Ultrafast Energy Migration Pathways in Self-Assembled Phospholipids

ARTICLE pubs.acs.org/JPCA

Ultrafast Energy Migration Pathways in Self-Assembled Phospholipids Interacting with Confined Water Nancy E. Levinger,*,†,‡ Rene Costard,† Erik T. J. Nibbering,† and Thomas Elsaesser† † ‡

Max Born Institut f€ur Nichtlineare Optik und Kurzzeitspektroskopie, D-12489 Berlin, Germany Department of Chemistry, Colorado State University, Fort Collins, Colorado 80523-1872, United States

bS Supporting Information ABSTRACT: Phospholipids self-assembled into reverse micelles in benzene are introduced as a new model system to study elementary processes relevant for energy transport in hydrated biological membranes. Femtosecond vibrational spectroscopy gives insight into the dynamics of the antisymmetric phosphate stretching vibration νAS(PO2)
, a sensitive probe of local phosphate
water interactions and energy transport. The decay of the νAS(PO2)
mode with a 300-fs lifetime transfers excess energy to a subgroup of phospholipid low-frequency modes, followed by redistribution among phospholipid vibrations within a few picoseconds. The latter relaxation is accelerated by adding a confined water pool, an efficient heat sink in which the excess energy induces weakening or breaking of water
water and water
phospholipid hydrogen bonds. In parallel to vibrational relaxation, resonant energy transfer between νAS(PO2)
oscillators delocalizes the initial excitation.

’ INTRODUCTION Lipid molecules form a critical part of membranes found in all living systems. Self-assembly of these molecules into organized bilayers provides a framework that delineates cellular and subcellular compartments. Model lipid bilayers have been explored to infer information about structure and function of biological membranes. The amphiphilic nature of the lipid molecules allows them to self-assemble not only into membranes but also into a wide variety of structures, such as reverse micelles, small particles that stabilize a pool of water in nonpolar solvents.1 Reverse micelles generally form when amphiphiles stabilize water in nonpolar environments. A well-chosen reverse micelle system allows precise control of hydration through the molar ratio of water to amphiphile, w0 = [H2O]/[amphiphile]. In this way, local molecular interactions and processes that are relevant in biological membranes can be studied under well-defined conditions in a wide hydration range. Although reverse micelles commonly form in synthetic systems, such structures can also exist transiently in lipid bilayers.2 Many studies have explored reverse micelles formed from various components and have interpreted results relative to biological membranes. The most common reverse micelles utilize sodium dioctylsulfosuccinate (AOT) as the surfactant.3,4 Here we report studies of dioleoylphosphatidylcholine (DOPC) molecules self-assembled into phospholipid reverse micelles as a model system for phosphate
water interactions. The interaction of water molecules with ionic phosphate groups is of particular interest as phosphate groups are primary hydration sites in membranes and in biomolecules such as DNA and RNA. Significantly less is known about reverse micelles formed r 2011 American Chemical Society

from phospholipids compared to the ones formed from AOT. Several groups report the formation of isolated water droplets stabilized by lecithin, a mixture of naturally occurring phosphatidylcholines, in alkanes,5
12 haloalkanes,13 and benzene.6,14
19 In alkane solvents, lecithin tends to form a gel phase with added water.5
12 In contrast, droplets remain isolated when reverse micelles form in benzene.6,14
19 Although no reports of DOPC reverse micelles have appeared, DOPC forms a significant component of the naturally occurring lecithin phospholipid so we expect strong similarities between DOPC and lecithin reverse micelles. As it has frequently been used in studies of vesicles20
22 or multibilayer films23
31 as models of membranes or bilayers, employing DOPC facilitates direct comparison of our results to these systems. Aqueous systems, including reverse micelles, are characterized by structural fluctuations and energy dissipation processes occurring in the femto- to picosecond time range.32
34 This behavior arises from the interplay of long-range Coulomb interactions, local hydrogen bonds, and anharmonic vibrational couplings. Nonlinear vibrational spectroscopy with femtosecond time resolution has developed into a leading technique for studying the dynamics of such processes and discerning the different interactions.32 In reverse micelles, such methods have focused on OH- and OD-stretch excitations of water molecules.34
40 Both neat and isotopically diluted water (HOD) have been studied in AOT micelles of different size to distinguish the properties of Received: June 28, 2011 Revised: September 9, 2011 Published: September 19, 2011 11952

