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Dehydration/rehydration cycles for mixing phospholipids without the use of organic solvents. Eric Oropeza-Guzman, and Jesus Carlos Ruiz Suarez Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.8b00799 • Publication Date (Web): 20 May 2018 Downloaded from http://pubs.acs.org on May 20, 2018

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is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

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Figure 1. Schematic workflow of the dehydration/rehydration method. Weighing of dry lipid powders (a), hydration and stirring with preheated deionized water (b), evaporation (c), and reconstitution of the mixed lipid film (d). 82x44mm (300 x 300 DPI)

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Figure 2. Molar heat capacity as a function of temperature for the four assayed mixtures using the dehydration/rehydration (D/S) method vs controls using organic solvent (OS) evaporation. Note the axis scaling in each box. 167x152mm (96 x 96 DPI)


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Figure 3. Molar heat capacity as a function of temperature for the DMPC/DPPC mixture using the dehydration/rehydration method. Discontinuous lines show the thermograms of each pure lipid species separately. The curve corresponding to ‘3 cycles’ is the same that the one depicted in the top-left corner of Figure 2. 167x116mm (96 x 96 DPI)


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For Table of Contents Only 82x44mm (300 x 300 DPI)


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Dehydration/rehydration cycles for mixing phospholipids without the use of organic solvents. Eric Oropeza-Guzman∗ and Jesús C. Ruiz-Suárez. Centro de Investigación y de Estudios Avanzados del IPN (CINVESTAV) Unidad Monterrey, Apodaca, Nuevo León 66600, México.

ABSTRACT. An environmentally friendly and straightforward dehydration/rehydration method for glycerophospholipid mixing that avoids the use of organic solvents, cosolvents or additives was developed. We prepared binary mixtures of zwitterionic and anionic glycerophospholipids using only deionized (DI) water in the entire mixing process. The resulting lipid films were subsequently reconstituted in vesicular form and compared to controls using differential scanning calorimetry (DSC). The calorimetric scans revealed no significant differences between mixing methods for any of the studied cases. These findings suggest that the developed dehydration/rehydration procedure creates a sample with equivalent compositional uniformity than the conventional solvent evaporation technique.

1. INTRODUCTION Phospholipid mixtures are essential for the preparation of physiologically relevant membrane models1 or effective drug delivery liposomes.2 Traditional lipid mixing has always relied on the use of suitable organic solvents for the dispersion and combination of phospholipid components.3


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Different phospholipids possess a variable degree of solubility in several organic phases which include, but are not limited to, acetone, short-chain alcohols, chlorinated solvents, n-alkanes, diethyl ether and combinations of these solvents in different ratios. The solubility of each phospholipid species depends on its molecular structure but is also modified by the presence of other species in the mixture.4 In the most time-consuming scenarios, each lipid component of a particular mixture must be dispersed separately in an appropriate solution of organic solvents. These single-lipid solutions are then mixed in the desired proportion, hoping that the whole will not phase separate and will result in a homogeneous film after solvent removal. As a final step of the sample preparation, traces of solvent are usually removed after a lengthy exposure under vacuum. Changing the proportion of any of the lipid components could in some cases compromise the stability of the mixture since different species may possess entirely different partition coefficients. The consequence is that each ensemble needs experimental characterization and fine-tuning of an appropriate solvent combination, which complicates the preparation process. Some of the organic solvents used for phospholipid solubilization are cataloged as undesirable due to their potential for human toxicity and environmental damage.5 Pharmaceutical liposome formulations that make use of these solvents in any step of the preparation process are regulated by the FDA and the US Pharmacopoeia.6 The use of ‘greener solvents’ with better environmental, health and safety (EHS) characteristics7 in both the laboratory and the industry remains a current topic of research that has significant economic implications.8 Several techniques using alternative solvents for liposome preparation have been proposed, including reverse phase evaporation,9 rapid injection,10 detergent depletion,11 supercritical fluids,12 membrane contactor,13 and the polyol dilution or heating method.14 However, to our


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knowledge, no technique has been developed that completely avoids the use of organic solvents, cosolvents or additives to mix phospholipids. In this work, we present a technically simple method to prepare mixed phospholipid films, that avoids the time-consuming process of characterizing an appropriate solvent combination for each case, since it only employs deionized water in every scenario. The method is based on the repeated dehydration and rehydration of a lipid-water system that contains the phospholipid species

to

be

mixed.

