Morphology-Induced Defects Enhance Lipid Transfer Rates

Article pubs.acs.org/Langmuir

Morphology-Induced Defects Enhance Lipid Transfer Rates Yan Xia,† Kamil Charubin,† Drew Marquardt,‡,§ Frederick A. Heberle,∥,⊥ John Katsaras,∥,⊥ Jianhui Tian,# Xiaolin Cheng,# Ying Liu,† and Mu-Ping Nieh*,†,∇,○ †

Department of Chemical and Biomolecular Engineering, ∇Polymer Program, Institute of Materials Science, and ○Department of Biomedical Engineering, University of Connecticut, Storrs, Connecticut 06269, United States ‡ Institute of Molecular Biosciences, Biophysics Division, NAWI Graz, University of Graz, Graz 8010, Austria § Department of Physics, Brock University, St. Catharines, Ontario L2S 3A1, Canada ∥ Biology and Soft Matter Division, Neutron Sciences Directorate, ⊥Joint Institute for Neutron Sciences, and #Center for Molecular Biophysics, Oak Ridge National Laboratory, Oak Ridge, Tennessee 37831, United States S Supporting Information *

ABSTRACT: Molecular transfer between nanoparticles has been considered to have important implications regarding nanoparticle stability. Recently, the interparticle spontaneous lipid transfer rate constant for discoidal bicelles was found to be very different from spherical, unilamellar vesicles (ULVs). Here, we investigate the mechanism responsible for this discrepancy. Analysis of the data indicates that lipid transfer is entropically favorable, but enthalpically unfavorable with an activation energy that is independent of bicelle size and longto short-chain lipid molar ratio. Moreover, molecular dynamics simulations reveal a lower lipid dissociation energy cost in the vicinity of interfaces (“defects”) induced by the segregation of the long- and short-chain lipids in bicelles; these defects are not present in ULVs. Taken together, these results suggest that the enhanced lipid transfer observed in bicelles arises from interfacial defects as a result of the hydrophobic mismatch between the long- and short-chain lipid species. Finally, the observed lipid transfer rate is found to be independent of nanoparticle stability.

■

ied,14−18 their kinetic properties (i.e., stability19 and lipid transfer rates20) have not attracted much attention. Molecular transfer kinetics have been shown to affect the formation of polymeric, surfactant and lipid-based micelles, and their stabilities.21−25 We recently reported a 150-fold enhancement in the interparticle lipid transfer rate constant, kinter, of dimyristoyl-PC (DMPC) bicelles (0.156 ± 0.011 h−1) compared to that of DMPC ULVs [(1.01 ± 0.06) × 10−3 h−1]. The dramatic difference was not expected to stem from lipid flip-flop; otherwise, the initial transfer rate between ULVs would have been similar to that of bicelles.26 Although an interesting result, the molecular origin of this difference in lipid transfer rate was not developed. To understand the mechanism responsible for this difference, we have performed a systematic study of lipid transfer between dipalmitoyl-PC (DPPC) and dihexanoyl-PC (DHPC) bicelles. 5 mol % of the negatively charged dipalmitoylphophotidylglycerol (DPPG) was added to these bicelles to minimize the possibility of them fusing with each other. Time-resolved smallangle neutron scattering (SANS) and differential scanning calorimetry (DSC) requiring only isotopic substitution were used to measure lipid transfer rate constants. Both SANS and

INTRODUCTION Biological membranes allow for compartmentalized biochemical processes to take place. The structural dynamics of biomembranes and the transfer kinetics of their molecular constituents, primarily lipids, sterols, and proteins, are among the most important physical parameters affecting these biochemical processes. For example, translocation and flipflop of phospholipids in biological membranes are known to play a significant role in endocytotic and exocytotic processes, as well as the determination of molecular pathways.1,2 Malfunctions in lipid transfer can lead to cardiovascular3 and autoimmune diseases,4 atherosclerosis,5 obesity,6 and diabetes,7 to name just a few. Because physical studies are often stymied by the chemical complexity of biological membranes, tractable model systems are used to investigate the physicochemical properties and dynamic behavior of lipid bilayers. Of special interest are unilamellar vesicles (ULVs) and discoidal bicelles, owing to their ease of preparation, well-characterized morphologies, and well-defined size distributions.8−12 It has been reported that bicelles composed of a mixture of long- and short-chain phosphatidylcholines (PC) have a uniform diameter and thickness, and spontaneously form under certain environmental conditions.13 The long-chain lipids constitute the planar bilayer disk, while the short-chain lipids are sequestered at the disk's rim. Although the structural properties of bicellar mixtures have been extensively stud© XXXX American Chemical Society

