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Nonsynchronicity Phenomenon Observed during the Lamellar-Micellar Phase Transitions of 1-Stearoyllysophosphatidylcholine Dispersed in Water Fu-Gen Wu, Nan-Nan Wang, Ji-Sheng Yu, Jun-Jie Luo, and Zhi-Wu Yu* Key Laboratory of Bioorganic Phosphorous Chemistry and Chemical Biology (Ministry of Education), Department of Chemistry, Tsinghua UniVersity, Beijing 100084, P. R. China ReceiVed: NoVember 10, 2009; ReVised Manuscript ReceiVed: December 26, 2009

Knowledge on the synchronicity or cooperativity of changes in different parts of the amphiphilic molecules is important to understand the molecular mechanisms of phase transformations of self-assembled aggregates. A long-standing and challenging question is to understand the roles individual groups/portions in an amphiphilic molecule play during phase transitions. To address this question, we selected a lysophospholipid, 1-stearoyllysophosphatidylcholine (SLPC), to study the transition mechanisms between its lamellar phase and micellar phase by using differential scanning calorimetry, small-angle X-ray scattering, Fourier transform infrared spectroscopy, and two-dimensional correlation analysis. It was found that during the lamellar to micellar transition the interfacial CdO groups and the lipid acyl tails change evidently with the former changing a little earlier, but the lipid headgroups remain unchanged in the hydration and conformation state. This means that the head, interface, and tail of SLPC molecules change nonsynchronously. Moreover, the results show that the lamellar to micellar transition is initiated by the interfacial groups. The molecular mechanism of the slow formation kinetics of the lamellar state from the micellar state was also discussed in the context of the nonsynchronicity phenomenon. The markedly different behaviors of the head and interface/tail groups during phase transitions are explained as the retention of the intermolecular attractive forces between the neighboring polar headgroups of the amphiphiles. 1. Introduction Amphiphilic lipids self-assemble into several supramolecular structures when dispersed in water. Cylindrical-shaped lipids, such as the normal biomembrane phosphatidylcholines (PCs) or phosphatidylethanolamines (PEs), favor a planar bilayer, while lipids with a conical or wedge-like shape, such as lysophosphatidylcholines (lysoPCs), tend to form micellar phases.1 LysoPCs have the same headgroup as PCs. Their hydrophobic tails, however, contain only a single acyl chain, and they form water-soluble aggregates. LysoPCs are important metabolites and occur as a minor constituent in various cell membranes. They may play a crucial role in cell proliferation and differentiation.2 At low concentrations, lysoPCs promote cell fusion, whereas at higher concentrations they destabilize membrane structure resulting in morphological changes, permeability property alterations, and ultimately cell lysis.3 It has been shown that lysoPCs, when added to the contacting monolayers of fusing membranes, inhibit the hemifusion observed between lipid vesicles and planar membranes.4 When inserted into planar phospholipid bilayers, the “wedge”shaped lysoPCs promote a micellar positive curvature and induce structural changes and lamellar disruption.3 Interactions of lysoPC with PC,5 PE,3 cholesterol,6 and drugs7 have been investigated. The ability of a lipid molecule to participate in a specific biological/biochemical function is related to its structure. The most widespread self-assembled lyotropic structure is the fluid lamellar (LR) phase. Other lyotropic phase structures include the inverse hexagonal (HII) phase and the cubic phase. These fluid lyotropic phases are of the most direct relevance to the * To whom correspondence should be addressed. Tel.: (+86)10 6279 2492. Fax: (+86)10 6277 1149. E-mail: [email protected].

structure and function of biomembranes. In addition, a large number of stable or metastable lamellar gel or crystal structures are adopted by different lipidsswith perpendicular or tilted chains with respect to the bilayer plane, with interdigitated, partially interdigitated, or noninterdigitated chains from the two leaflets, etc.8–12 Although numerous efforts have been made to study the various aspects of phase transitions, some long-standing and challenging questions still remain. We are curious to know the role of the individual groups/portions of an amphiphilic molecule played during the transformation process from one phase to another. To answer this question, it is important to consider the cooperativity of the change of individual groups/portions of the amphiphilic molecule during the phase transformation process, that is, whether the different groups/portions of the molecule change synchronously or nonsynchronously during the phase transitions. The cooperativity issue opens a broad window for us to challenge important questions including the kinetics, polymorphism, metastability, and reversibility of phase transitions. However, not enough attention has been paid to this issue so far. Inthiscontribution,weselectedalysophospholipid,1-stearoyllysophosphatidylcholine (SLPC) (Figure 1), to study its lamellar to micellar and micellar to lamellar transition mechanisms and, through which, to understand the specific role of the individual groups/portions of the amphiphilic molecule played during the transition processes. Various aspects regarding the lamellar and micellar phases of SLPC molecules have been studied using dynamic light scattering (DLS), differential scanning calorimetry (DSC), freeze-fracture electron microscopy (FFEM), 31P NMR, Raman, fluorescence, and X-ray scattering techniques.13–15 Other than using DSC and small-angle X-ray scattering (SAXS) to characterize the phase transitions, we have developed a strategy