We

assayed

four

different

binary

mixtures

of

saturated

glycerophospholipids in equimolar proportion, comprising five different polar head groups (PC, PA, PG, PS, and PE), which involve both zwitterionic and anionic phospholipids. We see that the ability of the method to produce an equivalent compositional homogeneity to the traditional solvent evaporation technique is a consequence of two physicochemical properties of phospholipids, namely the thermodynamic interaction parameter and the lateral diffusion coefficient in the lamellar phase (see Discussion). 2. EXPERIMENTAL SECTION 2.1 Materials 1,2-dimyristoyl-sn-glycero-3-phosphocholine

(DMPC),

1,2-dipalmitoyl-sn-glycero-3-

phosphocholine (DPPC), 1,2-dipalmitoyl-sn-glycero-3-phospho-(1'-rac-glycerol) (sodium salt) (DPPG), 1,2-dipalmitoyl-sn-glycero-3-phosphate (sodium salt) (DPPA), 1,2-dipalmitoyl-snglycero-3-phosphoethanolamine (DPPE) and 1,2-dipalmitoyl-sn-glycero-3-phospho-L-serine (sodium salt) (DPPA) were purchased in dry powder form from Avanti Polar Lipids (Alabaster, USA) and used as received. Chloroform (anhydrous, ≥99%, with 0.5-1.0% ethanol as stabilizer), methanol (anhydrous, 99.8%) and dichloromethane (ACS reagent, ≥99.5%, with 50 ppm amylene as stabilizer) were purchased from Sigma-Aldrich (Toluca, México). Twice distilled


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water was treated with the Milli-Q IQ 7000 Ultrapure Water System from Merck Millipore México (Naucalpan de Juarez, México) prior to use. Phosphate buffered saline tablets from Sigma-Aldrich (1 tablet in 200 mL water) were used to obtain a 137 mM NaCl, 2.7 mM KCl and 10 mM PBS solution with pH 7.4 at 25 °C. 2.2 Methods The assayed lipid mixtures were: DMPC/DPPC (50/50) mol %, DPPC/DPPA (50/50) mol %, DMPC/DPPG (50/50) mol %, and DPPS/DPPE (50/50) mol %. Multilamellar vesicle samples were prepared using the proposed dehydration/rehydration method as schematically depicted in Figure 1.

Figure 1. Schematic workflow of the dehydration/rehydration method. Weighing of dry lipid powders (a), hydration and stirring with preheated deionized water (b), evaporation (c), and reconstitution of the mixed lipid film (d). Each phospholipid in powder form was weighed (4 µmoles) inside a clear scintillation vial using an analytical balance (OHAUS Explorer EX224). The balance was always calibrated before each use using an analytical weight set certified by Troemner (Weight ID Number 4000017899 – Certificate Number 852839) by testing with NIST Traceable Reference Standards (NIST Test Number 822-275872-11). Variable volume manual pipettes (Eppendorf Reference 2)


