Cy3 in AOT Reverse Micelles I. Dimer Formation Revealed through

ARTICLE pubs.acs.org/JPCB

Cy3 in AOT Reverse Micelles I. Dimer Formation Revealed through Steady-State and Time-Resolved Spectroscopy Jeffrey T. McPhee, Eric Scott, Nancy E. Levinger,* and Alan Van Orden* Department of Chemistry, Colorado State University, Fort Collins, Colorado 80523, United States ABSTRACT: Cyanine-3 (Cy3) fluorescent dye molecules confined in sodium di-2-ethylhexyl sulfosuccinate (AOT) reverse micelles were examined using steady-state absorption and emission as well as time-resolved fluorescence spectroscopy to understand the effect of confinement on the spectroscopic properties of the dye. This study explored a wide range of reverse micelle sizes, with hydrodynamic radii ranging from ∼1.7 to ∼5 nm. The relative concentrations of Cy3 and AOT reverse micelles were such that, on average, one dye molecule was present for every 2 104 to 9 105 reverse micelles. In the smallest reverse micelles examined, observed changes in the absorption and emission spectra and fluorescence lifetime of the dye molecules indicated H-aggregation of Cy3 into side-by-side dimers. It is hypothesized that this dimerization is governed by the high local concentrations that result from the confinement of the Cy3 in the reverse micelles. What is notable about this study is that this dimer occurs even at overall dye concentrations in the nanomolar range. Such concentrations are too low for aggregation to occur in bulk solution. Hence, the reverse micelles serve as nanocatalysts for this aggregation process.

’ INTRODUCTION The sensitivity and versatility of fluorescence has made it an exceptionally effective tool for a wide range of studies. Fluorescence spectroscopies are especially effective for studies of biological processes.13 For example, by adding a fluorescent tag to a biomolecule, researchers can follow molecular recognition events,4 macromolecular folding,58 or intermolecular interaction.9 As we introduce these fluorophores into increasingly confined environments in biological samples, understanding the effect of confinement on their basic photophysics becomes critical. One model system providing significant and variable confinement is the reverse micelle, shown in Figure 1.10,11 Consisting of three different phases, these structures enable studies of the effects of spatial and orientational confinement on molecular processes.1215 For example, at the appropriate concentrations, sodium di-2ethylhexyl sulfosuccinate (AOT, structure shown in Figure 1) forms reverse micelles that consist of a water pool, an interfacial region created by the polar head groups and aliphatic tails of the surfactant, and the nonpolar solvent. The size of AOT reverse micelles is characterized by the parameter w010,11 w0 ¼

½H2 O ½AOT

ð1Þ

where [AOT] and [H2O] are the molar concentrations of each species in the solution. For AOT reverse micelles, w0 is directly proportional to the hydrodynamic radius, RH (nm) = 0.175w0 + 1.5,16 facilitating precise control of the particle size of these r 2011 American Chemical Society

systems and making them attractive for studying the role of confined local environment on various processes, such as catalysis,1720 intermolecular charge transfer,21 and redox reactions.22 Here, we present a study that investigates the role of confinement in AOT reverse micelles on the behavior of a water-soluble organic cyanine dye, i.e., Cy3, shown in Figure 1. The general class of cyanine dyes has a rich history of investigation in homogeneous solutions, including many studies on their spectroscopic properties.23,24 Most cyanine dyes display absorption bands in the visible spectrum consisting of a shoulder followed by an absorbance maximum.2326 Generally speaking, cyanine dyes exist as monomers at lower concentrations (submicromolar) in aqueous solutions. However, as the concentration of the dye increases or the solvent environment changes, these dyes can form dimers and higher order aggregates that alter the spectroscopic properties of the dye. These aggregation processes have been studied extensively in bulk solution.2740 In aqueous solution, cyanine dyes form both J- and H-aggregates, leading to shifts in their absorption maxima. J-aggregates arise from an end-to-end or slipped geometry of monomers, as shown in Figure 1, displaying a red-shifted absorption spectrum, while H-aggregates, characterized by a side-by-side arrangement of molecule display a blue shift in the absorption spectrum, relative to the monomer peak, and generally exhibit weak fluorescence in Received: January 5, 2011 Revised: June 15, 2011 Published: July 18, 2011 9576

dx.doi.org/10.1021/jp200126f | J. Phys. Chem. B 2011, 115, 9576–9584

The Journal of Physical Chemistry B

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Figure 1. Molecular structures for Cy3 dye and AOT surfactant used for experiments as well as a model structure for an AOT reverse micelle. This figure also shows the configuration adopted for dimers of H-aggregates or J-aggregates.

solution compared to the monomer.37 However, instances of highly fluorescent H-aggregated cyanine dyes have been reported, especially in confined environments such as LangmuirBlodgett layers, at low temperatures, and in rigid environments.36,37,4145 The enhanced emission most likely occurs because the rigid environment prevents rapid internal conversion that would quench the emission. For the purposes of this paper, we define an H-dimer as a pair of dye molecules in a side-by-side geometry as shown in Figure 1. Cy3 has two forms, a fluorescent trans-isomer and its nonfluorescent cis-isomer.4648 The transcis photoisomerization reaction has been studied extensively by a variety of techniques.25,4756 The Cy3 absorption maximum appears at 550 nm with a corresponding emission maximum at 570 nm.4648 In aqueous solution, the dye exists in its monomer form at concentrations below 1 mM. Cy3 exhibits a short fluorescence lifetime in aqueous solution, typically about 0.15 ns and has a low quantum yield (0.05).4648 The short lifetime and corresponding low quantum yield is thought to arise from the transcis isomerization reaction that the dye undergoes, which provides an efficient nonradiative decay pathway to compete with the radiative decay.4648 Studies of the cis trans isomerization reaction show its sensitivity to changes in the solvent viscosity as well as weak influence of solvent polarity.48 Our working hypothesis that motivates the present study is that the confined environment within the reverse micelle can alter the kinetics and mechanism of transcis isomerization and aggregation of Cy3. Thus, introducing Cy3 into reverse micelles can probe the effects of confinement on such processes. Our results show an interesting, unexpected behavior of aqueous Cy3 inside AOT/isooctane reverse micelles. The confined environment of the reverse micelles drives the formation of Cy3 H-dimers at highly dilute concentrations. Such dimerization would not occur in bulk water when the Cy3 molecules are at equal concentrations. Furthermore, the aggregation occurs predominantly in the smallest sized reverse micelles, w0=1. In larger reverse micelles, the Cy3 dye exists primarily in the monomer form. Here, we present a study of this