10.1021/jp9107014  2010 American Chemical Society Published on Web 01/21/2010

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Figure 1. Molecular structure of 1-stearoyllysophosphatidylcholine (SLPC).

to examine individual functional groups of the molecule during the phase transformation using time-resolved Fourier transform infrared (FTIR) spectroscopy combined with two-dimensional (2D) correlation analysis. It is found that, during the lamellar to micellar and micellar to lamellar transitions, the headgroups, the interface, and the tail portions of SLPC molecules change nonsynchronously. The discovery of the nonsynchronicity phenomenon in self-assembled structures composed of the medium-sized lipid molecules has profound significance in understanding the nature of the phase transformation processes of amphiphiles. 2. Experimental Section 2.1. Sample Preparation. SLPC was purchased from Avanti Polar Lipids (Birmingham, AL, USA). Double deionized water with a resistivity of 18.2 MΩ cm or D2O (99.9% of deuterium, from Cambridge Isotopes) was used for the preparation of lipid samples. The lipid/water ratio was 1/3 (w/w). Homogeneous lipid dispersion was prepared by repeated thermal cycling between -20 and 50 °C. The initial lamellar phase was obtained by incubating the well-hydrated sample at around 0 °C for 10 days according to the literature.13 2.2. DSC. Calorimetric data were obtained with a differential scanning calorimeter DSC821e equipped with the high-sensitivity sensor HSS7 (Mettler-Toledo Co., Switzerland). 2.3. SAXS. SAXS experiments were performed at the beamline 1W2A of the Beijing Synchrotron Radiation Facility (BSRF) (λ ) 1.54 Å). A standard silver behenate sample was used for the calibration of diffraction spacings. X-ray scattering intensity patterns were recorded during 60 s exposure of the samples to the synchrotron beam. A Linkam thermal stage (Linkam Scientific Instruments, the United Kingdom) was used for temperature control ((0.1 °C). The X-ray powder diffraction intensity data were analyzed using the program Fit2D. 2.4. FTIR Spectroscopy. FTIR spectra were recorded using a Nicolet 5700 Fourier transform infrared spectrometer with a DTGS detector in the range of 4000-900 cm-1 with a spectral resolution of 2 cm-1 and a zero filling factor of 2. The accuracy of the frequency is better than 0.1 cm-1. Samples were coated onto the inner surfaces of a pair of CaF2 windows, which were mounted on a Linkam heating-cooling stage for temperature control ((0.1 °C). For each spectrum, 16 scans were required, and the spectra were recorded every ∼30 s. The positions of various IR bands are determined by reading the center of gravity as suggested in the ref 16. 2.5. 2D IR Correlation Analysis. Sixteen spectra (from 20 to 35 °C) at an equal temperature interval (1 °C) over a selected wavenumber range were used for the 2D correlation analysis. Standard 2D correlation calculation was performed using Matlab 7.0 (Math Works Inc., Natick, MA), based on the algorithm developed by Noda.17 In the 2D correlation contour map, solid and dashed lines represent positive and negative correlation

Figure 2. DSC results of the SLPC-H2O and SLPC-D2O after storage at 0 °C for 10 days. The heating rate is 0.5 °C/min.

Figure 3. SAXS data of the SLPC-H2O system at the lamellar (15 °C) and micellar (50 °C) phases.

