Influence of Morphology on Polar Solvation Dynamics in Lecithin

J. Phys. Chem. B 2000, 104, 11075-11080

11075

Influence of Morphology on Polar Solvation Dynamics in Lecithin Reverse Micelles Dale M. Willard and Nancy E. Levinger* Department of Chemistry, Colorado State UniVersity, Fort Collins, Colorado 80523-1872 ReceiVed: June 8, 2000; In Final Form: August 31, 2000

The continuous nonpolar phase can change the morphology of lecithin reverse micelles dramatically. In alkane solvents, increasing the hydration leads to gelation as isolated water droplets transform into an entangled tube network. In contrast, increasing the hydration for lecithin in benzene maintains isolated, largely spherical traditional reverse micelles. Solvation dynamics experiments have been carried out in the two different micellar environments. These measurements show that solvation dynamics are considerably more restricted in the tubular micelles than in the spherical droplets. This is interpreted as evidence for water pool formation and disorder at the interface in the spherical micelles and supports the hypothesis of substantial water incorporation into the tubular micelles. To perform solvation dynamics experiments in the benzene/lecithin/water reverse micelles, we have synthesized a new headgroup-labeled probe molecule. This synthesis, coupling the standard solvation dynamics probe, coumarin 343, to phosphatidylethanolamine through an amide bond, is presented.

I. Introduction Reverse micelles can be formed from a number of different surfactants, among them are biologically relevant lipids such as phosphatidylcholine or lecithin. The forms of these micelles range from isolated spherical droplets to entangled tubular networks and depend on a number of different properties, including the surfactant, the nonpolar continuous phase, the water concentration, and the presence of other solutes such as enzymes, among others.14,34 Of particular interest for studies of reverse micelles is the variation in properties of water that is solubilized within them. Various methods have been used to investigate the structural and dynamical properties of intramicellar water.10 Vibrational and NMR spectroscopy have shown that interactions with the surfactant headgroups perturb the water structure. Researchers have proposed distinct water environments, or shells, based on their proximity to the surfactant interface.15 As the water content in the reverse micelles increases, the characteristics of the water approach those of bulk water. Although results vary, several studies have proposed three water types to exist in various reverse micellar systemssthat is, water strongly bound to the surfactant, water bound to the surfactant, and bulklike, free water.6,16 In addition to perturbing the structure of intramicellar water, the reverse micellar environment can significantly alter its dynamical properties. While a range of measurements have shown that the mobility of water at the lecithin interface is significantly lower than that in bulk water,12,19,21,36,39,43 the techniques used could not follow the ultrafast motion of water. Hence, these studies do not distinguish water interacting with the interface unless its dynamical response is slowed by orders of magnitude. One way of measuring the ultrafast dynamics of water motion is with ultrafast polar solvation dynamics.23 In this technique, an ultrashort laser pulse promotes a dye molecule with a small ground-state dipole to its excited state, where the dipole is large. Following the time-resolved fluorescence Stokes shift in time * Corresponding author. Email: [email protected].

allows measuring the water response to this instantaneous perturbation. While the dynamics of polar solvation have been measured in a wide range of bulk solvents,4,23 there are only a few references to the solvation dynamics in restricted environments such as reverse micelles. The overall result from all these experiments is that the solvation dynamics inside the reverse micelles are significantly slower than those in bulk aqueous media.22,24-26,28,30-33,44 However, in our previous work, we have observed that some of the micellar systems appear to immobilize water more effectively than others.26,28,30-32,44 For example, in aerosol OT (AOT) reverse micelles, bulklike water motion was observed for reverse micelles at a water-to-surfactant ratio where lecithin reverse micelles displayed no motion.32,44 In this paper, we report on the solvation dynamics of water at lecithin reverse micellar interfaces. We have studied two related reverse micellar systems, namely, lecithin reverse micelles in cyclohexane and benzene. Except for the nonpolar continuous phase, the microemulsions are identical. In cyclohexane, the microemulsion forms an entangled tube network; in benzene, for the same hydration level, the microemulsion displays isolated reverse micelles. We have probed polar solvation dynamics in each case to compare the effect that the mesoscopic morphology has on the dynamics. We find that the form of the microemulsion and the effect of the nonpolar solvent on the interfacial order strongly influence the dynamics. To facilitate our study of the solvation dynamics in benzene/lecithin/ water reverse micelles, we have synthesized a new amphiphilic probe molecule. Details of this synthesis and characterization are also presented. II. Experimental Methods II.A. Synthesis of Coumarin-Labeled Phosphatidylethanolamine (CLPE). Because the probe molecule of coumarin 343 (C343) that we have used in other reverse micelles experiments is highly soluble in benzene, we synthesized a headgroup-labeled surfactant molecule to serve as the probe in these experiments. Details of the synthesis and characterization follow.