aggregation phenomenon using a variety of spectroscopic techniques including fluorescence emission, time-correlated single photon counting (TCSPC) and absorption spectroscopy measurements.

’ EXPERIMENTAL METHODS Sample Preparation. Cy3 monoreactive dye pack (GE Life Science (Piscataway, NJ) and isooctane (99.8%, Sigma-Aldrich) were used as received. AOT (sodium di-2-ethylhexyl sulfosuccinate, 99%, Sigma-Aldrich) was purified using a process described elsewhere.57 For sample preparation, the solid dye was dissolved in deionized Millipore water (18.2 MΩ 3 cm) and the resulting solution stored under refrigeration. Aqueous Cy3 solutions were prepared with a concentration range from 9.9 105 to 9.9 106 M. Cy3 showed no solubility in the pure isooctane nonpolar phase in the absence of AOT. Reverse micelles were prepared by dissolving AOT in isooctane to form a 0.3 M stock solution to which aqueous dye solution and water were added. All samples were prepared using 2 mL of a 0.3 M AOT in isooctane solution and 10.8 μL of aqueous Cy3 solution. The size (w0) and concentration of the reverse micelles were adjusted by adding an appropriate amount of isooctane and water, totaling 1 mL, resulting in a reverse micelle sample with an overall AOT concentration of 0.2 M. For the experiments reported here, the AOT concentration in the final solution was 0.02 M, unless otherwise noted. To make these final samples, a 100 μL aliquot of a 0.2 M AOT sample was diluted into 900 μL of isooctane. The final overall dye concentration in the solutions ranged from 3.6 108 to 8.9 1010 M. Using data in the literature,58 we estimate that in the most concentrated samples there is at most one Cy3 molecule per 20 000 reverse micelles; at the lowest dye concentration, there is fewer than one dye molecule per 900 000 reverse micelles. Steady-State and Time-Resolved Measurements. UVvis absorption spectra of Cy3 in different-sized reverse micelles were measured using a Cary 500 UV/vis-NIR spectrophotometer (Varian Inc.). Spectra were collected using a 1 cm path length 9577

dx.doi.org/10.1021/jp200126f |J. Phys. Chem. B 2011, 115, 9576–9584

The Journal of Physical Chemistry B cuvette referenced to the spectrum of neat isooctane in the sample cuvette. Each sample was scanned a minimum of three times and the resulting spectra were averaged. Cy3 concentrations in samples used for emission experiments are too low to be measured by absorption spectroscopy. For absorption spectroscopy measurements, the overall Cy3 concentration was 3.6 107 M Cy3, approximately 1 order of magnitude greater than the concentrations used in the fluorescence spectroscopy experiments, accomplished by increasing the overall AOT concentration to 0.2 M AOT. All other experiments were performed using the sample preparation procedure described previously. Steady-state Cy3 emission spectra and excitation spectra were collected using a SPEX steady-state spectrofluorometer (HORIBA Jobin Yvon) using a 1 cm path length cuvette. Emission spectra were collected with excitation at 514 and 532 nm, to capture emission characteristics of various features observed in absorption spectra. Excitation spectra were measured by scanning the excitation wavelength while monitoring emission intensity at 561, 600, and 650 nm. Emission lifetimes were measured via time-correlated single photon counting using a Fluorocube instrument (HORIBA Jobin Yvon) to monitor changes in the Cy3 emission lifetime due to the surrounding environment. The instrument response function was measured and the sample was subsequently loaded into the instrument. All samples were excited at a 514 nm wavelength of an argon flash lamp selected by a monochromator. Subsequent emission was detected at 560, 600, and 650 nm. Lifetimes on the 110 ns time scale indicate fluorescence rather than phosphorescence emission from the samples. Total fluorescence emission from the Cy3 in w0 = 1 reverse micelles was collected using a Nikon Eclipse TE-2000-U microscope (Nikon Inc., Melville) irradiated with a CW laser beam at 514 nm (Ar ion laser, Melles-Griot) using a 100 1.3 numerical aperture microscope objective. The laser power was 0.05 mW, before entering the microscope. The sample was introduced on the microscope stage in a well-slide covered with a microscope cover slide. The resulting fluorescence was transmitted back through the microscope and imaged onto a 50 μm confocal pinhole. The resulting light was long-pass filtered (535 nm LP filters, Thorlabs Inc.) and focused onto a single-photon-counting avalanche photodiode detector (PerkinElmer Inc.). The resulting output of the detector was recorded using one channel of an ALV-5000E/EPP card (ALV-GmbH) interfaced to a computer.