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were checked to dispense within manufacturer reported tolerances before each working day, using the gravimetric tool programmed into the OHAUS Explorer EX224 balance. The vial containing the lipid powder was preheated for 15 min at the working temperature (i.e., 10 °C above the main transition temperature of the highest melting lipid in the mixture). Double distilled deionized water (100 µL) at the same temperature was added to the dry lipid powder inside the vial using the technique recommendations of the pipette manufacturer. The resulting lipid paste was stirred at 200 rpm with a small magnetic bar while keeping the vial above a lab stove at the same temperature. The vial was maintained on the stove for about 12 minutes until the lipid was dried to the naked eye. After drying, deionized water was added again, and this cycle was repeated three times. After three cycles of drying and rehydration, phosphate buffered saline (137 mM NaCl, 2.7 mM KCl and 10 mM PBS, pH 7.4) was added to the dry mixed lipid film to achieve a final total lipid concentration of 5 mM. Multilamellar vesicle controls for the same four mixtures were prepared using the solvent evaporation method. Each phospholipid was weighed in separate vials, and the appropriate organic solvent was added to disperse the dry powder according to Table 1. Table 1. Solubilization parameters for each phospholipid used in the mixtures. Phospholipid Solvent components

Solubilization temperature

DPPC

Chloroform

Room temperature

DMPC

Chloroform

Room temperature

DPPA

Chloroform/methanol/dichloromethane (2:1:1 v/v/v)

35 °C

DPPE

Chloroform/methanol (2:1 v/v)

35 °C

DPPG


Room temperature

DPPS


35 °C


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After dispersing each phospholipid, equimolar amounts were mixed in the same vial. The organic phase was then removed in a fume hood with a nitrogen stream until dry to the naked eye. The mixed dry lipid film was kept under vacuum for 60 minutes to remove remaining solvent traces. Afterward, phosphate buffered saline (137 mM NaCl, 2.7 mM KCl and 10 mM PBS, pH 7.4) was added to reach a final total lipid concentration of 5 mM. 2.3 Vesicle processing and characterization Multilamellar vesicles were processed the same way independently of the mixing method. A total of 1 mL of vesicles (5 mM) was loaded into a syringe and extruded 15 times using the Avanti Mini Extruder that was preheated to the working temperature. A polycarbonate membrane with a pore size of 100 nm was used. The quality of the extrusion products was checked for all samples using dynamic light scattering with a Zetasizer Nano ZS equipment (Malvern, Worcestershire, UK). After extrusion, all the samples were equilibrated to room temperature right before calorimetric analysis. Samples were loaded into a differential scanning calorimeter (Nano DSC, TA Instruments). After 800 seconds of equilibration time, a heating scan was performed at a rate of 1 °C/min and a cell pressure of 3 atmospheres. 3. RESULTS AND DISCUSSION Figure 2 shows representative scans after baseline subtraction. The obtained curves are characteristic of extruded unilamellar suspensions of liposomes, having a relatively broad transition half-width with a small cooperative unit size, which results in much lower maximum heat capacity values in comparison with those of multilamellar systems. Main transition temperature (Tm) and enthalpy change (∆H) were characterized from each of these scans. Table 2


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shows the arithmetic mean and standard deviation of these calorimetric parameters using three independent experiments from each mixture and mixing method.

Figure 2. Molar heat capacity as a function of temperature for the four assayed mixtures using the dehydration/rehydration (D/S) method vs controls using organic solvent (OS) evaporation. Note the axis scaling in each box. Table 2. Calorimetric parameters for each phospholipid mixture using the compared methods of lipid mixing. DMPC/DPPC (50/50) mol % Tm (°C)

∆H (kJ/mol*K)

Organic solvent evaporation

33.05 ± 0.15

28.37 ± 0.79

Dehydration/rehydration

32.85 ± 0.15

29.39 ± 0.74

Tm (°C)

∆H (kJ/mol*K)


54.64 ± 0.18

35.60 ± 0.74


54.56 ± 0.15

34.59 ± 0.73

DPPA/DPPC (50/50) mol %

DMPC/DPPG (50/50) mol %


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Tm (°C)

∆H (kJ/mol*K)


33.47 ± 0.12

28.07 ± 0.78


33.65 ± 0.14

29.09 ± 0.76

Tm (°C)

∆H (kJ/mol*K)