10.1021/jp002076n CCC: $19.00 © 2000 American Chemical Society Published on Web 11/05/2000

11076 J. Phys. Chem. B, Vol. 104, No. 47, 2000

Figure 1. Synthetic route for coupling the C343 probe molecule to PE. The presumed mechanism has been taken from ref 45.

Materials. 1,2-Dioleoyl-syn-glycero-3-phosphoethanolamine (PE, powder form, Avanti Polar Lipids, Inc), C343 (Exciton, Inc.), triphenylphosphine (PPh3, Aldrich), aldrithiol-2 (Aldrich), 4-(dimethylamino)pyridine (DMAP, Aldrich), methanol (Fisher, ACS grade), acetic acid (Aldrich), and chloroform (Fisher, HPLC grade) were used as received. Methylene chloride (Fisher, HPLC grade) used in the reaction was distilled over calcium hydride. Deuterated solvents for NMR analysis were obtained from Cambridge Isotopes Laboratories, Inc. High-purity water (MilliQ filtered, 18.2 MΩ-cm) was used in the synthesis and subsequent reverse micelle preparation. Chromatography columns were packed with silica gel (Silicycle, 230-400 mesh). Plates for thin-layer chromatography were coated with silica gel 60 F254 (EM Separations Technology, 250 µm layer thickness.) Synthesis. The reaction scheme is shown in Figure 1. C343 (0.27 mmol, 76 mg) was activated using PPh3 (0.27 mmol, 71 mg) and aldrithiol (0.27 mmol, 59 mg) in dry methylene chloride (5 mL). The mixture was stirred for 2 h under N2 at room temperature. Once activated, PE (0.134 mmol, 100 mg) and DMAP (0.13 mmol, 16 mg) were added to form the desired amide bond. This mixture was stirred under N2 at room temperature for 2 h. The reaction was quenched with 8 mL of saturated aqueous sodium bicarbonate solution, 11 mL of methanol, and 20 mL of chloroform. The organic phase was subsequently washed twice with 10 mL of the saturated sodium bicarbonate solution and separated, and the solvent was removed in vacuo. The sample was left under vacuum overnight to remove any residual water. Purification was accomplished via silica gel chromatography with a 90:8:2 methylene chloride: methanol:acetic acid mobile phase. On a thin-layer chromatographic plate, the product has a Rf ) 0.3 with this mobile phase.1 The purified product was dried in vacuo, followed by evaporation under vacuum overnight. The final product appeared as flaky yellow-brown crystals (136 mg, 98% yield). Product Characterization. 1H NMR (9:1 CD2Cl2:CD3OD at 25 °C): δ 9.19 (s, 1 H), 8.49 (s, 1 H), 7.02 (s, 1 H), 5.32 (m, 4 H), 5.17 (m, 1 H), 4.31 (broad m, 1 H), 4.00 (broad m, 5 H), 3.66 (s, 2 H), 3.32 (s, 4 H), δ 2.74 (broad m, 4 H), 2.23 (m, 4 H), 1.97 (m, 12 H), 1.52 (m, 4 H), 1.25 (m, 40 H), 0.86 (m, 6 H). All NMR spectra were recorded with a Varian Inova‚300 MHz spectrometer. Chemical shifts were referenced to tetramethylsilane (TMS). Mass spectra were measured using fast atom bombardment (FAB) with a Fisons VG AutoSpec instrument calibrated with PPG. Positive ion FAB m/z: (CLPE-‚2H+, 100%) calcd 1011.6439, found 1011.6437; (CLPE-‚H+‚Na+, 6.8%) calcd 1033.626, found 1033.629. II.B. Sample Preparation and Characterization. Reverse micellar samples were prepared with the CLPE probe in benzene (Fisher, ACS grade). The samples were also prepared in cyclohexane (Acros, spectrophotometric grade, 99+%) for