’ RESULTS Figure 2 displays the normalized absorption spectrum of 3.6 107 M Cy3 in water and in reverse micelles of varying size. In pure water, the Cy3 absorption spectrum matches reports in the literature46 displaying a peak at 550 nm and a shoulder at 514 nm. The Cy3 absorption spectrum shown in Figure 2 for the largest reverse micelle studied, w0 = 30, is almost identical to that of Cy3 in water. This suggests the Cy3 dye in w0 = 30 reverse micelles experiences an environment similar to that of bulk water, and is consistent with previous reports showing that water in reverse micelles assumes the properties of bulk water for w0 g 15.14 However, as the reverse micelle environment becomes more confined from w0 = 15 to 2, the peak at 550 nm gradually shifts to longer wavelength. Concomitantly, the shoulder at 514 nm gradually becomes more intense relative to the main peak, forming a second peak for the smallest reverse micelle sizes.

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Figure 2. Normalized absorption spectra of Cy3 in a variety of environments: bulk water (black) and five different reverse micelle sizes, w0 = 1 (blue), w0 = 2 (pink), w0 = 9 (turquoise), w0 = 15 (purple), and w0 = 30 (red). The concentration of Cy3 in the water sample was 9.9 105 M. In the five reverse micelle samples, the overall Cy3 concentration is held constant at 3.6 107 M. The concentration of AOT in the five reverse micelle samples is 0.2 M.

These changes in the absorption spectra suggest that the confined environment of the reverse micelles alters the spectroscopic properties of the dye. Possible sources of this perturbation include confinement effects on the water molecules surrounding the dye, interactions of the dye with the polar head groups in the interior walls of the reverse micelles, or interactions of the dye with the nonpolar tailgroups, but the absorption spectra do not distinguish between these possibilities. The most dramatic change occurs in the w0 = 1 sample, in which the primary spectral feature peaks at the longest wavelength of all the samples, and the relative intensity of the peak at 514 nm becomes equivalent to that of the primary peak. Indeed, there appears to be a qualitative difference in the spectra of dyes confined in w0 = 1 reverse micelles, compared to those confined in w0 = 230 reverse micelles and bulk water. Whereas confinement in w0 = 230 reverse micelles gives rise to gradual changes in the spectra with decreasing micelle size, the changes that occur in the w0 = 1 are much more dramatic. This suggests that, in addition to the dyereverse micelle interactions evident in all the reverse micelle samples, the Cy3 in w0 = 1 reverse micelles may be present in a different form than in the larger w0 = 230 micelles. Figure 3 shows fluorescence emission spectra obtained from samples containing 3.6 108 M Cy3 in bulk water and in reverse micelles of varying size. Spectra displayed in Figure 3a were excited at 532 nm while those in Figure 3b were excited at 514 nm. We chose these wavelengths to selectively excite the two prominent peaks observed in the absorption spectra (Figure 2) and to match the laser excitation wavelengths used in our fluorescence correlation spectroscopy experiments described in the accompanying paper.59 The fluorescence emission spectra of Cy3 in bulk water agree well with the known properties of the dye, with an intense maximum at ∼560 nm and a shoulder at ∼600 nm.46,47,60,61 The emission spectra shift to longer wavelength with respect to the bulk water spectrum as the micelles become smaller, consistent with the behavior observed in the absorption spectra. For 532 nm excitation (Figure 3a), these changes occur gradually within the w0 = 230 size range, while the qualitative shapes of the spectra remain relatively constant throughout this range. A larger red shift and significant broadening of the spectrum is observed for Cy3 in w0 = 1 reverse micelles, but the qualitative shape of the emission spectrum is similar to that of Cy3 in w0 = 230 reverse micelles and in bulk water. 9578



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Figure 4. Normalized fluorescence emission spectra of Cy3 in a 0.02 M AOT w0 = 1 reverse micelle sample at different overall concentrations of Cy3. From highest to lowest concentration, the five concentrations analyzed were 3.6 108 M (red), 1.8 108 M (blue), 8.9 109 M (green), 3.6 109 M (black), and 1.8 109 M (pink).

to a sample of Rhodamine 6G (R6G) in methanol (quantum yield ≈0.9562) using Figure 3. Normalized fluorescence emission spectra of Cy3 in different environments and at two different excitation wavelengths, 532 nm (a) and 514 nm (b). The different environments are as follows: bulk water (black) and five different reverse micelle sizes, w0 = 1 (blue), w0 = 2 (pink), w0 = 9 (turquoise), w0 = 15 (purple), and w0 = 30 (red). Inset in the bottom panel shows the overall fluorescence intensity in each of the six samples, calculated from the area under the unnormalized emission curves. The concentration of Cy3 in all samples is 3.6 108 M and the concentration of AOT in the reverse micelle samples is 0.02 M.

Similar trends are observed in the 514 nm excited spectra for Cy3 in w0 = 930 reverse micelles—namely, a gradual shift to the red with decreasing micelle size. However, the changes observed in the smaller sized reverse micelles are much more pronounced than those observed in the spectra excited at 532 nm. In the w0 = 2 sample, the shoulder at ∼600 nm becomes more intense relative to the primary peak near 560 nm, and a new feature at ∼640 nm begins to appear. The w0 = 1 sample exhibits fluorescence behavior that fundamentally differs from all the other samples. The main peak near 560 nm is suppressed relative to the peaks at ∼600 and ∼640 nm, which dominate the spectrum. This provides further evidence that the Cy3 molecules exist in a different form when confined in w0 = 1 reverse micelles compared to larger micelles and bulk water. Also, the spectral differences observed for Cy3 in w0 = 1 reverse micelles excited at 514 and 532 nm suggest that the features being excited at these wavelengths represent different forms of the dye present in the reverse micelles. Another important difference between Cy3 in w0 = 1 reverse micelles compared to the larger reverse micelles and bulk water comes from the dramatic increase in total fluorescence intensity observed in the w0 = 1 sample. This is demonstrated in the inset to Figure 3b, which shows approximately a 4-fold increase in emission intensity for Cy3 in w0 = 1 reverse micelles excited at 514 nm. To investigate the observed change in the emission intensity, we measured the fluorescence quantum yield of Cy3 in 0.2 M w0 = 1 reverse micelles at 514 and 532 nm and compared this to the fluorescence quantum yield in w0 = 30 reverse micelles and bulk water. We calculated the quantum yield by referencing