dx.doi.org/10.1021/jp206099a | J. Phys. Chem. A 2011, 115, 11952–11959

The Journal of Physical Chemistry A water molecules interacting directly with the polar head groups from those inside the water pool. The dynamics of vibrational dephasing, molecular reorientation, and vibrational relaxation display pronounced dependence on the micelle size, reflecting changes in the fluctuating water structure. In general, increasing rearrangement times of the hydrogen bond network occur with decreasing micelle size. In contrast to the OH- and OD-stretching vibrations, other reverse micelle vibrations have received far less attention.41
43 As far as we are aware, only Dlott and coworkers44,45 have investigated dynamics of lower frequency water vibrations or vibrations associated with the surfactant. Here we present the first ultrafast dynamics study of phosphate vibrations in a phospholipid assembly. Phosphate vibrations are highly sensitive to both local hydrogen bond interactions and electronic polarization effects originating from long-range Coulomb interactions.46
49 The antisymmetric phosphate stretching vibration is an excellent probe of energy relaxation and migration and of local hydrogen bonding in DOPC micelles. The femto- to picosecond dynamics of this mode as observed through temporally and spectrally resolved pump
probe measurements provide specific insight into the pathways of relaxation in self-assembled lipids and their interactions with the water pool.

’ EXPERIMENTAL TECHNIQUES Sample Preparation and Characterization. Reverse micelle samples were prepared from dioleoylphosphatidylcholine (DOPC, Avanti Polar Lipids), benzene (Sigma Aldrich, anhydrous 99.8%), and water (Acros, deionized reagent grade). Benzene and water were used as received. Prior to use, DOPC was dried under a vacuum over P2O5 overnight to minimize the inherent water in the sample. A 0.25 M stock solution of DOPC in benzene was prepared. Appropriate amounts of water measured by mass were added to the stock solution to create samples with w0 = 2, 4, 8, and 16. All solutions were stored at
20
C until use. Infrared spectroscopy (Varian 640-IR) characterized the samples. Samples were loaded into liquid cuvettes (Harrick) with 1 mm thick BaF2 windows and a 25 μm path length. This yielded absorbances of approximately 0.5
0.7 in the phosphate antisymmetric stretching region of the spectrum, near 1250 cm
1. Fresh samples were loaded into the IR cuvette for each timeresolved measurement. Sample spectra exhibited no changes before and after laser experiments. Spectra of diluted dry samples presented no significant changes for the position of the phosphate peak compared to the 0.25 M DOPC samples confirming that phosphate moieties in the reverse micelle aggregates do not interact with each other leading to collective excitation. In preliminary experiments (Stanford University, Colorado State University) we determined the size of DOPC reverse micelles formed in benzene using dynamic light scattering (DLS, Nanosizer, Malvern; Dynapro Titan, Wyatt). From diameters measured for w0 = 5 and 16.5 (46.5 ( 1.5 and 78.5 ( 5 Å, respectively), we extrapolate sizes (in Å) for reverse micelles formed here using dH = 2.8w0 + 32.6. This suggests that the average length of the DOPC in the reverse micelles is 32.6/2 = 16.3 Å. This is slightly smaller than the length indicated from DOPC lipid bilayers of 34.8/2 = 17.4 Å.24,25,28 Table 1 shows an estimation of the reverse micelle size for the particles examined here, an estimation of the number of lipid molecules comprising each aggregate and an estimation of the number of water

ARTICLE

Table 1. Sizes and Molecular Aggregation Estimates for DOPC/Benzene Reverse Micellesa w0

dH (Å)