intensities, respectively. The data manipulations, i.e., the subtraction, truncation, and baseline correction of the IR spectra, were also done using Matlab 7.0. 3. Results and Discussion 3.1. DSC and SAXS Measurements. Figure 2 shows the DSC results of SLPC-H2O and SLPC-D2O after storage at around 0 °C for 10 days. SLPC in excess H2O displays a sharp endothermic peak with a peak temperature of 28.7 °C and a transition enthalpy of 23.2 kJ/mol, similar to the data reported in the literature.13 The endothermic transition corresponds to the reorganization of lysophospholipid molecules from a multilamellar to a micellar structure.13 As for the SLPC in D2O medium, the peak temperature is 29.0 °C, and the transition enthalpy is 24.6 kJ/mol. These slight increases in phase transition temperature and enthalpy indicate that during the lamellar to micellar transition the polar region that is in contact with the water medium has also been affected. Upon immediate cooling from the micellar solution to 15 at 0.5 °C/min, the lysophospholipids did not return to the lamellar structure, and a near straight line was observed in the DSC trace (data not shown), indicating that the formation of the lamellar structure is slow and the lamellar to micellar transition is irreversible under the present experimental conditions. However, we found that the lamellar structure of SLPC could be formed by prolonged annealing of the micellar solution at around 0 or 15 °C for hours or days. Shown in Figure 3 is the SAXS data of the SLPC-H2O system. The lamellar repeat spacing d of the SLPC dispersion at 15 °C was determined to be 6.3 nm (d ) 2π/q), consistent with the data reported by Hui and Huang.15 The calculated bilayer thickness without the water layer by these authors was

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Figure 4. Time-resolved FTIR absorbance spectra of SLPC-H2O over three selected wavenumber regions (A, B, and C) during the lamellar to micellar transition upon heating at 0.5 °C/min. The spectra are 1 °C apart. The wavenumber shifts of some selected bands as functions of temperature during the transition are shown in (D).

only 3.5-3.6 nm. The short bilayer thickness as compared with the lamellar repeat spacing value indicates that there is a large amount of interlamellar water to hydrate the SLPC headgroups. Considering that the molecular length as determined by the MM2 force field simulation was only 3.0 nm, the packing geometry of SLPC must be in a form of full interdigitation. Hui and Huang proposed that in the fully interdigitated model the long C(18) acyl chain extends across the entire hydrocarbon width of the bilayer.15 On the other hand, the SAXS result of the micellar state at 50 °C is shown as a broad diffuse band at q ) 0.92 nm-1. For the diluted 1-palmitoyllysophosphatidylcholine (PLPC) micelles (30 mM) in aqueous solution,7 a band at q ) 1.2 nm-1 charateristic of the intramicellar form factor7,18 was observed. While in our highly concentrated SLPC dispersions (25 wt %), the difference of the band position (and band shape) as compared with that of the diluted PLPC micelles is due to the interactions between micelles. 3.2. FTIR Measurements. The SLPC molecule can be divided into three parts: the headgroups N(CH3)3+ and PO4-, the interface region mainly containing the CdO group, and the tail acyl chains (CH2). These functional groups were selected as the IR probes for monitoring the changes of the head, interface, and tail regions of SLPC molecules, respectively, to see how these groups change during the lamellar to micellar and micellar to lamellar transitions. Shown in Figure 4A-C are the temperature-dependent FTIR spectra of the selected groups of SLPC molecules observed

during the lamellar to micellar transition at a heating rate of 0.5 °C/min. The corresponding wavenumber shifts of the selected bands as functions of temperature are presented in Figure 4D. The phase transition temperature identified in Figure 4D using the CdO and CH2 as the probing groups correlates well with the DSC result. The 3000-2800 cm-1 region (Figure 4A) contains mainly the CH2 asymmetric and symmetric stretching bands. The lamellar to micellar transition is characterized by shifts toward high frequencies and broadening of the bands. Specifically, the CH2 asymmetric and symmetric stretching bands at 2917.3 and 2849.8 cm-1 at the lamellar state (15 °C) shift to higher wavenumbers at 2923.9 and 2853.4 cm-1 at the micellar state (50 °C). These special features have been used frequently to follow the conformational order of the lipid methylene chains and the trans-gauche isomerization of the CH2 groups in lipid tail regions.10,19 The increase of wavenumber is partly due to the increase in the gauche conformers of methylene chains and partly due to the change in density or packing state of the chain region.20 The origin of the increase in bandwidth is the augmentation of the rotational motion of the methylene chains.21 The lamellar structure of SLPC is highly ordered, which is reflected in the viewpoint that the single acyl chains of lysophospholipids are fully interdigitated in the lamella. At higher temperatures in the micellar state, the CH2 stretching bands are broader, indicating high mobility and disordered packing. The 1800-1400 cm-1 region (Figure 4B) contains mainly three bands. The band at 1750-1700 cm-1 is from the CdO stretching, which is partially overlapped by the H2O absorbance

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Figure 5. Synchronous (A) and asynchronous (B) 2D correlation contour maps of the νsCH2 and νCdO bands. The solid and dashed lines represent positive and negative correlation intensities, respectively.