Willard and Levinger comparison with previous results.44 The solvents were used as received. Lyophilized egg phosphatidylcholine (lecithin) was purchased from Avanti Polar Lipids, Inc. Lecithin was stored at -10 °C and used without further purification. The samples were always prepared from newly opened bottles to minimize water absorption. Materials, concentrations, and methods for preparing the lecithin reverse micelles were similar to those described earlier.44 The final concentration of the probe in the solutions was ∼200 µM, yielding an absorbance of ∼1 OD in the 1 mm path length optical cuvette. The major difference was the incorporation of CLPE into the lecithin. To ensure that CLPE was homogeneously distributed within the reverse micelles, we dispersed and dried lecithin and CLPE before introducing them into the supporting benzene or cyclohexane nonpolar phase. Specifically, 500 mg of lecithin and 2 mg of CLPE were codispersed in a 9:1 methylene chloride:methanol solvent, which was subsequently removed by vacuum evaporation. Unevaporated methylene chloride/methanol was removed by three cycles in which lipids were dissolved in ∼10 mL of benzene or cyclohexane followed by evaporation for one or more hours. 1H NMR analysis (Bruker AM‚500 MHz Spectrometer) revealed 0.4 ( 0.1 mol H2O/mol lecithin after 3 cycles of cyclohexane or benzene addition and vacuum evaporation. The samples showed the expected viscosity as a function of hydration, and the reverse micelles in benzene displayed hydrodynamic radii consistent with those in published reports.40 The hydration level of the reverse micelles was characterized according to

w0 )

[H2O] [lecithin]

(1)

Micelle sizes were determined using dynamic light scattering (DynaPro, MSTC). Absorption spectra were recorded with a Cary 2400 UVvis-NIR spectrophotometer. Steady-state emission spectra were measured with a home-built fluorometer.7 The concentration of the fluorescent probe in the reverse micelles was similar to that in our previous work on C343 in water/lecithin/cyclohexane reverse micelles.44 II.C Time-Resolved Measurements. Both time-resolved fluorescence anisotropy and Stokes shift measurements were performed with a fluorescence upconversion spectrometer that has been described in detail.32 The doubled output (410 nm) from an ultrafast mode-locked Ti:sapphire laser excites the dye to its first excited state; the resulting fluorescence is mixed with the residual laser fundamental gate pulse in a nonlinear crystal with variable time delay, thereby time-resolving the fluorescence. The time resolution of the spectrometer was measured via cross correlation of the pump and gate pulses to be 200 ( 30 fs. For time-resolved fluorescence Stokes shift measurements, the angle between the polarization of the pump and gate pulses is kept at the magic angle to eliminate effects from rotational diffusion. For time-resolved fluorescence anisotropy measurements, fluorescence decays were collected with the polarization of the pump and probe beams both parallel and perpendicular to each other. Time-resolved fluorescence Stokes shift data were collected at 10 different wavelengths from 440 to 530 nm at 10 nm intervals. Individual fluorescence decays were measured on three different time scales, 2, 33, and 500 ps, with 16.8 fs, 233 fs, and 3 ps time steps, respectively, to measure processes occurring in different timescales. Individual fluorescence scans were fit to a multiexponential function using an iterative-reconvolution

Lecithin Reverse Micelles

J. Phys. Chem. B, Vol. 104, No. 47, 2000 11077

fit program with the cross correlation of the pump and gate pulses as the instrument response function. From these fits, the time-resolved fluorescence spectra were reconstructed. The time correlation function, C(t),

C(t) )

ν(t) - ν(∞) ν(0) - ν(∞)

(2)

where ν(t), ν(0), and ν(∞) are the peaks of the fluorescence spectra at time t, instantaneously, and at equilibrium, respectively, was calculated using the peak positions of the reconstructed fluorescence spectra fitted to log normal line shapes. Time-resolved fluorescence anisotropy measurements were analyzed via

r(t) )

I||(t) - I⊥(t) I||(t) + 2I⊥(t)

(3)

