values for Imax (266 au in the absence of SDS and 1141 au in the presence of SDS) and the intrinsic TR emission intensity of 101 au, the amplification factor (the ratio of Imax to 101) is thus 2.6-fold and 11.3-fold for P1/ssDNA-TR in the absence of SDS and in the presence of 5 µM SDS, respectively. The amplification of signaling TR emission reflects the lightharvesting properties of conjugated polymers.4 In the overall FRET process, both the donor quantum yield and the acceptor fluorescence quenching can play important roles in determining the CCP-sensitized acceptor emission.9 A high quantum yield of CCP should be beneficial for FRET since donor fluorescence determines the Fo¨rster distance; whereas acceptor fluorescence quenching is an energy-wasting channel, which could reduce the CCP-sensitized acceptor emission. Despite the decreased quantum yield of P1 in the presence of SDS (Figure S2 in the Supporting Information), it could be possible that this apparent decrease in quantum yield has no inherent relationship with the P1-senstized TR emission if the total (radiative and nonradiative) rate of the excited-state recombination of P1 in the presence of SDS is slower than the rate of FRET from P1 to TR.8 Nevertheless, since the quantum yield of P1 decreases in the presence of SDS, the origin of SDSenhanced TR signal output could still be attributed to SDSreduced TR fluorescence quenching (Figure 4), and the increased number of polymer repeat units associated with ssDNA-TR (Figure 6) in the presence of SDS. Despite the favorable complexation between P1 and ssDNA-TR, the presence of high SDS concentration could more or less constrain the P1/ssDNATR complexation. This factor leads to the hampered FRET process at elevated SDS concentrations. As a result, SDS has a conflicting impact on the FRET process between P1 and ssDNATR, bringing about an optimum SDS concentration of ∼5 µM for P1-sensitzied TR emission (as shown in Figure 8). Conclusion We have demonstrated that introduction of SDS into a solution containing P1 and TR-labeled ssDNA could enhance the signal output of dye emission. Investigation of the surfactant effect on optical properties of P1 and TR fluorescence quenching indicated the conflicting effect of SDS on the donor and accepter fluorescent properties. Although the charge neutralization

J. Phys. Chem. B, Vol. 112, No. 31, 2008 9299

Figure 8. Imax as a function of [SDS].

between SDS and P1 led to decreased donor quantum yields, it suppressed the TR fluorescence quenching through reducing the acceptor aggregation within the complexes. Under optimized conditions, the amplification factor increased from 2.6 fold in the absence of SDS to 11.3-fold in the presence of 5 µM SDS. The SDS enhanced signal output of ssDNA-TR provided profound fundamental information for designing high-performance CCP-based optical sensors. It indicates that the optimization of donor-acceptor