band. This band changes from 1726.9 cm-1 in the lamellar state to 1721.3 cm-1 in the micellar state, and the red-shift suggests that more water molecules are associated with the CdO groups at the micellar state. More detailed analyses will be discussed in the SLPC-D2O system. The band centered at ∼1490 cm-1 is ascribed to the asymmetric deformation vibration of the headgroup methyl groups attached to the N+ atom.22,23 It is an important band for monitoring the structural behavior of the hydrophilic part of the amphiphilic substance, which is known to be sensitive to the extent of disorder and the packing of the headgroups.24,25 As shown by the dotted line, the position of this band is constant during the lamellar to micellar transition, indicating no conformation and hydration changes occur for the N(CH3)3+ headgroup. The third important band in this region is the CH2 scissoring band centered at 1467.7 cm-1 in the lamellar state and at around 1467.4 cm-1 in the micellar state. Although there is no significant change in band position, the band sharpness reduces markedly upon phase transition. This band is very sensitive to the intermolecular forces and can serve as a key band for examining the state of packing of the methylene chains in various phases.19 In the lamellar state, the single sharp peak centered at 1467.7 cm-1 indicates that the methylene trans-zigzag planes are packed in the hexagonal state.19 While in the micellar state, the broad band at 1467.4 cm-1 suggests that the methylene chains are in the disordered fluid or fused state.26 Figure 4C contains IR bands of PO2- and N(CH3)3+ groups in the SLPC head region. These IR bands are sensitive to the change of the conformation and hydration of the headgroups. We can see that almost no band position shift or band contour change occurs in the asymmetric and symmetric stretching vibrations of PO2- at around 1222 and 1085 cm-1, respectively. It is worth noting that a number of small peaks appear in the region 1300-1200 cm-1 at low temperatures (marked with arrows), which are assigned to the CH2 wagging progressions of all-trans methylene chains in the lamellar state.19,27 These peaks reside on the shoulder of the PO2- asymmetric stretching band and modify the band appearance. However, the position of the νasPO2- band center is almost unchanged. The fact that the symmetric stretching vibration of PO2- at 1085 cm-1 also does not change in band position and band contour further confirms this conclusion. Moreover, for the asymmetric stretching vibration of the N+-CH3 group at around 972 cm-1,28 no evident band position or band contour changes are observed, consistent with the result of the δasN+-CH3 band at ∼1490 cm-1. The wavenumber changes of the selected IR bands of the head, interface, and tail groups are depicted in Figure 4D. It is

Figure 6. (A) Time-resolved FTIR absorbance spectra of SLPC-D2O in the region of 1800-1650 cm-1 upon heating at 0.5 °C/min, showing the change of the CdO stretching band. The spectra are 1 °C apart. (B) The peak fitting results of the CdO bands at 15 °C (lamellar) and 50 °C (micellar).

thus clear that, during the lamellar to micellar transition, band positions of the N(CH3)3+ and PO2- headgroups are almost fixed except the slight drift along with the temperature increase. At the same time, the band position of the interface region (νCdO) shows a downward shift of 5.6 cm-1. For the acyl tails, the change in the wavenumber is 3.6 cm-1 for νsCH2 and 6.6 cm-1 for νasCH2. The data show unambiguously the abrupt change in the conformations of the lipid interfacial region and tails. In a previous work, we proposed to use half-time, which is defined as the time spent when half of the event takes place, to present the occurrence of an event. By doing so, sequential order of two or more events can be evaluated.29 Following this concept, we compared the temperature of the half change in the wavenumber shift of the interfacial CdO groups (νCdO) and that of the acyl tails (νsCH2 or νasCH2) as indicated with the vertical lines in Figure 4D. It is found that the former is about 1 °C earlier than the latter. This indicates that the change in the interfacial groups is somewhat prior to that of the acyl tails. Furthermore, as 2D correlation spectroscopy can be used effectively to determine the sequential order of events,30–35 we also carried out 2D correlation analysis to study the change sequence of the lipid interfacial groups (νCdO) and the lipid acyl tails (νsCH2), and the results are shown in Figure 5. The synchronous and asynchronous 2D correlation contour maps of νCdO and νsCH2 are given in Figure 5A and 5B, respectively. In the synchronous 2D correlation map, there is a positive cross peak at (2849 cm-1, 1736 cm-1). This indicates that the changes of the absorbance of νCdO and νsCH2 during the phase transition have the same direction of change. The main cross peak in the asynchronous 2D correlation map at (2850 cm-1, 1725 cm-1) is negative. According to Noda’s rule,36,37 the absorbance of νsCH2 varies after that of νCdO. The 2D correlation analysis also implies that the lipid interfacial groups change earlier than the lipid acyl tails during the lamellar to micellar transition. More details on the change of the CdO groups during the lamellar to micellar transition are given by investigating the change of the CdO stretching band in the SLPC-D2O system (Figure 6). In the initial lamellar state at 15 °C, the CdO stretching band centers at 1730 cm-1, and it can be fitted by two subcomponents at 1737 and 1723 cm-1 with the assistance of literature work,38 with the peak area percentage of 23% and 77%, respectively. While in the micellar state at 50 °C, the CdO