no. DOPC

no. H2O

0

35

55

0) transitions. Relaxation of the v = 1 population with a 300 fs lifetime (part (2) Figure 1b) results in a decay in ΔA with minor changes of their spectral envelopes. The v = 1 decay brings the excited νAS(PO2)
oscillator back to its ground state and populates a group of low-lying vibrational modes (orange states Figure 1b). This process is connected with a redistribution of excess energy and the formation of a “hot” ground state (part (2) of Figure 1b). The fact that the lifetime is the same for all w0 values, i.e., hydration levels, indicates a predominant relaxation into the vibrational manifold of the DOPC molecule and a negligible role of water for this initial fast relaxation. A similar intramolecular relaxation has been observed for νAS(PO2)
in DNA oligomers.48 Although we do not know the specific relaxation pathways of the v = 1 state of νAS(PO2)
in detail, we expect contributions from off-diagonal anharmonic couplings to combination bands and overtones of phosphate low-frequency modes. For instance, a combination band involving quanta of the PO2 twisting mode at 430 cm
1 and the diester-PO2 stretching mode around 800 cm
1 lies at an energy close to the v = 1 state of νAS(PO2)
and could serve as primary energy acceptor.26,51,52 Anharmonic coupling between low-frequency DOPC modes that are populated in the hot ground state and νAS(PO2)
modifies the potential energy surface leading to states v0 = 0 and v0 = 1 of the νAS(PO2)
oscillator (part (2) of Figure 1b). This mechanism causes a spectral shift and reshaping of the νAS(PO2)
absorption A0 (t) on the v0 = 0
1 transition that are proportional to the momentary population of the low-frequency modes and, in turn, make the νAS(PO2)
oscillator a probe of 11955

dx.doi.org/10.1021/jp206099a |J. Phys. Chem. A 2011, 115, 11952–11959

The Journal of Physical Chemistry A

ARTICLE

Figure 4. Time-resolved absorbance changes for w0 = 0, 2, 4, 8, 16 DOPC reverse micelles. The isotropic absorbance change ΔAiso(t) = (ΔA||(t) + 2ΔA^(t))/3 measured at fixed probe frequencies is plotted as a function of pump
probe delay (symbols). For each w0, the frequency positions are at the respective maximum of the transient v = 1 to 2 absorption (cf. Figure 3, blue symbols) and the maximum bleaching of the v = 0
1 transition (red symbols). Signals around delay zero are affected by contributions from nonresonant nonlinear effects. Solid lines: Numerical fits of the transient from which the time constants and amplitudes given in Table 2 have been derived.

Table 2. Time Constants τ1, τ2 and Relative Amplitudes A1/ A2 of the Two Kinetic Components Derived from Numerical Fits of the Time-Resolved Pump
Probe Tracesa w0

τ1

τ2

A1/A2

0

300 fs

1.5 ps

1.4

2

300 fs

1.3 ps

3.2

4

300 fs

1.2 ps

3.9

8

300 fs

1.0 ps

4.3

16

300 fs

1.1 ps

5.7

a The uncertainty of τ1 is of the order of (10% (all w0); the uncertainty of τ2 is between (10% (w0 = 0) and (30% (w0 = 16).

their time evolution.53 The transient spectra at a 1 ps delay (Figure 3) display a positive differential absorption at low and a negative differential absorption at high probe frequencies, corresponding to a red-shift of A0 (t) absorption, i.e., the v0 = 0
1 transition, of up to 40 cm
1. In the w0 = 0 sample, the subsequent population decay of the low-frequency modes transfers energy into other vibrations in the reverse micelle system, that is, additional modes in the original DOPC, modes on adjacent DOPC molecules, or into the surrounding benzene. The 1.5 ps decay time of the νAS(PO2)
absorption changes reflects this process. We observe small residual signals on time scales up to 500 ps suggesting that on this long time scale a ground state with