interactions within CCP/DNA-C* complexes played a vital role in improving the signal out of C*-labeled DNA. Further enhancement in overall signal amplification and sensitivity of CCP-based optical sensors could be realized with a methodology involving the concurrent virtues of enhancing the quantum yield of CCP and reducing the acceptor quenching. Acknowledgment. The authors are grateful to the National University of Singapore (NUS ARF R-279-000-197-112/133, R-279-000-233-123, and R-279-000-234-123) and the Singapore Ministry of Education (R-279-000-255-112) for financial support. Supporting Information Available: Scheme and text showing the synthesis of P1 and figures showing the NMR spectra, the quantum yields of P1 as a function of SDS concentrations, and the Stern-Volmer plots of ssDNA-TR quenched by P1. This material is available free of charge via the Internet at http:// pubs.acs.org. References and Notes (1) (a) 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. (b) Peter, K.; Nillsson, R.; Ingana¨s, O. Nat. Mater. 2003, 2, 419. (c) Ho, H.; Leclerc, M. J. Am. Chem. Soc. 2004, 126, 1384. (d) Tang, Y.; Feng, F.; He, F.; Wang, S.; Li, Y.; Zhu, D. J. Am. Chem. Soc. 2006, 128, 14972. (e) Li, C.; Numata, M.; Takeuchi, M.; Shinkai, S. Angew. Chem., Int. Ed. 2005, 44, 6371. (f) Haskins-Glusac, K.; Pinto, M. R.; Tan, C.; Schanze, K. S. J. Am. Chem. Soc. 2004, 126, 14964. (g) Wosnick, J. H.; Mello, C. M.; Swager, T. M. J. Am. Chem. Soc. 2005, 127, 3400. (h) Pinto, M. R.; Schanze, K. S. Proc. Nat. Acad. Sci. USA, 2004, 101, 7505. (i) Miranda, O. R.; You, C. C.; Phillips, R.; Kim, I. B.; Ghosh, P. S.; Bunz, U. H. F.; Rotello, V. M. J. Am. Chem. Soc. 2007, 129, 9856. (2) (a) Achyuthan, K. E.; Bergstedt, T. S.; Chen, L.; Jones, R. M.; Kumaraswamy, S.; Kushon, S. A.; Ley, K. D.; Lu, L.; McBranch, D.; Mukundan, H.; Rininsland, F.; Shi, X.; Xia, W.; Whitten, D. G. J. Mater. Chem. 2005, 15, 2648. (b) Ho, H. A.; Be´ra-Abe´rem, M.; Leclerc, M. Chem. Eur. J. 2005, 11, 1718. (c) Peter, K.; Nilsson, R.; Rydberg, J.; Baltzer, L.; Ingana¨s, O. Proc. Nat. Acad. Sci. U.S.A. 2003, 100, 10170. (d) Lee, K.; Povlich, L. K.; Kim, J. AdV. Funct. Mater. 2007, 17, 2580. (3) (a) Gaylord, B. S.; Heeger, A. J.; Bazan, G. C. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 10954. (b) Gaylord, B. S.; Heeger, A. J.; Bazan, G. C. J. Am. Chem. Soc. 2003, 125, 896. (c) Liu, B.; Bazan, G. C. Chem. Mater. 2004, 16, 4467. (d) Liu, B.; Bazan, G. C. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 589. (4) (a) McQuade, D. T.; Pullen, A. E.; Swager, T. M. Chem. ReV. 2000, 100, 2537. (b) Thomas, S. W., III; Joly, G. D.; Swager, T. M. Chem. ReV. 2007, 107, 1339.