Received: June 3, 2016 Revised: August 23, 2016

A

DOI: 10.1021/acs.langmuir.6b02099 Langmuir XXXX, XXX, XXX−XXX

Article

Langmuir

respectively] between 0.005 and 0.25 Å−1 was selected through a combination of a 4 m sample-to-detector distance (SDD) and the 30 Hz “frame-skipping” mode, with a minimum wavelength of 2.5 Å, giving rise to a second wavelength band starting at 9.4 Å. A 25 mm source and 10 mm sample apertures were used for all measurements. The resulting two-dimensional (2D) scattering profiles were reduced to one-dimensional (1D) SANS profiles using the Mantid program (http://www.mantidproject.org/) developed by ORNL. The data were then put on an absolute scale. During the reduction process, data were corrected for detector pixel sensitivity, dark current, and sample transmission. At NCNR, the SANS data were collected at three different SDDs (13, 4, and 1 m) using 8.4 Å wavelength neutrons and a neutron “focusing” lens for the longest SDD position (i.e., 13 m) and 6 Å wavelength neutrons for the two other SDDs. This yielded a q range from 0.003 to 0.4 Å−1. The 2D raw data were corrected for detector sensitivity, background, sample transmission, empty cell scattering and transmission. The corrected data were then circularly averaged, yielding the customary 1D profiles, which were then put on an absolute intensity scale using the measured incident beam flux. The h- and d-DPPC bicelles (1 wt %) with Q = 3 and R = 0.05 were prepared in a CM H2O/D2O mixture (see Bicelle Preparation). The lipid transfer rate at different incubation temperatures (i.e., 20, 25, and 30 °C) was monitored through a series of time-resolved scattering data. The effect of 5 mol % DSPE-PEG2000 on the lipid transfer rate of bicelles was studied using the same approach as neat bicelles. It should be noted that the structure factor S(q) of the solution composed of h- and d-bicelles in CM solvents is negligible, thus allowing us to use a disk form factor to fit the SANS data, even for a highly interacting (highly charged) system.1,33,34 As lipid transfer proceeds, the contrast factor, (Δρ)2 (Δρ = |ρs − ρBic|, where ρs and ρBic are the SLDs of the solvent and bicelle, respectively) decreases. This can be related to the spontaneous lipid transfer rate of the system. The morphologies of bicelles with different Q values were investigated using SANS. Similar to the aforementioned samples, hand d-DPPC bicelles (0.5 wt %) with Q = 2.5, 3, and 3.5 were dispersed in CM solvent, where S(q) is approximated to be 1. To compare to the DSC results, an equimolar mixture of h- and d-bicelles was studied at 20 °C. DSC Measurements. DSC experiments were conducted using a NanoDSC (TA Instruments, New Castle, DE). Different Q (i.e., 2.5, 3, and 3.5) h- and d-bicelles were diluted from 10 to 0.5 wt % using deionized (DI) water. At t = 0, h- and d-bicelles were mixed using equal molar ratios of each. The h-bicelle/d-bicelle mixtures were then stored at 20, 25, or 30 °C for time-resolved studies. A sample aliquot (∼0.5 mL) was taken at different times for DSC measurements, where the bicellar solution and DI water were loaded in the sample and reference cells, respectively. The pressure of all experiments was kept at 3 atm. Prior to each DSC measurement, samples were equilibrated at 5 °C for 10 min. DSC endotherms were collected from 5 to 55 °C at a rate of 1 °C/min. The raw DSC profiles were then corrected for solvent background, where both sample and reference cells were filled with DI water. Baselines were adjusted using the TA Instruments software. Simulation. Umbrella sampling MD simulations were conducted to compare the difference in the potential of mean force (PMF) of pulling one DPPC lipid out from a single component planar bilayer and from a bicelle. The bilayer system consisted of a total of 392 DPPC lipids (i.e., 196 lipids in each leaflet). One lipid molecule was pulled out from one leaflet along the membrane normal (z) direction. A total of 22 sampling windows were used, with a window width of 0.1 nm and a force constant of 1000 kJ mol−1 nm−2), thus guaranteeing sufficient overlap between neighboring windows. For the bicelle system, an “inverse” bicelle model (Figure 1) was built, made up of a total of 96 DHPC lipids and 312 DPPC lipids, with an equal number of lipids in each leaflet. DHPC lipids were mainly located at the boundary region, surrounding a water hole. The PMF of one DPPC lipid located near the DHPC-rich region (referred as to “interfacial domain” later) was simulated to be pulled out from the leaflet. A total of 42 sampling windows were used with a window width of 0.1 nm. The same force constant used for the bilayer system was used for the

DSC use the properties of deuterated lipids (e.g., different scattering strength and melting transition temperature, TM), which are distinct from their protiated counterparts. For example, by monitoring the time-dependent decay of the neutron scattering length density (NSLD) or TM difference (ΔTM) in a mixture of protiated and deuterated bicelles, the lipid transfer rate can be obtained.25,27−31 Through complementary molecular dynamics (MD) simulations, we find that the enhanced lipid transfer rate constant is primarily due to the reduced dissociation energy (from bilayer to aqueous phase) of lipids at the interface between long- and short-chain lipid patches, where significant bilayer thickness mismatch is present. In fact, it has been reported that the bilayer thickness mismatch could control lipid domain size in ULVs,9,32 which has drawn considerable attention because of its potential relevance to lipid rafts in biological membranes. The insights gained from the current study advance our understanding of the interplay between the structure and dynamics of biomimetic lipid mixtures.