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Figure 7. Time-resolved FTIR absorbance spectra of SLPC-H2O over three selected regions (A, B, and C) during the isothermal incubation at 15 °C. The spectra are 1 h apart. The time dependences of the normalized intensity changes of νasCH2 and νsCH2 during the incubation are shown in (D).

stretching band centers at 1725 cm-1, and it can also be fitted by two subcomponents at 1737 and 1723 cm-1, with the peak area percentage of 13 and 87%, respectively. The band at 1737 cm-1 is assigned to the stretching vibration of the CdO component without hydrogen bonding interactions, while the band at 1723 cm-1 is assigned to that hydrogen bonded with water molecules. The increase in the peak area of the band at 1723 cm-1 shows that the interfacial CdO groups increase in hydration degree during the lamellar to micellar transition. The reverse phase change process, the micellar to lamellar transition, was also studied using time-resolved FTIR spectroscopy during the isothermal process at 15 °C for 30 h. As shown in Figure 7 (A, B, and C), only the acyl tails (νasCH2/νsCH2 and δCH2) and the interfacial CdO groups (νCdO) change in the band positions. The acyl tails change from a disordered fluid or fused state to the ordered hexagonal packing state, while the interfacial CdO groups partially dehydrate. Meanwhile, other bands, such as the δasN+-CH3, νasPO2-, νsPO2-, and νasN+-CH3, do not change either in band position or in band contour, indicating that almost no conformation or hydration state changes in the lipid headgroups during the micellar to lamellar transition. The structural relaxation process is quite slow. As indicated in the change patterns of the normalized intensity changes of νsCH2 (2850.1 cm-1) and νasCH2 (2917.8 cm-1) (Figure 7D), the complete formation of the lamellar phase takes about 25 h. To sum up, these IR results indicate that during the lamellar to micellar and micellar to lamellar transitions the headgroups of SLPC molecules remain unchanged in their hydration state

and conformation; however, the interface groups change in the hydration state and tail regions change in the conformational order and packing state. To the best of our knowledge, for the first time the evidence was collected at the molecular/submolecular level that the different parts of an amphiphilic phospholipid molecule can change nonsynchronously during the phase transformation processes. 3.3. Nonsynchronicity Phenomenon. The stable phases of SLPC dispersions at low (15 °C) and high (50 °C) temperatures are lamellar and micellar phases, respectively. During the lamellar to micellar transition, the conformation and hydration degree of the headgroups do not change, while the interface increases in hydration and the tail portions change in conformation and packing state. Now the question is which one starts first, the lipid tail regions or the interfacial CdO groups? Here, we propose that the lamellar to micellar transition is triggered by the interfacial CdO groups: When approaching the onset temperature, the lipid acyl tails are so tightly packed in the interdigitated lamellar state that it is hard for them to rearrange first at the initial stage of the lamellar to micellar transition. On the other hand, the elevated temperature enhances the activity of the water molecules, and more water molecules may reach the lipid interface region. This will cause a loosening of the tail portion, which in turn induces more space in the interface region and allows more water molecules to associate with the interfacial CdO groups. The experimental results (Figure 4D and Figure 5) support the above transformation mechanism. Cooling the micellar phase from 50 to 15 °C produces merely a straight line in the DSC thermogram, indicating that no phase

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Figure 8. Schematic models showing the lamellar to micellar transition of lysoPCs (A) and lamellar (LR) to inverse micellar (HII) transition of PEs (B) in excess water.