9300 J. Phys. Chem. B, Vol. 112, No. 31, 2008 (5) (a) Liu, B.; Gaylord, B. S.; Wang, S.; Bazan, G. C. J. Am. Chem. Soc. 2003, 125, 6705. (b) Wang, S.; Liu, B.; Gaylord, B. S.; Bazan, G. C. AdV. Funct. Mater. 2003, 13, 463. (c) Xu, Q. H.; Gaylord, B. S.; Wang, S.; Bazan, G. C.; Moses, D.; Heeger, A. J. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 11634. (6) Bazan, G. C. J. Org. Chem. 2007, 72, 8615. (7) (a) Liu, B.; Bazan, G. C. J. Am. Chem. Soc. 2006, 128, 1188. (b) Woo, H. Y.; Vak, D.; korystov, D.; Mikhailovsky, A.; Bazan, G. C.; Kim, D. Y. AdV. Funct. Mater. 2007, 17, 290. (8) (a) Lakowicz, J. R. Principles of Fluorescence Spectroscopy; Kluwer Academic/Plenum: New York, 1999. (b) Tan, C. Y.; Atas, E.; Mu¨ller, J. G.; Pinto, M. R.; Kleiman, V. D.; Schanze, K. S. J. Am. Chem. Soc. 2004, 126, 13685. (9) (a) Liu, B.; Wang, S.; Bazan, G. C.; Mikhailovsky, A. J. Am. Soc. Chem. 2003, 125, 13306. (b) Liu, B.; Bazan, G. C. J. Am. Chem. Soc. 2004, 126, 1942. (c) Liu, B.; Bazan, G. C. Chem. Asian J. 2007, 2, 499. (d) Wang, Y.; Liu, B. Chem. Commun. 2007, 3553. (e) Pu, K. Y.; Fang, Z.; Liu, B. AdV. Funct. Mater. 2008, 18, 1321. (10) (a) Chen, L.; Xu, S.; McBranch, D.; Whitten, D. J. Am. Chem. Soc. 2000, 122, 9302. (b) Chen, L.; McBranch, D.; Wang, R.; Whitten, D. Chem. Phys. Lett. 2000, 330, 27. (c) Abe, S.; Chen, L. J. Polym. Sci., Part B: Phys. 2003, 41, 1676. (11) (a) Burrows, H. D.; Lobo, V. M. M.; Pina, J.; Ramos, M. L.; Seixas de Melo, J.; Valente, A. J. M.; Tapia, M. J.; Pradhan, S.; Scherf, U. Macromolecules 2004, 37, 7425. (b) Tapia, M. J.; Burrows, H. D.; Knaapila, M.; Monkman, A. P.; Arroyo, A.; Pradhan, S.; Scherf, U.; Pinazo, A.; Perez, L.; Moran, C. Langmuir 2006, 22, 10170. (c) Knaapila, M.; Almasy, L.; Garamus, V. M.; Pearson, C.; Pradhan, S.; Petty, M. C.; Scherf, U.; Burrows, H. D.; Monkman, A. P. J. Phys. Chem. B 2006, 110, 10248. (d) Burrows, H. D.; Tapia, M. J.; Silva, C. L.; Pais, A. A. C. C.; Fonseca, S. M.; Pina, J.; Seixas de Melo, J.; Wang, Y.; Marques, E. F.; Knaapila, M.; Monkman, A. P.; Garamus, V. M.; Pradhan, S.; Scherf, U. J. Phys. Chem. B 2007, 111, 4401. (e) Monteserin, M.; Burrows, H. D.; Valente, A. J. M.; Lobo, V. M. M.; Mallavia, R.; Tapia, M. J.; Garcia-Zubiri, I. X.; Di Paolo, R. E.; Macanita, A. L. J. Phys. Chem. B 2007, 111, 13560. (f) Tapia, M. J.; Burrows, H. D.; Valente, A. J. M.; Pradhan, S.; Scherf, U.; Lobo, V. M. M.; Pina, J.; Seixas de Melo, J. J. Phys. Chem. B 2005, 109, 19108. (12) (a) Dalvi-Malhotra, J.; Chen, L. J. Phys. Chem. B 2005, 109, 3873. (b) Lavigne, J. J.; Broughton, D. L.; Wilson, J. N.; Erdogan, B.; Bunz, U. H. F. Macromolecules 2003, 36, 7409. (c) Sholin, V.; Lopez-Cabarcos, E. J.; Carter, S. A. Macromolecules 2006, 39, 5830. (d) George, W. N.; Giles, M.; McCulloch, I.; de Mello, J. C.; Steinke, H. G. J. Soft Matter 2007, 3, 1381. (e) Al Attar, H. A.; Monkman, A. P. J. Phys. Chem. B 2007, 111, 12418. (13) Cooper, C. L.; Dubin, P. L.; Kayitmazer, A. B.; Turksen, S. Curr. Opin. Colloid Interface Sci. 2005, 10, 52. (14) (a) Tong, H.; Hong, Y.; Dong, Y.; Haeussler, M.; Lam, J. W. Y.; Li, Z.; Guo, Z.; Guo, Z.; Tang, B. Z. Chem. Commun. 2006, 3705. (b)