■

MATERIALS AND METHODS

Materials. 1,2-Dipalmitoyl-sn-glycero-3-phosphocholine (DPPC), 1,2-dipalmitoyl-d62-sn-glycero-3-phosphocholine (d-DPPC), 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC), 1,2-dimyristoyl-d54sn-glycero-3-phosphocholine (d-DMPC), 1,2-dipalmitoyl-sn-glycero-3phospho-(1′-rac-glycerol) (sodium salt) (DPPG), 1,2-dihexanoyl-snglycero-3-phosphocholine (DHPC), and 1,2-distearoyl-sn-glycero-3phosphoethanolamine-N-[methoxy(polyethylene glycol)-2000] (ammonium salt) (DSPE-PEG2000) were purchased from Avanti Polar Lipids (Alabaster, AL) and used without further purification. Deuterium oxide (D2O, 99.9% purity) was purchased from Cambridge Isotope Laboratories, Inc. (Andover, MA). Bicelle Preparation. Lipid mixtures of DPPC/DPPG/DHPC and d-DPPC/DPPG/DHPC were prepared at long- to short-chain lipid molar ratios, Q {≡([DPPC] + [DPPG])/[DHPC]}, of 2.5, 3, and 3.5, with a constant long-chain charged lipid ratio, R {≡[DPPG]/([DPPC] + [DPPG])}, of 0.05. The DPPC/DPPG/DHPC and d-DPPC/ DPPG/DHPC are denoted as h-DPPC bicelles and d-DPPC bicelles, respectively. In the case of lipid mixtures containing DSPE-PEG2000, four samples with 5.0 mol % DSPE-PEG2000, [DSPE-PEG2000]/ [total lipids] = 0.05, were prepared (i.e., h-DPPC, d-DPPC, h-DMPC, and d-DMPC PEGylated bicelles at Q = 3 and R = 0.05). The same protocol was used for all samples, as described below. Dry lipid powders were co-dissolved in chloroform. After removal of the bulk chloroform using a nitrogen stream, samples were placed under vacuum overnight. The dried lipid films were rehydrated to 10 wt % with either contrast matching (CM) solvent for SANS experiments or deionized water for DSC experiments, followed by temperature cycling between 4 and 60 °C, and vortexing. The CM solvent was made up of a mixture of H2O and D2O in amounts that yielded the same NSLD as that of the mixture of h- and d-bicelles, detailed compositions of which are listed in Table S1 of the Supporting Information. The final bicelle solutions were transparent and liquid-like at low T ( 0),44 where, in comparison to DMPC ULVs, a 20-fold increase in kinter for DMPC nanodiscs was observed at 27 °C. The enhanced lipid transfer for nanodiscs was attributed to a tighter packing of lipids at the edge of the nanodiscs caused by the rim tension induced by the lipoprotein “belt”.44 However, this rationale does not apply to bicelles whose lateral tension is expected to be minimized through bicelle fusion19,45 or ULV formation,46,47 rather than through enhanced lipid transfer. Here, we propose a different mechanism to account for the large increase in kinter, namely, the presence in bicelles of an interface region separating DPPC-rich from DHPC-rich domains. The first line of evidence for this proposed mechanism is the invariant activation energy for lipid transfer in bicelles at different Q values. We retrieved kinter using time-resolved DSC for bicelles at different Q (see time-resolved DSC data in Figure S1 and Tables S5−S13 of the Supporting Information). The difference in TM values (ΔTM) as a function of time exhibits an exponential decay from which kinter is determined, as shown in Figure 5 and Table 1, where kinter increases with decreasing Q (i.e., increased DHPC molar ratio).31,48 As a result of the difficulty in accurately differentiating the phase transition peaks associated with h- and d-rich bicelles at the late stages of lipid transfer, only a limited number of data points were obtained at high temperature (i.e., 30 °C). Ea for the transfer process obtained from the Arrhenius analysis (Figure 6) is independent of Q (Ea = 173.2 ± 4.5, 179 ± 13, and 168 ± 20 kJ/mol for Q = 2.5, 3, and 3.5 bicelles, respectively), revealing that the energy barrier for DPPC dissociating from bicelles into the water phase is independent of DHPC concentration. The increase in kinter observed with decreasing Q is most likely attributed to an increasing interface between DPPC-rich and DHPC-rich domains rather than a fundamental change in the energy landscape. The proposed mechanism of lipid transfer is further validated by analyzing the SANS data using a disk model. Analysis shows higher Q samples form larger diameter bicelles (radii of 73 ± 8, 82 ± 7, and 95 ± 6 Å for Q = 2.5, 3, and 3.5 bicelles, respectively; Figure S2 and Table S14 of the Supporting Information). In bicelles, DPPC and DHPC molecules are predominantly located at the disk’s plane and rim, respectively, as a result of differences in packing parameters but are not excluded from each others regions (as shown in Figure 7). As a result, the interface between DPPC-rich and DHPC-rich domains is mostly localized at the disk rim. The fraction of interfacial DPPC (∼4πR bicelle ) to total bilayer DPPC (∼2πRbicelle2) is proportional to (Rbicelle)−1. Lower Q (smaller) bicelles, therefore, have a greater total interfacial area, leading to enhanced lipid transfer. The reduced enthalpy of the DPPC gelto-Lα transition at lower Q further supports this notion (Figure