a common elevated temperature of the solute DOPC and the solvent benzene exists. A strikingly different behavior is found when the reverse micelles are large enough to form a water pool as in the w0 = 16 reverse micelles.38 Now, the differential spectra for t > 1 ps (Figure 3j) display a negative ΔA at low and a positive ΔA at high frequencies, corresponding to a blue-shift of A0 (t) and the v0 = 0
1 transition. Low-frequency modes of the water pool, in particular librations, hydrogen bond vibrations, and translations, act as an additional energy accepting manifold into which part of the excess energy flows (part (3) of Figure 1b). This energy quickly randomizes in the water pool and establishes an elevated vibrational temperature. This hot water ground state which has been studied in different aqueous systems is characterized by a larger fraction of weakened and/or broken intermolecular hydrogen bonds,42 both between water molecules and between water molecules and phosphate groups. Studies of hydrated DNA in which the water shell was heated selectively via femtosecond OH stretching excitation have shown that the νAS(PO2)
v0 = 0
1 absorption undergoes a blue shift upon heating.48 The same behavior is found here for the DOPC micelle system with a very similar blue-shift of the order of 30 cm
1. In addition to this second channel for vibrational energy relaxation, we note that the relative amplitude of the longer time signals decreases significantly for the w0 = 16 sample compared to w0 = 0, and the decay time shortens from 1.5 to ∼1 ps. This demonstrates that energy relaxation is more efficient when water fully hydrates the phosphate groups. In reverse micelles, we can easily control the amount of water present. This differs from other self-assembled lipid systems such as bilayers where control and precise measurement of the hydration level are much more difficult.50 At the lowest hydration levels w0 = 0 and 2, the system lacks a bulklike water bath and low-frequency modes of the DOPC molecules serve as the primary receptor for energy deposited in vibrational excitation of the νAS(PO2)
mode. At higher hydration levels, a bulklike water pool is formed in the reverse micelle interior, leading to faster and more efficient vibrational energy relaxation. This demonstrates the importance that water plays in accepting this energy and funneling it away from the phospholipid reverse micellar interface, a mechanism most likely also important for biological membranes. B. Energy Transfer between Phosphate Oscillators. In addition to vibrational relaxation, we have explored orientational randomization of νAS(PO2)
excitations in the samples via the pump
probe anisotropy, r(t). Figure 5 presents a logarithmic plot of r(t) as a function of delay time for DOPC micelles with w0 = 0, 2, and 16. The inset of Figure 5 shows parallel and perpendicular decays for w0 = 2 along with measured and calculated isotropic signals that agree well with each other. We observe an initial r(0) ≈ 0.4 for all hydration levels indicating a random distribution of transition dipoles. For w0 = 0 samples, the anisotropy decays almost completely by 4 ps and fits well to a single exponential decay with a time constant of 2 ps (dashed line). Measuring the anisotropy up to ten times the 300 fs lifetime of νAS(PO2)
is facilitated by the longer-lived thermal component of the bleach recovery. At higher water concentrations, this component is weaker, resulting in a shorter time interval over which r(t) can be derived. However, anisotropy traces in Figure 5 clearly show that the r(t) decay gets slower for increasing water concentrations, and time constants of 3 ps (w0 = 2) and 6 ps (w0 = 16) are estimated. 11956

dx.doi.org/10.1021/jp206099a |J. Phys. Chem. A 2011, 115, 11952–11959

The Journal of Physical Chemistry A

Figure 5. Time-resolved anisotropy decays, r(t) for νAS(PO2)
in w0 = 0, 2, and 16 DOPC reverse micelles as a function of delay time between excitation and probe pulses (solid lines). The fast vibrational relaxation for w0 = 16 samples limits our ability to measure the signals past 1.5 ps. Dashed lines: Single exponential decays with time constants of 2 ps (w0 = 0), 3 ps (w0 = 2), and 6 ps (w0 = 16). The inset shows the transient absorption decays for νAS(PO2)
excitation and probe pulses parallel (black symbols), perpendicular (red), and at the magic angle (blue) with respect to each other in w0 = 2 DOPC reverse micelles. The isotropic signal calculated from parallel and perpendicular traces, ΔAiso(t) = (ΔA||(t) + 2ΔA^(t))/3 (black line), matches the magic angle decay.

We consider several different possible mechanisms to account for the anisotropy decays, including (i) rotational relaxation, (ii) local heating and environmental effects, and (iii) F€orster resonance energy transfer (FRET). (i) Rotational reorientation of a molecule is perhaps the most common mechanism associated with anisotropy decays in molecules.54 The phosphate moiety that we probe through νAS(PO2)
exists as a part of a larger molecule (see Figure 1a). We use aqueous phosphate anion, PO43
, as a model system to place a lower limit to the rotational motion of the phosphate in DOPC, estimating the phosphate rotational time using the Debye
Stokes
Einstein expression r(t) = 0.4 exp(
6Drt) with the diffusion coefficient Dr = kBT/(8πηR3), where kB is the Boltzmann constant, T is temperature, and η is the dynamic viscosity (1.002 cP for water).55 In this expression, R is the radius of the sphere approximating the phosphate anion. Assuming that R = 1.5 Å, which is the P
O bond length in PO43
, leads to a rotational diffusion coefficient of 4.5  1010 s
1 and a reorientation time τor = 3.7 ps.54 Because the phosphate in our system comprises part of a larger molecule, this time would be significantly faster than the actual rotational time for the phosphate in DOPC; thus we reject rotational reorientation as the mechanism for anisotropy decay. Note that the time scale for rotational reorientation for the smallest micelle (w0 = 0, dH = 3.5 Å) should be >20 ns assuming η = 0.6055 cP for benzene.56 Thus, neither molecular nor micellar rotational relaxation can account for the randomization of the anisotropy signals observed for any samples. (ii) The impact of heat deposited into the system provides another mechanism that could potentially account for