59.90 ± 0.07

38.92 ± 0.58


59.97 ± 0.04

38.30 ± 0.66

DPPE/DPPS (50/50) mol %

We can see that the resulting thermograms for the dehydration/rehydration method show almost identical curves than those obtained from liposome samples prepared with the traditional organic solvent evaporation technique. This result suggests that, for these four mixtures, the selected process of phospholipid mixing shows no differential effect on the compositional uniformity of the system. Whenever an equimolar lipid mixture contains a random molecular distribution of its components (i.e., no lipid demixing), a single endothermic peak is obtained on thermal analysis.15 On the contrary, when lipid species in the mixture have a thermodynamic preference for phase separation, more than one peak appears.16 Each of the mixtures explored in this work generates a single endothermic peak whether one uses the traditional mixing technique or the proposed dehydration/rehydration method. It has been reported that lipid mixing methods that involve a step of film formation can have a high degree of compositional heterogeneity.17 This observation is evident when the phospholipid species in the mixture have a low degree of lipid to lipid miscibility (non-ideal interaction) because they suffer from demixing during drying and film formation. To circumvent this problem, mixing methods that do not involve a step of film formation have been reported in the literature.18 Even if they need to incorporate the use of cosolvents like ethanol, they avoid the use


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of substances with worst EHS profiles and seem to improve the compositional homogeneity of non-ideal mixtures. On the other hand, lipid mixtures where the interaction between its components is close to ideal, produce a degree of homogeneity that seems to be independent of the preparation route.18 The four binary equimolar mixtures that we selected are classified as having ‘high' to ‘very high' miscibility according to the modified Silvius scheme.19 In a thermodynamic context, the final supramolecular homogeneity of these mixtures should be a state function that is independent of the mixing process (path). From a practical standpoint, this conclusion would also mean that different lipid mixtures could be prepared by a standardized method. We decided to take advantage of this observation and hypothesized that mixtures of miscible species should be able to reach compositional homogeneity in the medium with the best EHS characteristics that there is (i.e., water), without the need to be dissolved in proper solution beforehand. However, upon direct hydration with an aqueous medium, and due to their amphiphilic character, phospholipids spontaneously arrange into a series of closed concentric bilayer structures (i.e., multilamellar vesicles) that restrict their ability to create a homogeneously combined system. For the phospholipid molecules to be able to reach equilibrium and distribute homogeneously, there must be a way to exchange phospholipid molecules between separate bilayers within the lipidwater system. This part is where the repeated drying and rehydration of the sample becomes critical. The optimal number of dehydration/rehydration cycles was determined empirically for the DMPC/DPPC mixture first and then applied equally to the other mixtures. Extruded liposomes obtained from dried lipid films with less than three cycles of dehydration/rehydration were prepared and analyzed. The resulting curves are depicted in Figure 3. As expected, simple direct


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hydration of mixed lipid powders is unable to produce a homogeneous system, but each dehydration/rehydration cycle progressively contributes to merge the endothermic peaks that initially correspond to each one of the pure lipid species.

Figure 3. Molar heat capacity as a function of temperature for the DMPC/DPPC mixture using the dehydration/rehydration method. Discontinuous lines show the thermograms of each pure lipid species separately. The curve corresponding to ‘3 cycles’ is the same that the one depicted in the top-left corner of Figure 2. The gradual merging of endotherms reflects the ability of the phospholipid molecules to distribute into the whole lamellar system provided they start to gain access to all the bilayers. During the drying phase of each mixing cycle, these bilayers intimately approximate as the water content is reduced. Published experiments using the surface forces apparatus (SFA) technique, point out to the conclusion that fusion between two bilayer surfaces can spontaneously occur when they come within about 1 nm of each other.20 The bilayers are thought to overcome the repulsive forces between them via highly localized instabilities in the membrane integrity. These instabilities promote the hydrophobic attraction between the internal alkyl chains of the apposed membranes, resulting in hemifusion (i.e., transmonolayer contact) and lipid mixing.