Figure 3. NSLD contrast decays of equimolar deuterated and protiated DPPC/DHPC/DPPG bicelles in contrast matched water. The data were fitted with exponential decay. Values of R2 for the single exponential fits of 20, 25, and 30 °C data are 0.9827, 0.9871 and 0.9717, respectively.

monomeric lipid transfer takes place under these experimental conditions. The best fit values are listed in Tables S2−S4 of the Supporting Information.26 An activation energy, Ea, of 119 ± 9.8 kJ mol−1 is obtained through the Arrhenius analysis, as shown in Figure 4. The value of kinter at 10 °C was also included in Figure 4 from previously obtained data.26 Next, we used transition state theory (TST) to obtain the thermodynamic parameters for lipid transfer.39 Specifically, the temperature dependence of the transfer rate constant is given by the Eyring−Polanyi equation

Figure 4. Arrhenius plot of kinter for DPPC/DHPC/DPPG (Q = 3) bicelles obtained from SANS measurements. DSC and SANS data are in good agreement with each other (inset). The R2 of the linear fit is 0.9845. D

DOI: 10.1021/acs.langmuir.6b02099 Langmuir XXXX, XXX, XXX−XXX

Article

Langmuir

Figure 5. DSC data of ln ΔTM as a function of time in different Q DPPC/DHPC/DPPG bicelles at T = 20, 25, and 30 °C. The error bars were derived from the Gaussian fits of DSC endotherms and the experimental errors. The values of R2 for the linear fits of Q = 2.5 at 20, 25, and 30 °C, Q = 3 at 20, 25, and 30 °C, and Q = 3.5 at 20, 25, and 30 °C are 0.9896, 0.9489, 0.9974, 0.9535, 0.9320, 0.9589, 0.9754, 0.9624, and 0.9142, respectively.

Table 1. Lipid Transfer Rate Constants (h−1) of DPPC/DHPC/DPPG Bicellesa 20 °C (DSC) Q = 2.5 Q = 3.0 Q = 3.5

0.0012 ± 7 × 10−5 0.0010 ± 1.1 × 10−4 0.00053 ± 5 × 10−5

20 °C (SANS)

25 °C (DSC)

0.0013 ± 5 × 10−5

0.0041 ± 6.8 × 10−4 0.0031 ± 4.8 × 10−4 0.0021 ± 2.1 × 10−4

25 °C (SANS)

30 °C (DSC)

30 °C (SANS)

0.0029 ± 5 × 10−5

0.012 ± 4.4 × 10−4 0.012 ± 0.00245 0.0051 ± 7.8 × 10−4

0.0084 ± 2.6 × 10−4

The error bars for the rate constants are obtained from the single exponential decays in Figure 3 for TR-SANS and linear fits of ln(ΔTM) for TRDSC in Figure 5.

a

DHPC-rich rim are likely to be in the liquid disordered phase and, thereby, do not undergo a phase transition. As a result, the exchange of DPPC between discoidal rims would not contribute to the decay of ΔTM. Further evidence for this is the 40−50% increase in kinter in bicelles doped with 5 mol % of distearoyl-phosphatidylethanolamine-PEG2000 (DSPEPEG2000) [(4.13 ± 0.15) × 10−4 and 0.233 ± 0.003 h−1 for DSPE-PEG2000-associated DPPC and DMPC bicelles at 10 °C, as derived from Figures S4 and S5 and Tables S17 and S18 of the Supporting Information) compared to the reported values of neat bicelles (2.81 ± 0.02 × 10−4 and 0.156 ± 0.011 h−1 for DPPC and DMPC bicelles at 10 °C, respectively),26 even though DSPE-PEG2000-doped bicelles appear more stable as a result of steric effects.49 This increase in kinter is most likely underpinned by the same mechanism found in neat bicelles, because the size of DSPE-PEG2000-containing bicelles is smaller than that of neat bicelles (Table S14 of the Supporting Information). MD simulations were performed to assess the energies of DPPC lipid dissociation from a planar DPPC bilayer or a DPPC−DHPC interface (Figure 1).35,36 A free energy penalty of 83.7 ± 0.42 kJ/mol was incurred in the case of planar DPPC bilayers, while the energy required for pulling out DPPC from the DPPC−DHPC interface was only 65.7 ± 0.42 kJ/mol (Figure 8). Both processes are, however, energetically unfavorable, consistent with the fact that the dissociation of lipids forming stable bilayers is the bottleneck for lipid transfer. The difference (18.0 kJ/mol) in the dissociation energy from MD simulations gives us a molecular understanding of the experimentally determined kinter, which is enhanced in the case of bicelles compared to vesicles. Moreover, in our simulations, we observed that pulling one lipid molecule from the interface appears to “drag” surrounding lipids out of the bilayer, inducing a local membrane curvature. Furthermore, the simulation results imply that the intrinsic lipid transfer rate constant at interfaces does not depend upon the DHPC concentration, consistent with the Arrhenius analysis.26