ϕCy3 ¼ ϕR6G

FCy3 AR6G FR6G ACy3

ð2Þ

where, ϕCy3 and ϕR6G are the quantum yields, FCy3 and FR6G are the relative fluorescence intensities and ACy3 and AR6G are the absorbance values for Cy3 and R6G, respectively. The values for the relative fluorescence intensities were calculated from the area under the fluorescence emission spectra, and the values for the absorbance were obtained from the absorption spectra for the respective wavelengths. In these experiments, we used an AOT concentration of 0.2 M to facilitate the absorbance measurements. Interestingly, the quantum yield for Cy3 in w0 = 1 reverse micelles excited at 514 nm is ∼0.3 and at 532 nm is ∼0.5, both of which represent a dramatic increase from the quantum yield of aqueous Cy3 (∼0.05).4648 Moreover, in the w0 = 30 reverse micelle sample we found a quantum yield of ∼0.09 at both 514 and 532 nm excitation. The low quantum yield of Cy3 in bulk aqueous solution has been attributed to photoinduced trans cis isomerization, which suppresses the fluorescence by nonradiative energy transfer to the nonfluorescent cis-isomer of the dye.17,4648 The similarly low quantum yield of Cy3 in w0 = 30 reverse micelles suggests dyes confined in these micelles sense a similar environment to bulk water. The dramatic enhancement of the quantum yield of Cy3 in w0 = 1 reverse micelles provides further evidence for a new form of the dye when confined in the smallest reverse micelles. We hypothesize that the spectral changes discussed above for Cy3 in w0 = 1 are consistent with the formation of Cy3 H-dimers in these reverse micelles, whereas the dyes in w0 = 230 reverse micelles and in bulk water exist predominantly as isolated monomers. We considered the possibility that the species probed in the w0 = 1 reverse micelles could be J-dimers instead of H-dimers. However, J-dimers absorb at longer wavelengths than Cy3 monomers,28,31,39,40 whereas our absorption spectra show an increased absorbance at shorter wavelength compared to the monomer peak, consistent with H-aggregation.30,3335,37,38 Furthermore, the emission spectra show that the new spectral features arising in smaller reverse micelles depend on the excitation wavelength. The spectral changes are much more dramatic when the excitation wavelength is 514 nm. With 532 nm excitation, the 9579



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Figure 5. Normalized fluorescence emission spectra of Cy3 in w0 = 1 reverse micelles as a function of AOT concentration and excitation wavelength, 532 nm (top) and 514 nm (bottom). The overall Cy3 concentration in all samples is 1.8 108 M. The AOT concentration for each spectrum is as follows: 0.01 M (red), 0.02 M (blue), 0.04 M (pink), 0.06 M (purple), 0.08 M (turquoise), 0.1 M (green), and 0.2 M (black).

spectrum is qualitatively similar to the other spectra. This suggests that both H-dimers and monomers of Cy3 coexist in the w0 = 1 samples and that 514 nm selectively excites the H-dimer form of the dye, while 532 nm excitation is more selective of the monomer. The fact the fluorescence quantum yields observed using both 514 and 532 nm excitation are enhanced relative to larger reverse micelles and bulk water indicates both the H-dimer and monomer forms of the dye undergo significant changes in their photophysical properties when confined in w0 = 1 reverse micelles. In the case of H-dimers, this observation is consistent with previous studies showing enhanced fluorescence emission in confined environments relative to unconfined monomers.37 In the case of the monomer Cy3 molecules confined in w0 = 1 reverse micelles, we suggest the photoisomerization process responsible for the low fluorescence quantum yield in bulk samples is suppressed under these highly confined conditions. Indeed, this effect of confinement on the monomeric form of the dye may be present even in the largest w0 = 30 reverse micelles due to the slight increase in quantum yield noted above. To further investigate whether the observed spectral features in w0 = 1 reverse micelles could be caused by Cy3 dimerization, we examined the fluorescence emission properties of the Cy3 as a function of overall Cy3 concentration. Spectral features arising from dimers should be more pronounced at higher concentrations. Spectra of Cy3 in w0 = 1 reverse micelles for different Cy3 concentrations are shown in Figure 4. The AOT concentration was held constant at 0.02 M. At the lowest Cy3 dye concentration,

the spectral features follow a similar trend to those seen for Cy3 in water and in larger reverse micelles, that is, the peak near 560 nm dominates the spectrum, and the features at 600 and 650 nm are much weaker compared to the peak at 560 nm. As the concentration increases, the emission maximum shifts to the feature at 600 nm and the relative intensity of the 560 nm peak diminishes. Also, the feature at 650 nm becomes increasingly prominent. Thus, as the concentration of Cy3 increases, the emission spectrum increasingly diverges from that seen in water and the larger micelles, indicating the presence of a new species at higher concentrations. Our conclusion from these concentration dependence experiments is that within w0 = 1 reverse micelles samples, the Cy3 molecules exist in both monomer and H-dimer form and that the equilibrium shifts toward H-dimers at higher concentrations. This is accompanied by a shift in the emission maximum from 560 to 600 nm, suggesting the monomer emission occurs at 560 nm and the dimer emission occurs at the longer wavelengths. To further investigate this aggregation process, we examined the effect of reverse micelle concentration on the fluorescence emission spectra. If the observed spectral features of Cy3 in w0 = 1 reverse micelles arise due to H-dimerization, then the spectra should also depend on the overall AOT concentration. As the AOT concentration increases, the number of reverse micelles increases. With more reverse micelles in a sample, the probability of at least two Cy3 molecules coming in contact decreases and thus we expect to see less aggregation behavior as the concentration 9580



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Table 1. Fluorescence Lifetime Data of Cy3 in Three Different Environments: Water, w0 = 30, and w0 = 1a sample H2O w0 = 30 w0 = 1

T1 (% intensity) (ns)

Table 2. Fluorescence Lifetime Data of Cy3 in w0 = 1a


0.3 (100)

N/A

0.3 (45) 1.8 (84)

3.0 (55) 6.9 (14)

a

The samples were excited at 514 nm and the emission was collected at 561 nm. The concentration of Cy3 is 3.6 108 M and the concentration of AOT in the reverse micelle sample is 0.02 M. The errors in the reported lifetimes are ∼0.1 ns.