ARTICLE

randomization of the anisotropic signals we observe. The phosphate group on DOPC, whose antisymmetric stretching mode we probe, resides in a highly perturbative environment in the reverse micelles due to the proximity of nearby charges of its own choline and adjacent surfactant phosphates and cholines. Indeed, the dipole moment induced by the phosphate
choline pair on a single surfactant leads the DOPC molecule to possess an 18.7 D dipole in the headgroup region.31 When the PO2
group on DOPC forms hydrogen bonds with nearby water molecules, the electron density on the oxygen molecules changes,57 and in response, the transition dipole, which lies along the bisector of the O
P
O
bond angle, can be redirected. Theoretical calculations in ref 57 have shown, however, that the O
P
O
bond angle changes by approximately 3
when going from an unhydrated to a fully hydrated (PO2)
group. An even smaller change of the angular geometry is expected for fluctuations of hydrogen bond geometries at a fixed hydration level. Although these effects should occur on a time scale fast enough to appear in our data, changes of dipole orientations by a few degrees translate in changes of r(t) distinctly smaller than 0.1. Additionally, if this mechanism dominated the orientational randomization in the samples, we would expect correlation of r(t) with the heating we observe in the reverse micelles. This fails to account for the anisotropy decays for two reasons: (1) the heating occurs on a longer time scale than the anisotropy decay and (2) water related effects are absent in the smallest micelles. Another possible way that heat flow could impact the anisotropy decay arises if a particular oscillator absorbs a photon and subsequently relaxes, releasing its energy into lower frequency modes. The heat released into the local environment could cause a shift in the vibrational frequency of another nearby phosphate moiety which transiently changes the absorption by the nearby phosphate oscillator whose dipole points in a different direction from the initially excited oscillator. Estimating the related absorption changes caused by this mechanism from the small pump
probe signals at delay times longer than 1 to 2 ps (Figure 4) indicates that these long-term signals are much weaker than the absorption changes caused by the initially excited oscillator and, thus, cannot account for the anisotropy decay within the first 3 ps (Figure 5). (iii) FRET involving hopping of energy from a νAS(PO2)
donor to an acceptor with a random in-plane orientation of their transition dipoles (Figure 1a) could also lead to orientational randomization.58
63 In the simplest approximation, oscillators are modeled as point dipoles with weak coupling between them. For the P
P distance between phosphate groups in lipid bilayers, approximately 6.5 Å,13,24
26,28,31,64
67 we estimate a minuscule 1.5 cm
1 coupling of νAS(PO2)
point dipoles; if the dipoles lie 5 Å apart, the coupling increases to 3.3 cm
1, still clearly quite small. This demonstrates that the DOPC reverse micelle system is in the weak coupling limit, making it a good candidate for FRET. The F€orster theory of energy transfer between point dipoles introduces the F€orster radius R0 as the key parameter describing the donor
acceptor coupling. We discuss this in more detail in the Supporting Information. The coupling strength depends on 11957