Figure 6. Arrhenius plots of different Q DPPC/DHPC/DPPG bicelles obtained from DSC data. R2 values for Q = 2.5, 3 and 3.5 are 0.999, 0.995 and 0.986, respectively.

Figure 7. Schematic of DPPC lipids (gray headgroups) at the interface between the gel and Lα DPPC in the vicinity of the DHPC (green headgroups), represented by blue (rim) and red (plane) rectangles, respectively.

S3 and Table S16 of the Supporting Information), suggesting the presence of fewer liquid ordered DPPC molecules, i.e., more liquid disordered DPPC molecules are found at the interface. It is noteworthy that DPPC molecules located at the E

DOI: 10.1021/acs.langmuir.6b02099 Langmuir XXXX, XXX, XXX−XXX

Article

Langmuir

DPPC lipids (presumably in the vicinity of the defects), instead of those in the gel phase.

■

CONCLUSION In conclusion, TR-SANS, TR-DSC and MD simulations suggest that the interface between hydrophobic mismatched DPPC and DHPC molecules accounts for the faster lipid transfer observed in bicelles compared to vesicles (as high as 2 orders of magnitude). This observation has enabled a molecular understanding as to how defects can substantially alter the physical characteristics of a system. The mechanism underpinning this phenomenon can also be applied to other biologically relevant and polymeric systems.

■

Figure 8. PMFs as a result of pulling one DPPC lipid out of a bilayer (black symbols) in the vicinity of the DHPC domain (red symbols).

ASSOCIATED CONTENT

S Supporting Information *

Further comparison of the DSC endotherms of DPPC/ DPPG ULVs and DPPC/DPPG/DHPC bicelles (Q = 3) (shown in Figure 9) confirms the notion of packing defects in

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.langmuir.6b02099. Contrast-matched solvents (S1), TR-SANS (S2), TRDSC (S3), determination of the morphology of different Q bicelles using SANS (S4), DSC endotherms of h-, d-, and h-/d-bicelles (S5), and lipid exchange between PEGylated bicelles (S6) (PDF)

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS Mu-Ping Nieh, Yan Xia, Kamil Charubin, and Ying Liu acknowledge the financial support from the National Science Foundation (NSF) (CMMI 1131587 and CBET 1433903) and NSF-DMR 1228817 for the acquisition of a high-sensitivity DSC. SANS experiments were conducted at SNS (ORNL) and NCNR (NIST). The authors thank Dr. Boualem Hammouda (NIST) for his help. John Katsaras is supported through the Scientific User Facilities Division of the United States Department of Energy (DOE) Office of Basic Energy Sciences (BES) under Contract DE-AC05-00OR22725, which also supported the MD simulation work. Xiaolin Cheng is partially supported by Laboratory Directed Research and Development (LDRD) Fund P7394 (ORNL).

Figure 9. DSC endotherms for DPPC/DPPG ULV (R = 0.05) and DPPC/DPPG/DHPC bicelle (Q = 3, and R = 0.05). The minor peak around 42 °C in the case of bicelles is presumably from vesicles.

bicelles. The melting transition enthalpy, ΔHfu, of bicelles (18.2 kJ/mol) is smaller than that of ULVs (35.2 kJ/mol), indicating that less DPPC molecules are in the gel phase and are presumably located in interfacial domains or in the vicinity of the disk’s rim. The broadening of the transition peak for bicelles is indicative of a loss in cooperativity.50,51 Interestingly, the TM of DPPC in bicelles is shifted roughly 3 °C higher than that of ULVs, presumably resulting from the reduction of Laplace pressure, which is regulated by curvature (i.e., flat disk’s plane) and affects the melting point depression in other micellar systems.22,52 The fact that ΔH⧧ of bicelles (116 kJ/mol) derived from TR-SANS is significantly larger than ΔHfu (18.2 kJ/mol) implies that a higher energy is required for dislodging acyl chains from the bilayer to the water phase than is required to melt them. On the other hand, the change in the entropy of fusion, ΔSfu, of bicelles (0.057 kJ K−1 mol−1) is larger than the ΔS⧧ (=0.03 kJ K−1 mol−1) obtained from TR-SANS, indicating that the transferred acyl chains are less ordered than those of gel DPPC prior to transfer. This is different from polymer exchange in polymeric micelles (reported by Zinn et al.), where ΔS⧧ was found to be higher than ΔSfu as a result of the gain in conformational degree of freedom of a n-alkane chain in solution.52,53 The lower ΔS⧧ of bicelles suggests that lipid transfer in this system takes place between loosely packed