Figure 6. Fluorescence lifetime data of Cy3 inside a w0 = 1 reverse micelle excited at 514 nm and collected at 561 nm (red) and 600 nm (blue). The concentration of Cy3 is 3.6 108 M and the concentration of AOT is 0.02 M.

of AOT increases. To investigate this hypothesis, we performed fluorescence emission experiments keeping the overall Cy3 concentration constant at 1.8 108 M and holding the reverse micelle size to w0 = 1, but varying the AOT concentration. Figure 5 shows the emission spectra of Cy3 as a function of AOT concentration and at two different excitation wavelengths, 532 and 514 nm. At higher AOT concentration the spectra exhibit less divergence from the Cy3 spectrum in bulk water. This result further supports our hypothesis that the extremely confined environment of the w0 = 1 reverse micelles drives the aggregation of the Cy3 molecules. Another experiment to confirm the presence of H-dimers measured the fluorescence lifetime of Cy3 in different environments. We expect longer lifetimes for the H-dimer compared to the Cy3 monomer because dimerization provides rigidification that suppresses nonradiative relaxation pathways.37 Table 1 shows the fluorescence lifetime data for Cy3 in water, and in w0 = 30 and w0 = 1 reverse micelle samples. The data for Cy3 in water agrees with literature reports.47 In the w0 = 30 reverse micelle environment, the fluorescence decay displays two distinct components, consistent with the dye experiencing different environments over the course of its lifetime. The shorter lifetime component is similar to that of bulk water, but the longer lifetime may be indicative of the different environments present in the reverse micelle that could influence the spectroscopic properties of the dye. The lifetime data is most interesting in the case of Cy3 in w0 = 1 reverse micelles, where not only is the decay biexponential but also the slower component is significantly longer than that of Cy3 in bulk water or larger reverse micelle sizes. To explore the nature of the longer component, we measured the fluorescence

wavelength (nm)



561

1.8 (84)

6.9 (14)

600

2.0 (62)

10.4 (38)

650

1.7 (60)

8.7 (40)

a

The samples were excited at 514 nm and the lifetimes measured at three different wavelengths, 561, 600, and 650 nm. The concentration of Cy3 is 3.6 108 M and the concentration of AOT is 0.02 M. The errors in the reported lifetimes are ∼0.1 ns.

lifetimes of Cy3 in the w0 = 1, exciting it at 514 nm and collecting the emission at 561, 600, and 650 nm, corresponding to the three peaks we observe in the emission spectrum. Figure 6 shows the decays measured at 561 and 650 nm and results from biexponential fits are given in Table 2. The lifetimes measured are longer when monitoring the longer wavelength emission; we attribute the longer fluorescence lifetime component to the presence of H-dimers, which is consistent with reports of H-dimers of other cyanine dyes.37 This data also suggests the emission peaks observed at 600 and 650 nm in Figure 3b for w0 = 1 and w0 = 2 reverse micelles are likely due to emission from Cy3 H-dimers, whereas the emission near 560 nm is due to Cy3 monomers. The suppression of the 560 nm peak relative to the 600 and 650 nm peaks in the w0 = 1 sample reveals the dyes exist predominantly as H-dimers in this sample. Finally, if the species excited at 514 nm is the H-dimer, then exciting the same samples at a longer wavelength should produce spectra that are consistent with the monomer Cy3 molecules that are also present in the system. This was confirmed by the fluorescence emission experiments shown in Figure 4, where different spectral features arise when exciting Cy3 in the w0 = 1 at 514 and 532 nm. To further investigate the excitation wavelength dependence of the Cy3 emission, we performed fluorescence excitation experiments at three different monitoring wavelengths, 561, 600, and 650 nm. Figure 6 shows that emission at 600 and 650 nm correlates with absorption at a shorter wavelength compared to emission at 560 nm. This data provides further evidence that two forms of the dye exist in the w0 = 1 reverse micelle samples and that each form can be accessed by exciting the samples at different wavelengths.

’ DISCUSSION The results presented above support our hypothesis that w0 = 1 reverse micelles provide a unique environment that drives Cy3 to undergo reactions that would not occur in bulk solutions of similar overall concentration. These micelles behave as nanoreactors that drive the formation of H-dimers, even at extremely low overall dye concentrations. In the presence of larger w0 = 930 reverse micelles, where the environment resembles that of bulk water more closely,14,63 this behavior is not seen. Our observations suggest the spectroscopic changes caused by aggregation may be present in w0 = 2 sized reverse micelles, but they are most obvious in w0 = 1 reverse micelles. We find this result highly unexpected, as the concentration of dye molecules is so low compared to the concentration of reverse micelles that the occupation number for our system ranges from one Cy3 molecule per 20 000 to 900 000 reverse micelles. To gain insight into this unexpected behavior, we consider some properties of a single reverse micelle containing one Cy3 molecule. A Cy3 molecule can be approximated as a prolate 9581