dx.doi.org/10.1021/jp206099a |J. Phys. Chem. A 2011, 115, 11952–11959

The Journal of Physical Chemistry A the dielectric properties of the system which—on the simplistic level of F€orster theory—are described by the refractive index at the frequency of the vibrational transition. For reverse micelles, the refractive index is a highly complex quantity because of the ionic groups of the DOPC molecule and the spatial heterogeneity of the DOPC/water/benzene system. For w0 = 16, the phosphate groups of DOPC are fully hydrated by the water pool, and thus, we use the refractive index n = 1.291 of water at 1250 cm
1 to estimate a F€orster radius of 3.22 Å. In the spherical micelle geometry (Figure 1a), each donor is surrounded by six acceptors, resulting in a 6-fold enhancement of the single donor
single acceptor energy transfer rate. Taking a P
P distance of 6.5 Å as in lipid bilayers and the measured donor lifetime of 300 fs, we find an energy transfer time from a phosphate donor to its six nearest neighbors of 3.38 ps. This time scale is of the same order of magnitude as the anisotropy decays in Figure 5. Thus, FRET is a viable mechanism for the anisotropy decay (part (4) of Figure 1b). For micelles with smaller w0, the anisotropy decay becomes faster. Here, a somewhat smaller P
P distance and a reduced screening of donor
acceptor interaction (smaller refractive index) at the reduced water level may play a role. To clarify this issue, independent structural information of the micelle geometry is required. The presence of resonant energy transfer between DOPC molecules in the reverse micelles provides the relevant length and time scales for energy migration. This knowledge allows assessment of the relevance of this process in other biomolecular structures containing phosphate groups. For instance, the phosphate groups in the backbone of DNA and RNA show nearest neighbor distances between approximately 4 and 7 Å. By use of the F€orster radius determined here and the νAS(PO2)
lifetime of 340 fs measured in hydrated DNA oligomers,48 we estimate transfer times between 1.3 and 36 ps. In the backbone of DNA, the orientation of the νAS(PO2)
is well-defined, thus in contrast to the micelle geometry studied here, even a series of transfer steps does not fully randomize the dipole orientations.

’ CONCLUSIONS In conclusion, the results presented establish the time scales, pathways, and mechanisms of vibrational relaxation and energy migration in self-assembled phospholipid (DOPC) micelles. While the lifetime of the antisymmetric (PO2)
stretching vibration of 300 fs is independent of the degree of hydration, the subsequent redistribution of excess energy changes with water content. At low hydration, energy is redistributed mainly within the vibrational manifold of the DOPC molecules. At high hydration, the water pool serves as an efficient heat sink, resulting in an acceleration of energy redistribution and a weakening of hydrogen bonds between the phosphate groups and the first water layer. In parallel to such relaxation processes, resonant energy transfer between the weakly coupled νAS(PO2)