■

REFERENCES

(1) Ivanov, A. I. Pharmacological Inhibition of Endocytic Pathways: is it Specific Enough to be Useful? Methods Mol. Biol. 2008, 440, 15−33. (2) Gruenberg, J.; Maxfield, F. R. Membrane Transport in the Endocytic Pathway. Curr. Opin. Cell Biol. 1995, 7 (4), 552−63. (3) Maranhão, R. C.; Freitas, F. R. HDL Metabolism and Atheroprotection: Predictive Value of Lipid Transfers. Adv. Clin. Chem. 2014, 65, 1−41. (4) Alpy, F.; Tomasetto, C. START Ships Lipids Across Interorganelle Space. Biochimie 2014, 96, 85−95. (5) Neculai, D.; Schwake, M.; Ravichandran, M.; Zunke, F.; Collins, R. F.; Peters, J.; Neculai, M.; Plumb, J.; Loppnau, P.; Pizarro, J. C.; et al. Structure of LIMP-2 Provides Functional Insights with Implications for SR-BI and CD36. Nature 2013, 504, 172−6. (6) Rashid, S.; Genest, J. Effect of Obesity on High-density Lipoprotein Metabolism. Obesity 2007, 15, 2875−88. F

DOI: 10.1021/acs.langmuir.6b02099 Langmuir XXXX, XXX, XXX−XXX

Article

Langmuir (7) Balla, T. Phosphoinositides: Tiny Lipids with Giant Impact on Cell Regulation. Physiol. Rev. 2013, 93, 1019−137. (8) Pan, J.; Cheng, X.; Monticelli, L.; Heberle, F. A.; Kučerka, N.; Tieleman, D. P.; Katsaras, J. The Molecular Structure of a Phosphatidylserine Bilayer Determined by Scattering and Molecular Dynamics Simulations. Soft Matter 2014, 10, 3716−25. (9) Heberle, F. A.; Petruzielo, R. S.; Pan, J.; Drazba, P.; Kučerka, N.; Standaert, R. F.; Feigenson, G. W.; Katsaras, J. Bilayer Thickness Mismatch Controls Domain Size in Model Membranes. J. Am. Chem. Soc. 2013, 135, 6853−9. (10) Dürr, U. H.; Gildenberg, M.; Ramamoorthy, A. The Magic of Bicelles Lights up Membrane Protein Structure. Chem. Rev. 2012, 112, 6054−74. (11) Hu, W.; Webb, L. J. Direct Measurement of the Membrane Dipole Field in Bicelles using Vbrational Stark Effect Spectroscopy. J. Phys. Chem. Lett. 2011, 2, 1925−1930. (12) Hamada, T.; Hagihara, H.; Morita, M.; Vestergaard, M. C.; Tsujino, Y.; Takagi, M. Physicochemical Profiling of SurfactantInduced Membrane Dynamics in a Cell-Sized Liposome. J. Phys. Chem. Lett. 2012, 3, 430−5. (13) Sanders, C. R.; Hare, B. J.; Howard, K. P.; Prestegard, J. H. Magnetically-Orientated Phospholipid Micelles as a Tool for the Study of Membrane-Associated Molecules. Prog. Nucl. Magn. Reson. Spectrosc. 1994, 26, 421−444. (14) Sanders, C. R.; Prosser, R. S. Bicelles: a Model Membrane System for all Reasons? Structure 1998, 6, 1227−1234. (15) Sanders, C. R.; Oxenoid, K. Customizing Model Membranes and Samples for NMR Spectroscopic Studies of Complex Membrane Proteins. Biochim. Biophys. Acta, Biomembr. 2000, 1508, 129−45. (16) Marcotte, I.; Auger, M. Bicelles as Model Membranes from Solid- and Solution-State NMR Studies of Membrane Peptides and Proteins. Concepts Magn. Reson., Part A 2005, 24A, 17−37. (17) Vestergaard, M.; Kraft, J. F.; Vosegaard, T.; Thogersen, L.; Schiott, B. Bicelles and Other Membrane Mimics: Comparison of Structure, Properties, and Dynamics from MD Simulations. J. Phys. Chem. B 2015, 119, 15831−43. (18) Lee, H.; Pastor, R. W. Coarse-Grained Model for PEGylated Lipids: Effect of PEGylation on the Size and Shape of Self-Assembled Structures. J. Phys. Chem. B 2011, 115, 7830−7. (19) Hu, A.; Fan, T.-H.; Katsaras, J.; Xia, Y.; Li, M.; Nieh, M.-P. Lipid-based Nanodiscs as Models for Studying Mesoscale Coalescence–a Transport Limited Case. Soft Matter 2014, 10, 5055−60. (20) Yang, P.-W.; Lin, T.-L.; Hu, Y.; Jeng, U.-S. A time-Resolved Study on the Interaction of Oppositely Charged Bicelles–Implications on the Charged Lipid Exchange Kinetics. Soft Matter 2015, 11, 2237− 42. (21) Dong, H.; Lund, R.; Xu, T. Micelle Stabilization via Entropic Repulsion: Balance of Force Directionality and Geometric Packing of Subunit. Biomacromolecules 2015, 16, 743−7. (22) Kastantin, M.; Ananthanarayanan, B.; Karmali, P.; Ruoslahti, E.; Tirrell, M. Effect of the Lipid Chain Melting Transition on the Stability of DSPE-PEG(2000) Micelles. Langmuir 2009, 25, 7279−86. (23) Dong, H.; Shu, J. Y.; Dube, N.; Ma, Y.; Tirrell, M. V.; Downing, K. H.; Xu, T. 3-Helix Micelles Stabilized by Polymer Springs. J. Am. Chem. Soc. 2012, 134 (28), 11807−14. (24) Jain, S.; Bates, F. S. Consequences of Nonergodicity in Aqueous Binary PEO-PB Micellar Dispersions. Macromolecules 2004, 37, 1511− 1523. (25) Lu, J.; Choi, S.; Bates, F. S.; Lodge, T. P. Molecular Exchange in Diblock Copolymer Micelles: Bimodal Distribution in Core-block Molecular Weights. ACS Macro Lett. 2012, 1, 982−985. (26) Xia, Y.; Li, M.; Charubin, K.; Liu, Y.; Heberle, F. A.; Katsaras, J.; Jing, B.; Zhu, Y.; Nieh, M.-P. Effects of Nanoparticle Morphology and Acyl Chain Length on Spontaneous Lipid Transfer Rates. Langmuir 2015, 31, 12920−8. (27) Lund, R.; Willner, L.; Richter, D.; Dormidontova, E. E. Equilibrium Chain Exchange Kinetics of Diblock Copolymer Micelles: Tuning and Logarithmic Relaxation. Macromolecules 2006, 39, 4566− 4575.