Pu et al. Semenov, V. E.; Voloshina, A. D.; Toroptzova, E. M.; Kulik, N. V.; Zobov, V. V.; Giniyatullin, R. K.; Mikhailov, A. S.; Nikolaev, A. E.; Akamsin, V. D.; Reznik, V. S. Eur. J. Med. Chem. 2006, 41, 1093. (c) Ono, Y.; Shikata, T. J. Phy. Chem. B 2006, 110, 9426. (d) Liu, X.-M.; Yang, B.; Wang, Y.; Wang, J.-Y. Chem. Mater. 2005, 17, 2792. (15) (a) Swager, T. M.; Gil, C. J.; Wrighton, M. S. J. Phys. Chem. 1995, 99, 4886. (b) Xue, C.; Chen, Z.; Luo, F.-T.; Palaniappan, K.; Chesney, D. J.; Liu, J.; Chen, J.; Liu, H. Biomacromolecules 2005, 6, 1810. (16) Holmberg, K.; Jonsson, B.; Kronberg, B.; Lindman, B. Surfactants and Polymers in Aqueous Solution, 2nd ed.; Wiley: Chichester, U.K., 2003. (17) (a) Schwartz, B. J. Annu. ReV. Phys. Chem. 2003, 54, 141. (b) Chu, Q.; Pang, Y. Macromolecules 2005, 38, 517. (c) Nguyen, T. Q.; Schwartz, B. J. J. Chem. Phys. 2002, 116, 8198. (d) Peng, K. Y.; Chen, S. A.; Fann, W. S. J. Am. Chem. Soc. 2001, 123, 11388. (e) Zahn, S.; Swager, T. M. Angew. Chem., Int. Ed. 2002, 41, 4225. (18) (a) Chen, S. H.; Su, C. H.; Su, A. C.; Chen, S. A. J. Phys. Chem. B 2004, 108, 8855. (b) Chen, S. H.; Su, A. C.; Chang, C. S.; Chen, H. L.; Ho, D. L.; Tsao, C. S.; Peng, K. Y.; Chen, S. A. Langmuir 2004, 20, 8909. (19) (a) Zojer, E.; Pogantsch, A.; Hennebicq, E.; Bre´das, J. -L.; Scandiucci de Freitas, P.; Scherf, U.; List, E. J. W. J. Chem. Phys. 2002, 117, 6794. (b) Scherf, U.; List, E. J. W. AdV. Mater. 2002, 14, 477. (c) Romaner, L.; Pogantsch, A.; Scandiucci de Freitas, P.; Scherf, U.; Gaal, M.; Zojer, E.; List, E. J. W. AdV. Funct. Mater. 2003, 13, 597. (d) Kulkarni, A. P.; Kong, X.; Jenekhe, S. A. J. Phys. Chem. B 2004, 108, 8689. (e) Becker, K.; Lupton, J. M.; Feldmann, J.; Nehls, B. S.; Galbrecht, F.; Gao, D. Q.; Scherf, U. AdV. Funct. Mater. 2006, 16, 364. (f) Montilla, F.; Mallavia, R. AdV. Funct. Mater. 2007, 17, 71. (20) Gu, Z.; Bao, Y. J.; Zhang, Y.; Wang, M.; Shen, Q. D. Macromolecules 2006, 3125. (21) Huang, Y. Q.; Fan, Q. L.; Zhang, G. W.; Chen, Y.; Lu, X. M.; Huang, W. Polymer 2006, 47, 5233. (22) Brismar, H.; Trepte, O.; Ulfhake, B. J. Histochem. Cytochem. 1995, 43, 699. (23) (a) Fischer, D.; Dautzenberg, H.; Kunath, K.; Kissel, T. Int. J. Pharm. 2004, 280, 253. (b) Howard, K. A.; Dash, P. R.; Read, M. L.; Ward, K.; Tomkins, L. M.; Nazarova, O.; Ulbrich, K.; Seymour, L. W. Biochim. Biophys. Acta 2000, 1475, 245. (c) Strand, S. P.; Danielsen, S.; Christensen, B. E.; Va˚rum, K. M. Biomacromolecules 2005, 6, 3357. (d) Liu, X.; Yang, J. W.; Miller, A. D.; Nack, E. A.; Lynn, D. M. Macromolecules 2005, 38, 7907. (e) Rungsardthong, U.; Ehtezazi, T.; Bailey, L.; Armes, S. P.; Garnett, M. C.; Stolnik, S. Biomacromolecules 2003, 4, 683. (f) Furgeson, D. Y.; Chan, W. S.; Yockman, J. W.; Kim, S. W. Bioconjugate Chem. 2003, 14, 840. (g) Erbacher, P.; Roche, A. C.; Monsigny, M.; Midoux, P. Biochim. Biophys. Acta 1997, 1324, 27. (h) Wang, Y.; Dubin, P. L.; Zhang, H. Langmuir 2001, 17, 1670.

JP8019717