oscillators delocalizes the initial excitation on a time scale of a few picoseconds. Our results underline the potential of phosholipid reverse micelles as a model system for studying basic nonequilibrium processes of biological membranes under controlled and variable conditions. ’ ASSOCIATED CONTENT

bS

Supporting Information. Details of resonant energy transfer between vibrational dipoles. This material is available free of charge via the Internet at http://pubs.acs.org.

ARTICLE

’ AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected].

’ ACKNOWLEDGMENT We gratefully acknowledge Prof. M. D. Fayer (Stanford University) for providing the opportunity to make preliminary measurements of the DOPC reverse micelles. N.E.L. acknowledges Colorado State University for a sabbatical leave and NSF Grant No. CHE 0628260 for funding allowing preliminary studies of the DOPC reverse micelles. The research leading to these results has received funding from the European Research Council under the European Union's Seventh Framework Programme (FP7/2007-2013)/ERC Grant No. 247051. ’ REFERENCES (1) Luisi, P. L.; Giomini, M.; Pileni, M. P.; Robinson, B. H. Biochim. Biophys. Acta 1988, 947, 209–246. (2) van Meer, G.; Voelker, D. R.; Feigenson, G. W. Nat. Rev. Mol. Cell Biol. 2008, 9, 112–124. (3) De, T. K.; Maitra, A. Adv. Colloid Interface Sci. 1995, 59, 95–193. (4) Levinger, N. E.; Swafford, L. A. Annu. Rev. Phys. Chem. 2009, 60, 385–406. (5) Schurtenberger, P.; Magid, L. J.; King, S. M.; Lindner, P. J. Phys. Chem. 1991, 95, 4173–4176. (6) Shervani, Z.; Maitra, A.; Jain, T. K.; Dinesh. Colloids Surf. 1991, 60, 161–173. (7) Schurtenberger, P.; Cavaco, C. Langmuir 1994, 10, 100–108. (8) Shchipunov, Y. A.; Shumilina, E. V. Colloid J. 1996, 58, 117–125. (9) Jerke, G.; Pedersen, J. S.; Egelhaaf, S. U.; Schurtenberger, P. Phys. Rev. E 1997, 56, 5772–5788. (10) Angelico, R.; Palazzo, G.; Olsson, U.; Ambrosone, L.; Ceglie, A. Structural investigation of lecithin/cyclohexane solutions. In Trends in Colloid and Interface Science XIII (Progress in Colloid and Polymer Science); Tezak, D., Martinis, M., Eds.; Springer-Verlag: Berlin, 1999; Vol. 112; pp 1
4. (11) Tung, S. H.; Huang, Y. E.; Raghavan, S. R. J. Am. Chem. Soc. 2006, 128, 5751–5756. (12) Lee, H. Y.; Diehn, K. K.; Ko, S. W.; Tung, S. H.; Raghavan, S. R. Langmuir 2010, 26, 13831–13838. (13) Grdadolnik, J.; Kidric, J.; Hadzi, D. Chem. Phys. Lipids 1991, 59, 57–68. (14) Davenport, J. B.; Fisher, L. R. Chem. Phys. Lipids 1975, 14, 275–290. (15) Kumar, V. V.; Kumar, C.; Raghunathan, P. J. Colloid Interface Sci. 1984, 99, 315–323. (16) Walde, P.; Giuliani, A. M.; Boicelli, C. A.; Luisi, P. L. Chem. Phys. Lipids 1990, 53, 265–288. (17) Schurtenberger, P.; Peng, Q.; Leser, M. E.; Luisi, P. L. J. Colloid Interface Sci. 1993, 156, 43–51. (18) Willard, D. M.; Levinger, N. E. J. Phys. Chem. B 2000, 104, 11075–11080. (19) Milhaud, J.; Bouchemal, N.; Rog, T.; Hantz, E. Chemphyschem 2010, 11, 590–598. (20) Moyano, F.; Silber, J. J.; Correa, N. M. J. Colloid Interface Sci. 2008, 317, 332–345. (21) Veatch, S. L.; Keller, S. L. Biophys. J. 2003, 85, 3074–3083. (22) Gilat, T.; Somjen, G. J. Biochim. Biophys. Acta 1996, 1286, 95–115. (23) Pohle, W.; Selle, C.; Fritzsche, H.; Binder, H. Biospectroscopy 1998, 4, 267–280. (24) Tristram-Nagle, S.; Petrache, H. I.; Nagle, J. F. Biophys. J. 1998, 75, 917–925. 11958