(28) Choi, S. H.; Lodge, T. P.; Bates, F. S. Mechanism of Molecular Exchange in Diblock Copolymer Micelles: Hypersensitivity to Core Chain Length. Phys. Rev. Lett. 2010, 104, 047802. (29) Zinn, T.; Willner, L.; Lund, R.; Pipich, V.; Richter, D. Equilibrium Exchange Kinetics in n-alkyl-PEO Polymeric Micelles: Single Exponential Relaxation and Chain Length Dependence. Soft Matter 2012, 8, 623−626. (30) Nakano, M.; Fukuda, M.; Kudo, T.; Endo, H.; Handa, T. Determination of Interbilayer and Transbilayer Lipid Transfers By Time-resolved Small-angle Neutron Scattering. Phys. Rev. Lett. 2007, 98, 238101. (31) Bayerl, T. M.; Schmidt, C. F.; Sackmann, E. Kinetics of Symmetric and Asymmetric Phospholipid Transfer between Small Sonicated Vesicles Studied by High-sensitivity Differential Scanning Calorimetry, NMR, Electron Microscopy, and Dynamic Light Scattering. Biochemistry 1988, 27, 6078. (32) Heberle, F. A.; Doktorova, M.; Goh, S. L.; Standaert, R. F.; Katsaras, J.; Feigenson, G. W. Hybrid and Nonhybrid Lipids Exert Common Effects on Membrane Raft Size and Morphology. J. Am. Chem. Soc. 2013, 135 (40), 14932−5. (33) Williams, C. E.; Nierlich, M.; Cotton, J. P.; Jannink, G.; Boué, F.; Daoud, M.; Farnoux, B.; Picot, C.; DeGennes, P. G.; Rinaudo, M.; Moan, M.; Wolff, C. Polyelectrolyte Solutions-intrachain and Interchain Correlations Observed by SANS. J. Polym. Sci., Polym. Lett. Ed. 1979, 17 (6), 379−384. (34) Benoît, H.; Picot, C.; Benmouna, M. Structure Factor of a Polymer in a Complex System by Small-Angle Neutron-Scattering. J. Polym. Sci., Polym. Phys. Ed. 1984, 22 (8), 1545−1548. (35) Pronk, S.; Pall, S.; Schulz, R.; Larsson, P.; Bjelkmar, P.; Apostolov, R.; Shirts, M. R.; Smith, J. C.; Kasson, P. M.; van der Spoel, D.; Hess, B.; Lindahl, E. GROMACS 4.5: a High-Throughput and Highly Parallel Open Source Molecular Simulation Toolkit. Bioinformatics 2013, 29, 845−54. (36) Klauda, J. B.; Venable, R. M.; Freites, J. A.; O’Connor, J. W.; Tobias, D. J.; Mondragon-Ramirez, C.; Vorobyov, I.; MacKerell, A. D., Jr.; Pastor, R. W. Update of the CHARMM All-Atom Additive Force Field for Lipids: Validation on Six Lipid Types. J. Phys. Chem. B 2010, 114, 7830−43. (37) Guinier, A.; Fournet, G. Small-Angle Scattering of X-rays; Wiley: New York, 1955; pp 268. (38) Nagle, J. F.; Wiener, M. C. Structure of Fully Hydrated Bilayer Dispersions. Biochim. Biophys. Acta, Biomembr. 1988, 942, 1−10. (39) Glasstone, S.; Laidler, K. J.; Eyring, H. The Theory of Rate Processes; McGraw-Hill: New York, 1941; pp 100. (40) Homan, R.; Pownall, H. J. Transbilayer Diffusion of Phospholipids: Dependence on Headgroup Structure and Acyl Chain Length. Biochim. Biophys. Acta, Biomembr. 1988, 938, 155−66. (41) Aniansson, E. A. G.; Wall, S. N.; Almgren, M.; Hoffmann, H.; Kielmann, I.; Ulbricht, W.; Zana, R.; Lang, J.; Tondre, C. Theory of the Kinetics of Micellar Equilibria and Quantitative Interpretation of Chemical Relaxation Studies of Micellar Solutions of Ionic Surfactants. J. Phys. Chem. 1976, 80, 905−922. (42) McLean, L. R.; Phillips, M. C. Kinetics of Phosphatidylcholine and Lysophosphatidylcholine Exchange between Unilamellar Vesicles. Biochemistry 1984, 23, 4624−30. (43) Ihm, J.; Quinn, D. M.; Busch, S. J.; Chataing, B.; Harmony, J. A. Kinetics of Plasma Protein-Catalyzed Exchange of Phosphatidylcholine and Cholesteryl Ester between Plasma-Lipoproteins. J. Lipid Res. 1982, 23, 1328−1341. (44) Nakano, M.; Fukuda, M.; Kudo, T.; Miyazaki, M.; Wada, Y.; Matsuzaki, N.; Endo, H.; Handa, T. Static and Dynamic Properties of Phospholipid Bilayer Nanodiscs. J. Am. Chem. Soc. 2009, 131, 8308− 12. (45) Wang, H.; Nieh, M.-P.; Hobbie, E. K.; Glinka, C. J.; Katsaras, J. Kinetic Pathway of the Bilayered-Micelle to Perforated-Lamellae Transition. Phys. Rev. E: Stat. Phys., Plasmas, Fluids, Relat. Interdiscip. Top. 2003, 67, 060902. (46) Mahabir, S.; Small, D.; Li, M.; Wan, W.; Kučerka, N.; Littrell, K.; Katsaras, J.; Nieh, M.-P. Growth Kinetics of Lipid-based Nanodiscs to G