where I||(t) and I⊥(t) are the time-dependent intensities of the upconverted fluorescence with the polarization of the pump and probe pulses parallel and perpendicular to each other, respectively. The initial anisotropy was close to 0.4 for the CLPE probe, showing that it is an ideal anisotropy probe and that samples are isotropic. III. Results and Discussion III.A. Assessment of the CLPE Probe. Before the solvation dynamics measurements were made, CLPE was tested for viability as a solvation dynamics probe. It was particularly important to ensure that the CLPE probe effectively reports information about the dynamics in the micellar interior. We assessed the utility of CLPE in these experiments by comparing the steady-state absorption and fluorescence spectra and the time-resolved fluorescence measurements using CLPE in lecithin/ cyclohexane reverse micelles to our previous results using the free C343 dye in the same reverse micelles. Figure 2 displays the steady-state absorption and emission spectra of C343 and CLPE in cyclohexane/lecithin/water reverse micelles. The C343 parent fluorophore shows strong spectral dependence on the solvent,11,29,37; thus, we use the spectrum to help identify the location of the probe. We have previously determined that C343 resides inside the reverse micelles.44 The CLPE absorption spectrum displays a single peak at 424.5 nm, the same position but slightly narrower than that for C343 in the same environment. The CLPE emission spectrum, peaking at 479 nm, resembles the emission of C343 in the same environment but shifted slightly (∼2 nm) to the blue. We would expect the larger shifts, broadening, or appearance of additional peaks in both spectra if a percentage of CLPE molecules were aligned counter to the lecithin orientation or if the CLPE molecules resided in a range of differing locations in the microemulsion. On the contrary, the slight narrowing in the CLPE absorption spectrum may indicate a more homogeneous environment for CLPE compared to that for C343. Moreover, results from our earlier work on C343 in cyclohexane/lecithin/ water reverse micelles, in which the C343 spectra show similar widths for aqueous solution and reverse micelles,44 and the lack of evidence of coumarin aggregation both in our own experiments and in the literature indicate that the differences in spectral width for CLPE and C343 do not arise from the aggregation of C343. The similarities in both the emission and absorption spectra indicate that the CLPE fluorophore resides inside the micelles.

Figure 2. Steady-state (a) absorption and (b) emission spectra of the free dye, C343, and the headgroup-labeled dye, CLPE, in w0 ) 6.4 cyclohexane/lecithin/water reverse micelles.

Figure 3. (a) Absorption and (b) emission spectra of CLPE in cyclohexane/lecithin/water reverse micelles as functions of hydration (w0).

Figure 3 shows the absorption and emission of CLPE in cyclohexane/lecithin/water reverse micelles as a function of hydration. Previously, we observed red-shifting with increasing hydration for the C343 absorption and emission spectra in the same reverse micelles. The CLPE emission spectra show similar

11078 J. Phys. Chem. B, Vol. 104, No. 47, 2000

Willard and Levinger

TABLE 1: Measured Solvation Dynamics Components in Lecithin Reverse Micelles as Functions of Probe and Hydrationa probe

nonpolar solvent

w0

a1

τ1 (fs)

a2

τ2 (ps)

a3

τ3 (ps)

C343 CLPE C343 CLPE

cyclohexane cyclohexane cyclohexane benzene

5.8 6.4 6.8 6.4

0.100 ( 0.003 0.11 ( 0.01 0.13 ( 0.01 0.60 ( 0.01

420 ( 30 620 ( 10 570 ( 70 85 ( 3

0.17 ( 0.03 0.22 ( 0.01 0.25 ( 0.01 0.16 ( 0.01

17 ( 7 11 ( 1 14 ( 1 5.4 ( 0.6

0.74 ( 0.07 0.68 ( 0.01 0.62 ( 0.02 0.24 ( 0.01

340 ( 100 210 ( 10 320 ( 40 160 ( 20

a

Fits to a multiexponential function, C(t) ) ∑iaie-t/τi.

TABLE 2: Time-resolved Fluorescence Anisotropy for the Rotational Relaxation Parameters of C343 and CLPE in Pure Solvents and Various Reverse Micellar Environmentsa probe

solvent

wb

C343 C343 C343 CLPE C343 CLPE

benzene waterc cyclohexane cyclohexane benzene benzene

5e 5e 5 6.4

contribution (%) 100 100 72.0 ( 0.5 18 ( 2

τ1 (ps) 38.5 ( 6 83 35.3 ( 0.5 90 ( 20

contribution (%)

100 100 28.0 ( 0.3 82 ( 2

τ2 (ns)

2.4 ( 0.2 2.9 ( 0.3 2.5d 2.5d

a Anisotropy decays are fitted to a sum of exponentials function,r(t) ) ∑ a e-t/τi. b This column refers to reverse micellar samples only. c From i i ref 20. d Variable held fixed during fitting. e Gel phase.