dx.doi.org/10.1021/jp206099a |J. Phys. Chem. A 2011, 115, 11952–11959

The Journal of Physical Chemistry A (25) Costigan, S. C.; Booth, P. J.; Templer, R. H. Biochim. Biophys. Acta 2000, 1468, 41–54. (26) Binder, H.; Pohle, W. J. Phys. Chem. B 2000, 104, 12039–12048. (27) Binder, H. Eur. Biophys. J. Biophys. Lett. 2007, 36, 265–279. (28) Caracciolo, G.; Pozzi, D.; Caminiti, R. Appl. Phys. Lett. 2007, 90, 183901–183903. (29) El Kirat, K.; Dufrene, Y. F.; Lins, L.; Brasseur, R. Biochemistry 2006, 45, 9336–9341. (30) Volkov, V. V.; Palmer, D. J.; Righini, R. J. Phys. Chem. B 2007, 111, 1377–1383. (31) Orsi, M.; Michel, J.; Essex, J. W. J. Phys. Condens. Matter 2010, 22, 155106. (32) Nibbering, E. T. J.; Elsaesser, T. Chem. Rev. 2004, 104, 1887–1914. (33) Bakker, H. J.; Skinner, J. L. Chem. Rev. 2010, 110, 1498–1517. (34) Fayer, M. D.; Levinger, N. E. Ann. Rev. Anal. Chem. 2010, 3, 89–107. (35) Patzlaff, T.; Janich, M.; Seifert, G.; Graener, H. Chem. Phys. 2000, 261, 381–389. (36) Seifert, G.; Patzlaff, T.; Graener, H. Phys. Rev. Lett. 2002, 88, 147402. (37) Tan, H. S.; Piletic, I. R.; Riter, R. E.; Levinger, N. E.; Fayer, M. D. Phys. Rev. Lett. 2005, 94, 057405. (38) Piletic, I. R.; Moilanen, D. E.; Spry, D. B.; Levinger, N. E.; Fayer, M. D. J. Phys. Chem. A 2006, 110, 4985–4999. (39) Dokter, A. M.; Woutersen, S.; Bakker, H. J. Proc. Natl. Acad. Sci. U. S. A. 2006, 103, 15355–15358. (40) Cringus, D.; Bakulin, A.; Lindner, J.; Vohringer, P.; Pshenichnikov, M. S.; Wiersma, D. A. J. Phys. Chem. B 2007, 111, 14193–14207. (41) Ashihara, S.; Huse, N.; Espagne, A.; Nibbering, E. T. J.; Elsaesser, T. Chem. Phys. Lett. 2006, 424, 66–70. (42) Ashihara, S.; Huse, N.; Espagne, A.; Nibbering, E. T. J.; Elsaesser, T. J. Phys. Chem. A 2007, 111, 743–746. (43) Huse, N.; Ashihara, S.; Nibbering, E. T. J.; Elsaesser, T. Chem. Phys. Lett. 2005, 404, 389–393. (44) Deak, J. C.; Pang, Y. S.; Sechler, T. D.; Wang, Z. H.; Dlott, D. D. Science 2004, 306, 473–476. (45) Pang, Y.; Deak, J. C.; Huang, W. T.; Lagutchev, A.; Pakoulev, A.; Patterson, J. E.; Sechler, T. D.; Wang, Z. H.; Dlott, D. D. Int. Rev. Phys. Chem. 2007, 26, 223–248. (46) Falk, M.; Hartman, K. A.; Lord, R. C. J. Am. Chem. Soc. 1963, 85, 387–391. (47) Klahn, M.; Mathias, G.; Kotting, C.; Nonella, M.; Schlitter, J.; Gerwert, K.; Tavan, P. J. Phys. Chem. A 2004, 108, 6186–6194. (48) Szyc, L.; Yang, M.; Elsaesser, T. J. Phys. Chem. B 2010, 114, 7951–7957. (49) Tsuboi, M. J. Am. Chem. Soc. 1957, 79, 1351–1354. (50) Gauger, D. R.; Andrushchenko, V. V.; Bour, P.; Pohle, W. Anal. Bioanal. Chem. 2010, 398, 1109–1123. (51) Guan, Y. F.; Wurrey, C. J.; Thomas, G. J. Biophys. J. 1994, 66, 225–235. (52) Prescott, B.; Steinmetz, W.; Thomas, G. J. Biopolymers 1984, 23, 235–256. (53) Hamm, P.; Ohline, S. M.; Zinth, W. J. Chem. Phys. 1997, 106, 519–529. (54) Fleming, G. R.; Morris, J. M.; Robinson, G. W. Chem. Phys. 1976, 17, 91–100. (55) Yaws, C. L. Yaws’ Handbook of Thermodynamic and Physical Properties of Chemical Compounds; Knovel, 2003. (56) CRC Handbook of Chemistry and Physics, 91st ed.; CRC: Boca Raton, FL, 2011. (57) Mrazkova, E.; Hobza, P.; Bohl, M.; Gauger, D. R.; Pohle, W. J. Phys. Chem. B 2005, 109, 15126–15134. (58) Forster, T. Ann. Phys.-Berlin 1948, 2, 55–75. (59) Forster, T. Z. Naturfors. Sect. A-J. Phys. Sci. 1949, 4, 321–327. (60) Eisenthal, K. B. Chem. Phys. Lett. 1970, 6, 155–157. (61) Woutersen, S.; Bakker, H. J. Nature 1999, 402, 507–509. (62) Piatkowski, L.; Eisenthal, K. B.; Bakker, H. J. Phys. Chem. Chem. Phys. 2009, 11, 9033–9038.

ARTICLE

(63) Timmer, R. L. A.; Bakker, H. J. J. Phys. Chem. A. 2010, 114, 4148–4155. (64) Lopez, C. F.; Nielsen, S. O.; Klein, M. L.; Moore, P. B. J. Phys. Chem. B 2004, 108, 6603–6610. (65) Rosso, L.; Gould, I. R. J. Comput. Chem. 2008, 29, 24–37. (66) Saiz, L.; Klein, M. L. Biophys. J. 2001, 81, 204–216. (67) Stepniewski, M.; Bunker, A.; Pasenkiewicz-Gierula, M.; Karttunen, M.; Rog, T. J. Phys. Chem. B 2010, 114, 11784–11792. (68) Kaindl, R. A.; Wurm, M.; Reimann, K.; Hamm, P.; Weiner, A. M.; Woerner, M. J. Opt. Soc. Am. B 2000, 17, 2086–2094.

11959

dx.doi.org/10.1021/jp206099a |J. Phys. Chem. A 2011, 115, 11952–11959