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Theoretical models of intimate bilayer apposition seem to corroborate this finding using molecular dynamics simulations.21 These authors show that even when all the phospholipid molecules in the bilayers are saturated with water, the proximity to each other seems to induce a reorientation of the alkyl chains. Some of the chains were seen to lie parallel to the membrane plane or to protrude out of the bilayer. The number of misoriented alkyl chains was counted after a total simulation time of 2 ns, but reorientations occurred almost instantaneously (within several tens of picoseconds) after the two membranes approached each other just due to thermal fluctuations. Evidence of lipid mixing has also been retrieved in other experimental conditions that compromise membrane integrity like the case where sonicated liposomes are dried,22 or when they are freeze-thawed.23 In the former case, the investigators report that the size and unilamellarity of the liposomes (SUVs) seem to be a necessary condition for successful mixing during drying. Whereas in the latter case, the authors conclude that lipid mixing is the result of the ‘breakdown and reformation' of the bilayer structure during freeze-thawing. In our experiments, close apposition between bilayers certainly occurs during the evaporation of the aqueous phase, long before the lipid-water system has dried completely. On the other hand, the rapid escape of water molecules during this process has the potential to increase the rate of local instabilities and bilayer breakdowns. This mechanism would permit the lateral diffusion of phospholipid molecules between neighboring vesicles in the system, even when the evaporating liposomes are multilamellar and polydisperse in size. Experimental data from published pulse field gradient (PFG) NMR measurements of lipid selfdiffusion,24 and FRAP methods using fluorescent probes25,26 report that the phospholipid lateral diffusion coefficient in lamellar phases of DMPC or DPPC is always above 15 µm2s-1 for


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temperatures around 60 ºC. Theoretical estimations from coarse-grained models of DPPC bilayers arrive at a lateral diffusion value of about 20 µm2s-1 for the fluid phase at the same temperature.27 These simulations also reveal that the diffusion constants seem to be very sensitive to the number of lattice defects in the bilayers, with up to 5-fold increments in the diffusion constants compared to ideal defect-free membranes. With such values in mind, it is reasonable to assume that an increasingly homogeneous lipid distribution can be expected within the time frame of each evaporation cycle in our samples. We paid particular attention to the mixture using DPPS and DPPE since the latter lipid cannot be dispersed from its dried crystal powder using deionized water alone, even when the temperature of the system is above the temperature of its melting transition. The finding that a mixture with this lipid can be prepared by the dehydration/rehydration method implies that the energy of interaction between DPPS and DPPE molecules, in the aqueous lamellar phase, is lower than the energy of interaction between DPPE molecules themselves in the dry crystal phase Lc. This conclusion would be consistent with a negative value for the free energy of mixing in the case of the DPPS-DPPE pair. It has been reported that this is in fact, the case.28 The negative value is interpreted as a net attractive interaction between unlike lipids in the mixture, which may be the reason for the aqueous solvation of otherwise insoluble DPPE crystals. It would be ideal to find a negative value for the free energy of mixing in the case of the DPPC-Chol pair. Successful incorporation of cholesterol by this method would permit a technically simple and solvent-free preparation amenable for example to lipid raft modeling. However, this could hardly be the case, since the solubility of cholesterol in aqueous solutions is negligible29 and it is known that an energetically favorable interaction between the PC/Chol pair