hydration-dependent spectral shifting peaks at 473 nm for w0 ) 2.4 and 479 nm for w0 ) 6.4. The sensitivity of the CLPE fluorphore provides strong evidence that CLPE aligns with the coumarin probe pointing into the intramicellar pool. If a percentage of CLPE molecules were incorrectly aligned, we would expect a corresponding proportion of the emission spectra to be insensitive to the hydration level. The spectra also indicate that CLPE resides in an environment similar to that of C343. In addition to steady-state spectroscopy, fluorescence quenching experiments also indicate contact of the CLPE chromophore with the micellar interior. The most crucial test for CLPE’s viability as a solvation dynamics probe was whether its spectral response accurately reflects solvent motion. In standard solvation dynamics probes, the intramolecular reorganization is small relative to the solvent reorganization and occurs on a time scale faster than we can detect. However, it is possible that covalent attachment of the nonrigid PE to the fluorophore could introduce more accessible intramolecular vibrational relaxation pathways for probe relaxation. To assess this issue and its effectiveness as a probe of solvation dynamics, we reproduced the solvation dynamics measurements of cyclohexane/lecithin/water reverse micelles obtained from C343 with CLPE as the probe. Table 1 lists the results from these experiments. If CLPE resided in a slightly different position or location within the micelles, we would expect some discrepancy between the dynamics observed with CLPE versus those observed with C343. However, as shown in Table 1, the results are remarkably similar. These results prove that CLPE aligns correctly within the reverse micelle, and the results detect the same dynamics as those for C343. In addition to using time-resolved fluorescence Stokes shift measurements to assess the solvation dynamics, we also compared results from the time-resolved fluorescence anisotropy of CLPE within cyclohexane/lecithin/water w0 ) 4.4 reverse micelles with related measurements of C343. Like C343, CLPE has only one ∼2 ns rotational relaxation component (see Table 2). From these results, we conclude that C343 and CLPE reside in very similar rotationally restricted environments. Taken together, the results strongly suggest that (1) the CLPE fluorophore resides in the reverse micellar interior, (2) CLPE is sensitive to the water motion inside reverse micelles, and (3) CLPE reports essentially the same information as does C343. Therefore, we feel confident that results generated with this molecule are a good measure of the dynamics inside the micelles.

III.B. Lecithin Reverse Micelles in Benzene. Our original reason for synthesizing CLPE was to make it possible to measure solvation dynamics in lecithin/benzene reverse micelles. In contrast to the lecithin/cyclohexane reverse micelles, which form a gel phase consisting of tubular nanostructures,34 the isolated drop structure persists with increasing hydration when microemulsions are formed with lecithin and water in benzene.41 We characterized the lecithin/benzene reverse micellar systems in a manner similar to that for the characterization of lecithin/ cyclohexane reverse micelles. First, the isolated reverse micellar structure was verified by dynamic light scattering (DLS). At w0 ) 6.4, we find that our reverse micelles in benzene have a radius of ∼2.8 nm. Assuming that the water molecular volume is ∼30 Å3 and that the reverse micelle aggregation number is 80, we find that this result agrees well with the results of Wachtel et al., and it predicts w0 ) 6 reverse micelles to have a radius of 2.9 nm.40 Similar to the spectra for lecithin reverse micelles in cyclohexane, the steady-state absorption and emission spectra of CLPE in the benzene micelles were also measured and are shown in Figure 4. These spectra showed similar trends to CLPE and C343 in lecithin/cyclohexane reverse micellessthat is, with increasing hydration the emission spectrum peak shifts from 469 nm at w0 ) 2.4 to 476 nm at w0 ) 6.4. The absorption spectrum shifts somewhat less than the spectra for cyclohexane systems, suggesting differences in the two micellar systems. The sensitivity of the dye to the hydration level indicates that the dye resides on the aqueous side of the interface rather than being self-assembled inside-out. Moreover, experiments introducing KI as a quencher in the intramicellar water pool indicate contact of the CLPE fluorophore with the micellar interior. In contrast, the results of time-resolved fluorescence anisotropy measurements for the lecithin/benzene system differed distinctly from those for lecithin/cyclohexane systems (see Table 2). Specifically, the anisotropy in the lecithin/cyclohexane reverse micelles displays only a single, long-time decay component; in benzene, an additional fast relaxation time appears. In related studies, we interpreted anisotropy decays with two time components (as evidence for the two locations for dye in the micelles, bound to the interface and free within the water pool32) because the time scales matched those for the dye in water32 and for the micelle rotational correlation.20 In the benzene/lecithin/water reverse micelles, the fast component of the anisotropy decay changes with hydration level and does not match the rotation time for the C343 either in benzene or in

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Figure 5. Time correlation function, C(t), indicating solvent relaxation in benzene/lecithin/water and cyclohexane/lecithin/water reverse micelles, (w ) 6.4).