DOI: 10.1021/acs.langmuir.6b02099 Langmuir XXXX, XXX, XXX−XXX

Article

Langmuir Unilamellar Vesicles-a Time-Resolved Small Angle Neutron Scattering (SANS) Study. Biochim. Biophys. Acta, Biomembr. 2013, 1828, 1025− 35. (47) Nieh, M.-P.; Harroun, T. A.; Raghunathan, V. A.; Glinka, C. J.; Katsaras, J. Concentration-Independent Spontaneously forming Biomimetric Vesicles. Phys. Rev. Lett. 2003, 91, 158105. (48) Drazenovic, J.; Ahmed, S.; Tuzinkiewicz, N. M.; Wunder, S. L. Lipid Exchange and Transfer on Nanoparticle Supported Lipid Bilayers: Effect of Defects, Ionic Strength, and Size. Langmuir 2015, 31, 721−31. (49) Liu, Y.; Li, M.; Yang, Y.; Xia, Y.; Nieh, M.-P. The Effects of Temperature, Salinity, Concentration and PEGylated Lipid on the Spontaneous Nanostructures of Bicellar Mixtures. Biochim. Biophys. Acta, Biomembr. 2014, 1838, 1871−80. (50) Shaw, A. W.; McLean, M. A.; Sligar, S. G. Phospholipid phase transitions in homogeneous nanometer scale bilayer discs. FEBS Lett. 2004, 556, 260−4. (51) Marsh, D.; Watts, A.; Knowles, P. F. Cooperativity of the phase transition in single- and multibilayer lipid vesicles. Biochim. Biophys. Acta, Biomembr. 1977, 465, 500−14. (52) Zinn, T.; Willner, L.; Lund, R. Nanoscopic Confinement through Self-assembly: Crystallization within Micellar Cores Exhibits Simple Gibbs-Thomson Behavior. Phys. Rev. Lett. 2014, 113 (23), 238305. (53) Zinn, T.; Willner, L.; Pipich, V.; Richter, D.; Lund, R. Effect of Core Crystallization and Conformational Entropy on the Molecular Exchange Kinetics of Polymeric Micelles. ACS Macro Lett. 2015, 4, 651−655.

H

DOI: 10.1021/acs.langmuir.6b02099 Langmuir XXXX, XXX, XXX−XXX