transition occurs during the cooling process. In other words, the phase state at the end point (15 °C) of the cooling process is still the micellar state. This micellar state at 15 °C is supercooled and thus metastable and will eventually transform into the lamellar state upon incubation at this temperature or other lower temperatures for hours or days. The conversion of lysophospholipid molecules from a micellar structure to a two-dimensional lamella involves the ordering and interdigitation of the long acyl chains, as well as the partial dehydration in the interfacial CdO region. The slow kinetics of the lamellar formation process from the micellar state can be discussed in terms of the joining of a nucleation and growth process.14 The nucleation process is favored at relatively lower temperatures, such as 0 or 15 °C in the SLPC-H2O system, which is well below the lamellar to micellar transition temperature (∼29 °C). We observed slight downward shifts of the νasCH2 (from 2924.0 to 2921.9 cm-1) and νsCH2 (from 2853.2 to 2852.2 cm-1) during cooling from 50 to 15 °C (although a straight line was observed in the DSC cooling thermogram), indicating that some new trans conformers are formed. This will facilitate the occurrence of local structural order and thus the occurrence of the nuclei of the lamellar state. Meanwhile, almost no band position or band contour changes were observed in the interfacial CdO groups or the polar headgroups during cooling. Thus, this kind of nuclei is not the ordering of the whole SLPC molecules but only the ordering of some united SLPC tail portions. During the isothermal incubation period, the presence of the nuclei of the lamellar state provides the SLPC molecules more chance to change their conformational and packing mode to rearrange into the ordered lamellar structure. The change in the lipid packing mode will induce the change of the interfacial CdO group. However, this kind of growth process of the lamellar state still involves only the tail and interface regions, not the whole lipid molecules. Thus, the dehydration of the interfacial CdO groups is induced by the ordering of the tail region, and the change in lipid tails should be the key factor that controls the micellar to lamellar transition. There have been similar reports that the interfacial glycerol backbone conformation can be principally governed by the vicinally arranged acyl chains in phospholipid aggregates under certain conditions.39,40 To put these in the context of the nonsynchronicity phenomenon, we emphasize that the lipid tails are the factor controlling the micellar to lamellar transition. We now compare the hydration degree changes in two kinds of lamellar to nonlamellar transitions. Shown in Figure 8 are the schematic models showing the lamellar to micellar transition of lysoPCs and lamellar (LR) to inverse micellar (HII) transition of PEs in excess water. For a lipid possessing positive spontaneous curvature, such as the SLPC molecule, it can form

J. Phys. Chem. B, Vol. 114, No. 6, 2010 2163 micelles (spherical or ellipsoidal shape) at proper concentrations and temperatures. Under our conditions, heating the initial SLPC lamellar structure results in the formation of the micellar structure. In the micellar structure, the headgroups are in good contact with water and are well hydrated as in the lamellar state. No hydration change occurs during the lamellar to micellar transition. As for the other lamellar to nonlamellar phase transition (LR-HII transition), there have been reports concerning the changes of hydration properties of the nonbilayer-forming PE membranes. For the monounsaturated stearoyloleylphosphatidylethanolamine (SOPE), it was found that the number of water molecules residing at the lipid/water interface in the HII phase reduces as compared with the preceding LR phase.41 In another study, Channareddy et al. directly determined the hydration change during the LR-HII transition process of dioleoylphosphatidylethanolamine (DOPE).42 They found that in the LR phase of DOPE (at 2 °C) 7.2 water molecules per phospholipid were bound, while in the HII phase (at 16 and 30 °C) 5.4 and 5.6 water molecules per phospholipid were bound. A dehydration of approximately 2 water molecules per PE occurs upon the LR-HII transition. The overall hydration behavior of the hexagonal phase is governed mainly by the spontaneous curvature of the monolayers, thus a temperature increase reduces the number of water molecules/lipid.41,43 As PE molecules tend to form strong hydrogen bonding networks between the positively charged amine and the negatively charged phosphate groups at proper distances,44,45 the increase in the curvature of the HII phase will further increase this tendency. In conclusion, the geometrical constraints in the hexagonal phase and the strong intermolecular interactions between the PE headgroups contribute to the dehydration of the lipid headgroups during the LR-HII transition. The different behavior of the hydration changes in the lipid headgroups during the two kinds of lamellar to nonlamellar phase transitions emphasizes the important roles of the membrane geometry. In a previous study, we found that during the liquid crystalline to coagel phase transition of a simple structured cationic lipid, dioctadecyldimethylammonium bromide (DODAB), the lipid tails change prior to the headgroups.23 In that work, we attributed the origin of the nonsynchronous change of the lipid head and tail to the lack of special intermolecular attractive forces (hydrogen bonding and electrostatic interactions) between the neighboring polar headgroups of DODAB molecules. While in this SLPC system, we suggest that the lack of change in the intermolecular attractive electrostatic interactions between the neighboring polar headgroups of SLPC molecules is the origin of the occurrence of the nonsynchronicity phenomenon during the lamellar to micellar and micellar to lamellar transitions. The previous DODAB system is a typical nonsynchronicity phenomenon found in a lamellar to lamellar phase transition process. Our present work demonstrates that the nonsynchronicity phenomenon can also occur in a lamellar to nonlamellar transition in the SLPC molecules, which is, to the best of our knowledge, the first work to observe such a nonsynchronicity phenomenon in a phospholipid system. Furthermore, although there are numerous studies on the properties and applications of micelles, seldom efforts have been made on the tracking of the dynamic micelle formation process especially where the different parts of the amphiphilic molecules are concerned. Our work provides molecular evidence that the lamellar to micellar transition of SLPC molecules is associated with no change in the solvation of the hydrophilic heads and their conformation, which is related to the permeability,