Figure 4. (a) Absorption and (b) emission spectra of CLPE in benzene/ lecithin/water reverse micelles as functions of hydration (w0).

water26 (see Table 232). Furthermore, because the probe molecule is covalently attached to the phosphatidylethanolamine, the additional fast component cannot indicate the presence of free dye either in the benzene or inside the reverse micelles. Rather, we explain the fast component as a “wobbling-in-a-cone” motion of the fluorophore on the CLPE molecule. The amplitude of the wobble component indicates the degree to which the fluorophore can rotate in a conical volume, and the associated time component is the rate at which the fluorophore samples the possible configurations within the cone. Several models exist that determine the wobble cone angle on the basis of measured amplitudes. The model of Kinosita et al.17

r∞ ) [1/2cos θc(1 + cos θc)]2 r0

(4)

can be used to predict θc, the wobble cone angle, where r∞ represents the long time amplitude estimated by the amplitude of the longest decay component and r0 represents the initial amplitude at time zero. This model predicts that the coumarin fluorophore on CLPE wobbles in a 21° cone at w0 ) 6.4. This result indicates that while the CLPE fluorophore is immobilized in the lecithin/cyclohexane micelles, the structure of the lecithin/ benzene micelle interface allows substantially greater motion of the CLPE fluorophore. This suggests significantly less order at the lecithin/water interface in the benzene/lecithin/water reverse micelles than in the cyclohexane/lecithin/water system. The solvation dynamics in the benzene/lecithin/water reverse micelles at the same hydration level were measured, and the results are given in Table 1. Figure 5 contrasts the response for the dynamics in the w0 ) 6.4 reverse micelles for both water/ lecithin/cyclohexane and water/lecithin/benzene systems. The most notable feature of these decays is that the relaxation in the reverse micelles in benzene appears significantly faster than that in the cyclohexane reverse micelles.

One explanation for the differences we measured for the benzene system is the formation of a water pool in the reverse micellar interior. The tubular reverse micelles formed in cyclohexane do not support water droplet formation, while the spherical reverse micelles formed in benzene should allow water droplet formation in the micellar interior. We have attributed the appearance of an ultrafast solvation dynamics response in AOT reverse micelles to the formation of a bulklike water pool. This interpretation is strongly supported by the recent calculations of Faeder and Ladanyi, who observe a bulklike water pool forming for w0 ) 5 in a model AOT reverse micelle.13 We also observe faster solvation dynamics in spherical reverse micelles formed with the nonionic surfactants.27 The presence of a bulklike water pool in the micellar interior could lead to bulklike ultrafast water relaxation. Another explanation for faster dynamics we observe in the benzene/lecithin/water reverse micelles over the cyclohexane/ lecithin/water systems is a higher degree of disorder in the benzene/lecithin interface. Other studies suggest that benzene may reside at the surfactant interface, acting as a cosurfactant for the reverse micelles.2 In this case, benzene intercalation between lecithin molecules could disrupt the order or increase the spacing between headgroups, leading to less strongly bound water at the benzene interface because the interfacial order is reduced. This interpretation is supported by our time-resolved fluorescence anisotropy data. It is likely that the response we measure is a combination of these two effects. On the basis of previous studies by other workers, the differences we observe between benzene and cyclohexane reverse micelles are surprising. In other studies, researchers measured the properties of a particular chemical moiety within the reverse micelle interior and observed how each property changed with hydration. In general, those studies find that the properties of the particular chemical moiety are insensitive to the nonpolar solvent in which the reverse micelle is prepared. For example, 31P NMR studies of lecithin reverse micelles in both benzene5,18 and cyclohexane8,35 show that phosphorus mobility steadily increases with hydration. Likewise, the IR spectra of headgroup PdO and CdO stretching frequencies show increased water interaction with increasing hydration for both benzene and cyclohexane reverse micelles.3,9,35,38 Also, the OH stretching of water behaves similarly with increasing hydration.2,38 While some of these studies report differences at higher hydration levels in the cyclohexane reverse micelles, at the w0 ) 6.4 hydration level, these studies predict similar behavior for the reverse micelles in cyclohexane and benzene.