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Figure 8. Total fluorescence for varying concentrations of Cy3 in 0.02 M AOT w0 = 1 reverse micelle samples excited at 514 nm. The concentration of Cy3 ranges from 1.78 109 to 3.56 108 M. The data were analyzed according to eq 11 and the resulting fit line is shown. Figure 7. Normalized fluorescence excitation spectra for Cy3 in a 0.02 M AOT w0 = 1 reverse micelle sample, monitored at three different emission wavelengths: 561 nm (blue), 600 nm (pink), and 650 nm (red). The concentration of Cy3 is 3.6 108 M.

ellipsoid with short and long axes of ∼0.5 and ∼1.3 nm, respectively. The w0 = 1 reverse micelle can be approximated as a sphere with hydrodynamic radius ∼1.7 nm.16 If the reverse micelle water pool had the same density as bulk water, a 1.7 nm hydrodynamic radius implies that the water pool of the w0 = 1 reverse micelle contains ∼600700 water molecules. Thus, it is reasonable to assume that a single reverse micelle is large enough to enclose a Cy3 molecule and contains enough water to solvate the dye. However, we cannot rule out the possibility that the Cy3 may be associated with the charged head groups or the aliphatic tail groups of the reverse micelle, and thus may not be fully solvated by water. Formation of Cy3 H-dimers may occur when two reverse micelles, each containing a single Cy3 molecule, interact so that they allow the Cy3 molecules to interact. This would result in a local concentration for the two interacting Cy3 molecules within two reverse micelles of ∼0.1 M, which is sufficient to stabilize the Cy3 molecules in an H-dimer. In contrast, the local concentration of two Cy3 molecules in w0 = 9 reverse micelles is only ∼0.01 M. Using the assumption that two w0 = 1 reverse micelles containing one Cy3 monomer each must collide to form an H-dimer, we estimated a range of initial rates for such a collision. Assuming the reaction is diffusion controlled, these rates are given by the equation: rate0 ¼ kD ½A0 2

8kB T 3η

τ1=2 ¼

ð5Þ

½A0 ¼ ½A þ 2½A 2

ð6Þ

where [A]0 is the overall Cy3 concentration, [A] is the concentration of Cy3 monomer, and [A2] is the concentration of Cy3 H-dimer. The equilibrium constant governing the association/ dissociation reaction is then given by

ð4Þ

where kB is the Boltzmann constant, T is the laboratory temperature (∼294 K), and η is the viscosity of the isooctane solvent (0.473 mPa s).64 The set of conditions used in our experiments yields kD = 1.4 1010 L mol1 s1. Using this value in eq 2, we obtain initial rates ranging from 1.8 105 to

1 kD ½A0

which ranges from ∼2 to 40 ms. Thus, although the occupation number for our system is extremely low, the rate of collisions between reverse micelles containing Cy3 molecules is high enough to allow for this aggregation to occur. We hypothesize that, as two individual reverse micelles come in contact, each containing a Cy3 molecule, the local concentration is so high that the reaction is favored toward the formation of the Cy3 dimer. The fact that changes in the spectroscopy of Cy3 occur in the w0 = 2 reverse micelle (see Figures 2 and 3) indicates this could be a point where the formation of H-dimers starts to become significant. To further characterize the thermodynamic properties of Cy3 H-dimer formation in w0 = 1 reverse micelles, we also carried out a set of experiments to determine the overall equilibrium constant for the association/dissociation of the dimer. To do so, we measured the total fluorescence emission intensities above 530 nm for varying concentrations of Cy3 in w0 = 1 reverse micelles excited at 514 nm, shown in Figure 8. Assuming Cy3 dimer formation, we derived a series of equations to analyze the data shown in Figure 8. First, the mass balance equation for our system is

ð3Þ

where kD is the diffusion limited rate constant and [A]0 is the initial Cy3 monomer concentration in the bulk solution. The parameter kD is given by kD ¼

4.5 108 M s1. Furthermore, the half-life, τ1/2, is given by the equation

K ¼

½A 2 ½A 2 2 ¼ 2 ½A ½A0 4½A 2 ½A0 þ 4½A 2 2

ð7Þ

Solving eq 7 for [A2] yields pffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi ð4K½A0 þ 1Þ 8K½A0 þ 1 ½A 2 ¼ 8K 9582

ð8Þ



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Second, we assumed the fluorescence emission rate shown in Figure 7 is determined by the equation F ¼ εm ½A þ εd ½A 2

ð9Þ

where F is the total fluorescence emission rate, εm is the molar brightness of the monomer, and εd is the molar brightness of the H-dimer. The molar brightness depends on the absorption cross section and quantum yield of the species being probed as well as the collection efficiency of the optical setup. Given the mass balance in eq 6, we obtain the following expression F ¼ εm ½A0 þ ðεd 2εm Þ½A 2

ð10Þ

and substituting eq 8 into eq 10 gives F ¼ εm ½A0

pffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi! ð4K½A0 þ 1Þ 8K½A0 þ 1 þ ðεd 2εm Þ 8K ð11Þ

The fluorescence intensity for varying Cy3 concentrations is shown in Figure 7. This data was fit to eq 11 using K and εd as adjustable parameters. To calculate εm, we assumed the fluorescence emission at the lowest Cy3 concentration, 8.93 1010 M, arose predominantly from the Cy3 monomer. From the measured fluorescence intensity of this solution, we obtained a value for εm ≈ 6 109 kHz M1. To provide an initial estimate for εd, we assumed the fluorescence emission at the highest Cy3 concentration was predominantly from the Cy3 dimer, which gives an initial estimate of εd ≈ 2.4 1010 kHz M1. On the basis of this fitting procedure, we determined K ≈ (1.4 ( 0.7) 108 M1 and εd ≈ (2.9 ( 0.16) 1010 kHz M1. In the accompanying paper,59 we use fluorescence correlation spectroscopy to investigate the kinetics of the Cy3 H-aggregation in reverse micelles. We find that the formation of Cy3 H-dimers is accompanied by the formation of a transient dimer complex between two reverse micelles.