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flexibility, and fusion ability of the head region in the molecule. In fact, various nonbilayer micellar structures have been proposed as an obligatory intermediate during membranemembrane fusion.14,46,47 The kinetic study of the micellar to lamellar transition may therefore provide useful information which will aid in elucidating the mechanism of membrane fusion.14 4. Conclusions We have studied in detail the thermotropic phase behavior of the SLPC aqueous dispersions by using DSC, SAXS, FTIR, and 2D correlation analysis. The stable phases at low and high temperatures are lamellar (interdigitated gel phase) and micellar phases, respectively. Upon heating, the lamellar phase converts to the micellar phase, while on cooling, the micellar state does not transform into the lamellar phase instantaneously. The formation of the lamellar state requires incubation at around 0 °C for hours or days. At 15 °C, we observed that the lamellar state can also form upon incubation for around 25 h. Transformation from lamellar to micellar upon heating involves both a disordering rearrangement in the acyl tails and a hydrating process in the interface region, but no evident conformational or hydration changes of the lipid headgroups are observed. That is, the head, interface, and tail regions of SLPC molecules do not change synchronously during the process. We further suggest that the lamellar to micellar transition is initiated by the interfacial CdO groups. The molecular mechanism of the slow formation kinetics of the lamellar state from the micellar state was also discussed in the context of the nonsynchronicity phenomenon. It is emphasized that the lipid tail region is the bottleneck controlling the micellar to lamellar transition. The retention of the intermolecular attractive forces between the neighboring polar headgroups of the amphiphiles may be the origin of the nonsynchronicity phenomenon. Such a nonsynchronicity phenomenon in the self-assembled aggregates composed of the medium-sized lipid molecules reflects the regional (head, interface, and tail) imbalance in molecular interactions. Acknowledgment. This work was supported by grants from the Natural Science Foundation of China (NSFC: 20633080, 20973100) and a “973” National Key Basic Research Program of China (Grant No. 2006CB806203). The SAXS data were collected at the beamline 1W2A of the Beijing Synchrotron Radiation Facility (BSRF) with the assistance of Dr. ZhongHua Wu and Zhi-Hong Li. References and Notes (1) La Rosa, C.; Grasso, D.; Checchetti, A.; Golemme, A.; Chidichimo, G.; Westerman, P. W. Biophys. Chem. 1998, 70, 11–20. (2) Bhamidipati, S. P.; Hamilton, J. A. Biochemistry 1995, 34, 5666– 5677. (3) Checchetti, A.; Golemme, A.; Chidichimo, G.; La Rosa, C.; Grasso, D.; Westerman, P. W. Chem. Phys. Lipids 1996, 82, 147–162. (4) Chernomordik, L.; Chanturiya, A.; Green, J.; Zimmerberg, J. Biophys. J. 1995, 69, 922–929. (5) Kumar, V. V.; Malewicz, B.; Baumann, W. J. Biophys. J. 1989, 55, 789–792. (6) Ramsammy, L. S.; Brockerhoff, H. J. Biol. Chem. 1982, 257, 3570– 3574.