11080 J. Phys. Chem. B, Vol. 104, No. 47, 2000 Previous reports have denoted and quantified different water types within the reverse micelle interiorsthat is, bound or free. Estimating the proportion of free water within w0 ) 6.4 reverse micelles on the basis of our dynamical results shows that IR and NMR predictions match the reverse micelles in benzene much better than in cyclohexane. On the basis of previous results,44 we estimate that only about 1 water molecule per headgroup in cyclohexane reverse micelles can be considered free, bulklike water at the w0 ) 6.4 hydration level. Conversely, solvation dynamics results for benzene reverse micelles indicate that 60% of the solvent relaxation, or about 3-4 water molecules per headgroup, occurs on the subpicosecond time scale and can be attributed to “free” water. The evidence for this quantity of free water in water/lecithin/benzene reverse micelles conflicts with the results of other studies. Several other studies concluded that free water does not appear in benzene reverse micelles until hydration levels exceed w0 ) 8-10.9,18,42 Moreover, the differences we observe between the cyclohexane and benzene systems contradicts the results of previous work by others. Comparing their IR and NMR results, Maitra et al.2 estimate that the fraction of bound and free water molecules are virtually equal in benzene and cyclohexane reverse micelles at the w0 ) 6.4 hydration level. We believe that these differences can be attributed to the methods used to evaluate the water. In previous work, we have shown that results from IR spectroscopy disagree with our solvation dynamics results.30 Because NMR and IR experiments reflect shifts attributed to hydrogen bonding, they may not be sensitive to the immobilization observed for timeresolved solvation dynamics experiments. In summary, we have synthesized an amphiphilic probe molecule that has allowed us to investigate the solvation dynamics of water in benzene/lecithin/water reverse micelles. The CLPE probe in cyclohexane/lecithin/water produces indistinguishable solvation dynamics and anisotropy decays as the starting dye molecule C343. However, a comparison of the benzene-based reverse micelles at higher hydration levels shows that the solvation dynamics are significantly faster than in the cyclohexane gel at the same hydration level. These results are similar to our previous results, in which the dynamical response of a solvent confined within reverse micelles cannot be reliably predicted from NMR and IR data. Furthermore, the dynamical response of confined water appears to be highly sensitive to the morphology of the reverse micelles and the penetration of the nonpolar solvent into the interface. Acknowledgment. The authors thank Dr. Thomas A. Wynn, Dr. Jeffrey D. Kahl, Dr. Dustin McMinn, Mr. Joseph P. Bullock, Mr. Paul Gansle, Dr. Erik Kuester, and Prof. David W. Grainger at Colorado State University and Dr. Steve W. Burgess at Avanti Technical Support for their patient assistance with the CLPE synthesis. The authors thank Profs. E. R. Fisher, L. S. Hegedus, and S. H. Strauss for lending us equipment that made the synthesis and characterization possible. Support for this project came from the National Science Foundation References and Notes (1) Reagent spots were revealed with a phosphate stain and ninhydrin reagent: (a) Ellingson, J. S.; Lands, W. E. M. Lipids 1968, 3, 111-120. (b) Skipski, V. P.; Peterson, R. F.; Barclay, M. J. Lipid Res. 1962, 3, 467. (2) Maitra, A.; Jain, T. K.; Shervani, Z. Colloid Surf. 1990, 47, 25567. (3) Arcoleo, V.; Aliotta, F.; Goffredi, M.; La Manna, G.; Turco Liveri, V. Mater. Sci. Eng. C 1997, 5, 47-53. (4) Barbara, P. F.; Walker, G. C.; Kang, T. J.; Jarzeba, W. Proc. SPIEInt. Soc. Opt. Eng. 1990, 1209, 18-31.