’ CONCLUSION We have demonstrated the effect of reverse micelle environment on Cy3 dye molecules. We observed that the dye forms dimers in the extremely confined environment provided by w0 = 1 reverse micelles. This aggregation alters the spectroscopic properties and fluorescence lifetime of the dye compared to that of isolated Cy3 monomers. Furthermore, it is possible to access each form of the dye, depending on the excitation wavelength. What is most interesting about these results is that this behavior is not seen at the same overall concentrations in bulk water or in the larger, w0 = 930, reverse micelles. This suggests that the w0 = 1 reverse micelles provide a unique environment for the dye to undergo reactions that would not occur in bulk solution. Moreover, this unexpected aggregation is very interesting considering that the occupation number of the reverse micelles ranges from one Cy3 molecule per 20 000 to 900 000 reverse micelles. The results of this study indicate that the reverse micelle environment can provide a catalyst to cause reactions to occur that would not occur under bulk solution conditions. These results are relevant for studying molecular behavior in extremely

crowded and confined biological environments such as cells, organelles, and cellular nuclei.

’ AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected] (A.V.O.); levinger@ lamar.colostate.edu (N.E.L.).

’ ACKNOWLEDGMENT Funding for this work was provided by National Science Foundation CRC grant 0628260. Funds to purchase the Fluorolog 3 instrument used here were made possible through National Science Foundation grant DBI-0302571. We thank Prof. B. George Barisas and Peter Winter for helpful discussions and technical assistance with fluorescence lifetime measurements, and members of the Crans group for assistance with the purification of the AOT surfactant. ’ REFERENCES (1) Terai, T.; Nagano, T. Curr. Opin. Chem. Biol. 2008, 12, 515–521. (2) Prinz, A.; Reither, G.; Diskar, M.; Schultz, C. Proteomics 2008, 8, 1179–1196. (3) Lavis, L. D.; Raines, R. T. ACS Chem. Biol. 2008, 3, 142–155. (4) Kim, H. M.; Cho, B. R. Acc. Chem. Res. 2009, 42, 863–872. (5) Zhao, R.; Rueda, D. Methods 2009, 49, 112–117. (6) Lilley, D. M. J. The Structure and Folding of Branched RNA Analyzed by Fluorescence Resonance Energy Transfer. In Methods In Enzymology, Vol. 469: Biophysical, Chemical, and Functional Probes of RNA Structure, Interactions and Folding, Part B; Academic Press: New York, 2009; Vol. 469, pp 159187. (7) Okamoto, K.; Sannohe, Y.; Mashimo, T.; Sugiyama, H.; Terazima, M. Bioorg. Med. Chem. 2008, 16, 6873–6879. (8) Schuler, B. Chemphyschem 2005, 6, 1206–1220. (9) Sun, Y. S.; Landry, J. P.; Fei, Y. Y.; Zhu, X. D. Langmuir 2008, 24, 13399–13405. (10) De, T. K.; Maitra, A. Adv. Colloid Interface Sci. 1995, 59, 95–193. (11) Luisi, P. L.; Giomini, M.; Pileni, M. P.; Robinson, B. H. Biochim. Biophys. Acta 1988, 947, 209–246. (12) Pileni, M. P. J. Phys. Chem. 1993, 97, 6961–6973. (13) Baruah, B.; Swafford, L. A.; Crans, D. C.; Levinger, N. E. J. Phys. Chem. B 2008, 112, 10158–10164. (14) Fayer, M. D.; Levinger, N. E. Analysis of Water in Confined Geometries and at Interfaces. In Annual Review of Analytical Chemistry; Annual Reviews: Palo Alto, CA, 2010; Vol. 3; pp 89107. (15) Martinez, A. V.; DeSensi, S. C.; Dominguez, L.; Rivera, E.; Straub, J. E. J. Chem. Phys. 2011, 134. (16) Zulauf, M.; Eicke, H. F. J. Phys. Chem. 1979, 83, 480–486. (17) Abuin, E.; Lissi, E.; Solar, C. J. Colloid Interface Sci. 2005, 283, 87–93. (18) Correa, N. M.; Durantini, E. N.; Silber, J. J. J. Org. Chem. 1999, 64, 5757–5763. (19) Martinek, K.; Levashov, A. V.; Klyachko, N.; Khmelnitski, Y. L.; Berezin, I. V. Eur. J. Biochem. 1986, 155, 453–468. (20) Menger, F. M.; Yamada, K. J. Am. Chem. Soc. 1979, 101, 6731–6734. (21) Novaira, M.; Moyano, F.; Biasutti, M. A.; Silber, J. J.; Correa, N. M. Langmuir 2008, 24, 4637–4646. (22) Correa, N. M.; Zorzan, D. H.; D’Anteo, L.; Lasta, E.; Chiarini, M.; Cerichelli, G. J. Org. Chem. 2004, 69, 8231–8238. (23) Deligeorgiev, T.; Vasilev, A.; Kaloyanova, S.; Vaquero, J. J. Color. Technol. 2010, 126, 55–80. (24) Mishra, A.; Behera, R. K.; Behera, P. K.; Mishra, B. K.; Behera, G. B. Chem. Rev. 2000, 100, 1973–2011. 9583