Wu et al. (7) Barbosa, L. R. S.; Caetano, W.; Itri, R.; Homem-de-Mello, P.; Santiago, P. S.; Tabak, M. J. Phys. Chem. B 2006, 110, 13086–13093. (8) Tenchov, B.; Koynova, R.; Rapp, G. Biophys. J. 2001, 80, 1873– 1890. (9) Seddon, J. M.; Templer, R. H. Handbook of Biological Physics; Lipowsky, R., Sackmann, E., Eds.; Elsevier Science B.V.: Amsterdam, 1995; Vol. 1, pp 97-160. (10) Wu, F. G.; Chen, L.; Yu, Z. W. J. Phys. Chem. B 2009, 113, 869– 872. (11) Qin, S. S.; Yu, Z. W.; Yu, Y. X. J. Phys. Chem. B 2009, 113, 8114–8123. (12) Gao, W. Y.; Chen, L.; Wu, R. G.; Yu, Z. W.; Quinn, P. J. J. Phys. Chem. B 2008, 112, 8375–8382. (13) Wu, W. G.; Huang, C. H.; Conley, T. G.; Martin, R. B.; Levin, I. W. Biochemistry 1982, 21, 5957–5961. (14) Wu, W. G.; Huang, C. H. Biochemistry 1983, 22, 5068–5073. (15) Hui, S. W.; Huang, C. H. Biochemistry 1986, 25, 1330–1335. (16) Binder, H.; Pohle, W. J. Phys. Chem. B 2000, 104, 12039–12048. (17) Noda, I. Appl. Spectrosc. 1993, 47, 1329–1336. (18) Itri, R.; Amaral, L. Q. Phys. ReV. E 1993, 47, 2551–2557. (19) Lewis, R. N. A. H.; McElhaney, R. N. Methods in Molecular Biology; Dopico, A. M., Ed.; Humana Press: Totowa, NJ, 2007; Vol. 400, pp 207-226. (20) Umemura, J.; Cameron, D. G.; Mantsch, H. H. Biochim. Biophys. Acta 1980, 602, 32–44. (21) Kawai, T.; Umemura, J.; Takenaka, T.; Kodama, M.; Seki, S. J. Colloid Interface Sci. 1985, 103, 56–61. (22) Kawai, T.; Umemura, J.; Takenaka, T.; Kodama, M.; Ogawa, Y.; Seki, S. Langmuir 1986, 2, 739–743. (23) Wu, F. G.; Wang, N. N.; Yu, Z. W. Langmuir 2009, 25, 13394– 13401. (24) Wang, W.; Li, L. M.; Xi, S. Q. J. Colloid Interface Sci. 1993, 155, 369–373. (25) Wong, T. C.; Wong, N. B.; Tanner, P. A. J. Colloid Interface Sci. 1997, 186, 325–331. (26) Snyder, R. G. J. Chem. Phys. 1967, 47, 1316–1360. (27) Senak, L.; Moore, D.; Mendelsohn, R. J. Phys. Chem. 1992, 96, 2749–2754. (28) Chen, C. F.; Tripp, C. P. Biochim. Biophys. Acta 2008, 1778, 2266– 2272. (29) Jia, Q.; Wang, N. N.; Yu, Z. W. Appl. Spectrosc. 2009, 63, 344– 353. (30) Noda, I. J. Am. Chem. Soc. 1989, 111, 8116–8118. (31) Zhang, J. M.; Tsuji, H.; Noda, I.; Ozaki, Y. J. Phys. Chem. B 2004, 108, 11514–11520. (32) Zhang, J. M.; Duan, Y. X.; Shen, D. Y.; Yan, S. K.; Noda, I.; Ozaki, Y. Macromolecules 2004, 37, 3292–3298. (33) Zhang, J. M.; Sato, H.; Noda, I.; Ozaki, Y. Macromolecules 2005, 38, 4274–4281. (34) Yu, Z. W.; Chen, L.; Sun, S. Q.; Noda, I. J. Phys. Chem. A 2002, 106, 6683–6687. (35) Wang, N. N.; Jia, Q.; Li, Q. Z.; Yu, Z. W. J. Mol. Struct. 2008, 883, 55–60. (36) Noda, I. J. Mol. Struct. 2006, 799, 41–47. (37) Noda, I.; Ozaki, Y. Two-dimensional Correlation SpectroscopyApplications in Vibrational and Optical Spectroscopy; Wiley: Chichester, 2004. (38) Lewis, R. N. A. H.; McElhaney, R. N. Chem. Phys. Lipids 1998, 96, 9–21. (39) Hauser, H.; Pascher, I.; Sundell, S. Biochemistry 1988, 27, 9166– 9174. (40) Arora, A.; Gupta, C. M. Biochim. Biophys. Acta 1997, 1324, 47– 60. (41) Rappolt, M.; Hodzic, A.; Sartori, B.; Ollivon, M.; Laggner, P. Chem. Phys. Lipids 2008, 154, 46–55. (42) Channareddy, S.; Jose, S. S.; Janes, N. J. Am. Chem. Soc. 1997, 119, 2345–2347. (43) Harper, P. E.; Mannock, D. A.; Lewis, R. N.; McElhaney, R. N.; Gruner, S. M. Biophys. J. 2001, 81, 2693–2706. (44) Suits, F.; Pitman, M. C.; Feller, S. E. J. Chem. Phys. 2005, 122, 244714-1-244714-9. (45) Lewis, R. N. A. H.; McElhaney, R. N. Biophys. J. 1993, 64, 1081– 1096. (46) Lucy, J. A. Nature 1970, 277, 814–817. (47) Cullis, P. R.; Hope, M. M. Nature 1978, 271, 672–674.

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