Willard and Levinger (5) Boicelli, C. A.; Conti, F.; Giomini, M.; Giuliani, A. M. Chem. Phys. Lett. 1982, 89, 490-5. (6) Boicelli, C. A.; Giomini, M.; Giuliani, A. M. Appl. Spectrosc. 1984, 38, 537-9. (7) Brucker, G. A.; Kelley, D. F. J. Phys. Chem. 1987, 91, 28562861. (8) Capitani, D.; Rossi, E.; Segre, A. L.; Giustini, M.; Luisi, P. L. Langmuir 1993, 9, 685-689. (9) Davenport, J. B.; Fisher, L. R. Chem. Phys. Lipids 1975, 14, 27590. (10) De, T.; Maitra, A. AdV. Colloid Interface Sci. 1995, 59, 95-193. (11) Drexhage, K. H.; Erikson, G. R.; Hawks, G. H.; Reynolds, G. A. Opt. Comm. 1975, 15, 399-403. (12) Enders, A.; Nimtz, G. Ber. Bunsen-Ges. Phys. Chem. 1984, 88, 512-517. (13) Faeder, J.; Ladanyi, B. M. J. Phys. Chem. B 2000, 104, 10331046. (14) Fendler, J. H. Membrane Mimetic Chemistry: Characterizations and Applications of Micelles, Microemulsions, Monolayers, Bilayers, Vesicles, Host-Guest Systems, and Polyions; Wiley-Interscience: New York, 1982. (15) Finer, E. G. J. Chem. Soc., Faraday Trans. 2 1973, 69, 15901600. (16) Jain, T. K.; Varshey, M.; Maitra, A. J. Phys. Chem. 1989, 93, 74097416. (17) Kinosita, K., Jr.; Kawato, S.; Ikegami, A. Biophys. J. 1977, 20, 289-305. (18) Klose, G.; Arnold, K.; Peinel, G.; Binder, H.; Gawrisch, K. Colloids Surf. 1985, 14, 21-30. (19) Klosgen, B.; Reichle, C.; Kohlsmann, S.; Kramer, K. D. Biophys. J. 1996, 71, 3251-3260. (20) Krishnakumar, S.; Somasundaran, P. J. Coll. Interface Sci. 1994, 162, 425-430. (21) Lakowicz, J. R.; Hogen, D. Biochemistry 1981, 20, 1366-1373. (22) Mandal, D.; Datta, A.; Pal, S. K.; Bhattacharyya, K. J. Phys. Chem. B 1998, 102, 9070-9073. (23) Maroncelli, M. J. Mol. Liq. 1993, 57, 1-37. (24) Pal, S. K.; Mandal, D.; Sukul, D.; Bhattacharyya, K. Chem. Phys. Lett. 1999, 312, 178-184. (25) Pal, S. K.; Sukul, D.; Mandal, D.; Bhattacharyya, K. J. Phys. Chem. B 2000, 104, 4529-4531. (26) Pant, D.; Levinger, N. E. J. Phys. Chem. B 1999, 103, 7846-7852. (27) Pant, D.; Levinger, N. E. Langmuir, in press. (28) Pant, D.; Riter, R. E.; Levinger, N. E. J. Chem. Phys. 1998, 109, 9995-10003. (29) Reynolds, G. A.; Drexhage, K. H. Opt. Comm. 1975, 13, 222225. (30) Riter, R. E.; Undiks, E. P.; Kimmel, J. R.; Levinger, N. E. J. Phys. Chem. B 1998, 102, 7931-7938. (31) Riter, R. E.; Undiks, E. P.; Levinger, N. E. J. Am. Chem. Soc. 1998, 120, 6062-6067. (32) Riter, R. E.; Willard, D. M.; Levinger, N. E. J. Phys. Chem. B 1998, 102, 2705-2714. (33) Sarkar, N.; Das, K.; Datta, A.; Das, S.; Bhattacharyya, K. J. Phys. Chem. 1996, 100, 10523-10527. (34) Schurtenberger, P.; Scartazzini, R.; Magid, L. J.; Leser, M. E.; Luisi, P. L. J. Phys. Chem. 1990, 94, 3695-3701. (35) Shervani, Z.; Jain, T. K.; Maitra, A. Colliod Polym. Sci. 1991, 269, 720-726. (36) Takahashi, A.; Takizawa, T.; Nakata, Y. J. Phys. Soc. Jpn. 1996, 65, 635-642. (37) Tominaga, K.; Walker, G. C. J. Photochem. Photobiol. A 1995, 87, 127-133. (38) Tsai, Y.-S.; Ma, S.-M.; Kamaya, H.; Ueda, I. Mol. Pharmacol. 1987, 31, 623-630. (39) Volke, F.; Eisenblaetter, S.; Galle, J.; Klose, G. Chem. Phys. Lipids 1994, 70, 121-131. (40) Wachtel, E.; Federman, S.; Greenspoon, N. Israel J. Chem. 1992, 32, 113-9. (41) Walde, P.; Giuliani, A. M.; Boicelli, C. A.; Luisi, P. L. Chem. Phys. Lipids 1990, 53, 265-88. (42) Walter, W. V.; Hayes, R. G. Biochim. Biophys. Acta 1971, 249, 528-38. (43) Hodges, M. W.; Cafiso, D. S.; Polnaszek, C. F.; Lester, C. C.; Bryant, R. G. Biophys. J. 1997, 73, 2575-2579. (44) Willard, D. M.; Riter, R. E.; Levinger, N. E. J. Am. Chem. Soc. 1998, 120, 4151-4160. (45) Carey, F. A.; Sundberg, R. J. AdVanced Organic Chemistry, Part B: Reactions and Synthesis, 3rd ed.; Plenum Press: New York, 1990.