The Journal of Physical Chemistry B (25) Dempster, D. N.; Thompson, G. F.; Morrow, T.; Rankin, R. J. Chem. Soc., Faraday Trans. 2 1972, 68, 1479–&. (26) Dipaolo, R. E.; Scaffardi, L. B.; Duchowicz, R.; Bilmes, G. M. J. Phys. Chem. 1995, 99, 13796–13799. (27) Bergmann, K.; Okonski, C. T. J. Phys. Chem. 1963, 67, 2169. (28) West, W.; Pearce, S. J. Phys. Chem. 1965, 69, 1894. (29) Cooper, W.; Liebert, N. B. Photog. Sci. Eng. 1972, 16, 25. (30) Chambers, R. W.; Kajiwara, T.; Kearns, D. R. J. Phys. Chem. 1974, 78, 380–387. (31) Rentsch, S. K.; Danielius, R. V.; Gadonas, R. A.; Piskarskas, A. Chem. Phys. Lett. 1981, 84, 446–449. (32) Sundstrom, V.; Gillbro, T. J. Chem. Phys. 1985, 83, 2733–2743. (33) Vanderauweraer, M.; Biesmans, G.; Deschryver, F. C. Chem. Phys. 1988, 119, 355–375. (34) Chibisov, A. K.; Zakharova, G. V.; Gorner, H. Phys. Chem. Chem. Phys. 1999, 1, 1455–1460. (35) Mandal, A. K.; Pal, M. K. J. Colloid Interface Sci. 1997, 192, 83–93. (36) Mandal, A. K.; Pal, M. K. Chem. Phys. 2000, 253, 115–124. (37) Rosch, U.; Yao, S.; Wortmann, R.; Wurthner, F. Angew. Chem., Int. Ed. 2006, 45, 7026–7030. (38) Zhang, Y. Z.; Xiang, J. F.; Tang, Y. L.; Xu, G. Z.; Yan, W. P. Dyes Pigm. 2008, 76, 88–93. (39) Gadde, S.; Batchelor, E. K.; Kaifer, A. E. Chem.—Eur. J. 2009, 15, 6025–6031. (40) Nikolenko, L. M.; Ivanchihina, A. V.; Brichkin, S. B.; Razumov, V. F. J. Colloid Interface Sci. 2009, 332, 366–372. (41) Chen, Y. L.; Lv, Y. X.; Han, Y.; Zhu, B.; Zhang, F.; Bo, Z. S.; Liu, C. Y. Langmuir 2009, 25, 8548–8555. (42) Chen, Z. J.; Lohr, A.; Saha-Moller, C. R.; Wurthner, F. Chem. Soc. Rev. 2009, 38, 564–584. (43) Davis, R.; Kumar, N. S. S.; Abraham, S.; Suresh, C. H.; Rath, N. P.; Tamaoki, N.; Das, S. J. Phys. Chem. C 2008, 112, 2137–2146. (44) Lau, V.; Heyne, B. Chem. Commun 2010, 46, 3595–3597. (45) Seibt, J.; Lohr, A.; Wurthner, F.; Engel, V. Phys. Chem. Chem. Phys. 2007, 9, 6214–6218. (46) Jia, K.; Wan, Y.; Xia, A. D.; Li, S. Y.; Gong, F. B.; Yang, G. Q. J. Phys. Chem. A 2007, 111, 1593–1597. (47) Sanborn, M. E.; Connolly, B. K.; Gurunathan, K.; Levitus, M. J. Phys. Chem. B 2007, 111, 11064–11074. (48) Widengren, J.; Schwille, P. J. Phys. Chem. A 2000, 104, 6416–6428. (49) Conley, N. R.; Biteen, J. S.; Moerner, W. E. J. Phys. Chem. B 2008, 112, 11878–11880. (50) Hoebeke, M.; Piette, J.; Vandevorst, A. J. Photochem. Photobiol., B 1990, 4, 273–282. (51) Kuzmin, V. A.; Darmanyan, A. P. Chem. Phys. Lett. 1978, 54, 159–163. (52) Momicchioli, F.; Baraldi, I.; Berthier, G. Chem. Phys. 1988, 123, 103–112. (53) Seret, A.; Hoebeke, M.; Vandevorst, A. Photochem. Photobiol. 1990, 52, 601–604. (54) Sundstrom, V.; Gillbro, T. J. Phys. Chem. 1982, 86, 1788–1794. (55) Sundstrom, V.; Gillbro, T.; Bergstrom, H. Chem. Phys. 1982, 73, 439–458. (56) Widengren, J.; Seidel, C. A. M. Phys. Chem. Chem. Phys. 2000, 2, 3435–3441. (57) Stahla, M. L.; Baruah, B.; James, D. M.; Johnson, M. D.; Levinger, N. E.; Crans, D. C. Langmuir 2008, 24, 6027–6035. (58) Chowdhury, P. K.; Ashby, K. D.; Datta, A.; Petrich, J. W. Photochem. Photobiol. 2000, 72, 612–618. (59) McPhee, J. T.; Scott, E.; Levinger, N. E.; Van Orden, A. J. Phys. Chem. B 2011, in press. DOI: http://dx.doi.org/10.1021/jp2001282. (60) Mujumdar, R. B.; Ernst, L. A.; Mujumdar, S. R.; Lewis, C. J.; Waggoner, A. S. Bioconjugate Chem. 1993, 4, 105–111. (61) Levitus, M.; Ranjit, S. Q. Rev. Biophys. 2011, 44, 123–151. (62) Kubin, R. F.; Fletcher, A. N. J. Lumin. 1982, 27, 455–462. (63) Baruah, B.; Crans, D. C.; Levinger, N. E. Langmuir 2007, 23, 6510–6518.

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(64) Padua, A. A. H.; Fareleira, J. M. N. A.; Calado, J. C. G. J. Chem. Eng. Data 1996, 41, 1488–1494.

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Cy3 in AOT Reverse Micelles I. Dimer Formation Revealed through

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