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is observed only in the liquid-ordered phase (Lo).30 Besides, it has been reported that cholesterol dramatically suffers from the problem of demixing during drying and film formation that was also observed in the case of mixtures containing lipid components with low lipid to lipid miscibility.31 Pre-experiments performed in our laboratory readily confirmed our suspicions, and cholesterols crystals were easily seen after macroscopic inspection of the samples, independently of the number of D/R cycles performed. 4. CONCLUSION We proposed a standardized lipid mixing method that uses deionized water as the only mixing agent in every case, avoiding the need to characterize an appropriate organic solvent combination for different sets of lipid components. After three cycles of drying and rehydration, the DSC characterization of our samples showed equivalence with the traditional solvent evaporation technique. From a technical point of view, the method provides a valuable advantage over the traditional method, since it simplifies and speeds up the preparation process. It also benefits from requiring only simple laboratory tools without the need for high-pressure equipment or special installations. From the environmental health and safety standpoint, we are confident that the technique can contribute to the interest that the research community has expressed on the implementation of sustainable chemical practices,32 or situations where organic solvent use is closely regulated. We are sure that this technique constitutes a fast, solvent-free and technically simple standard method for mixing miscible lipids, especially useful in situations where several iterations must be made to study the effect that the titration of a lipid constituent has on the physicochemical properties of a mixed system. Moreover, it can be implemented as a complement to previously published techniques that make use of alternative solvents with


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reasonable EHS profiles when the need to incorporate components with negligible solubilities in aqueous solutions arises. AUTHOR INFORMATION Corresponding Author [email protected] Notes The authors declare no competing financial interests. ACKNOWLEDGMENT This work was supported by the “Consejo Nacional de Ciencia y Tecnología” CONACYT, under project Fronteras de la Ciencia-2-1132. E. Oropeza-Guzman acknowledges a scholarship from CONACYT. REFERENCES (1) van Meer, G.; Voelker, D. R.; Feigenson, G. W. Membrane lipids: where they are and how they behave. Nat. Rev. Mol. Cell Biol. 2008, 9, 112-124. (2) Pattni, B. S.; Chupin, V. V.; Torchilin, V. P. New Developments in Liposomal Drug Delivery. Chem. Rev. 2015, 115, 10938-10966. (3) Bangham, A. D. Membrane models with phospholipids. Prog. Biophys. Mol. Biol. 1968, 18, 29-95. (4) Dervichian, D. G. The physical chemistry of phospholipids. Prog. Biophys. Mol. Biol. 1964, 14, 263-342.


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(5) Dwivedi, A. M. Residual solvent analysis in pharmaceuticals. Pharm. Technol. 2002, 133, 42-46. (6) B’Hymer, C. Residual solvent testing: a review of gas-chromatographic and alternative techniques. Pharm. Res. 2003, 20, 337-344. (7) Capello, C.; Fischer, U.; Hungerbühler, K. What is a green solvent? A comprehensive framework for the environmental assessment of solvents. Green Chem. 2007, 9, 927-934. (8) Clarke, C. J.; Tu, W.-C.; Levers, O.; Bröhl, A.; Hallett, J. P. Green and Sustainable Solvents in Chemical Processes. Chem. Rev. 2018, 118, 747-800. (9) Cortesi, R.; Esposito, E.; Gambarin, S.; Telloli, P.; Menegatti, E.; Nastruzzi, C. Preparation of liposomes by reverse-phase evaporation using alternative organic solvents. J. Microencapsul. 1999, 16, 251-256. (10) Batzri, S.; Korn, E. D. Single bilayer liposomes prepared without sonication. Biochim. Biophys. Acta - Biomembr. 1973, 298, 1015-1019. (11) Brunner, J.; Skrabal, P.; Hausser, H. Single bilayer vesicles prepared without sonication physico-chemical properties. Biochim. Biophys. Acta - Biomembr. 1976, 455, 322-331. (12) Otake, K.; Imura, T.; Sakai, H.; Abe, M. Development of a New Preparation Method of Liposomes Using Supercritical Carbon Dioxide. Langmuir 2001, 17, 3898-3901. (13) Charcosset, C.; El-Harati, A.; Fessi, H. Preparation of solid lipid nanoparticles using a membrane contactor. J. Control. Release 2005, 108, 112-120.


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(14) Mozafari, M. R.; Reed, C. J.; Rostron, C.; Kocum, C.; Piskin, E. Construction of stable anionic liposome-plasmid particles using the heating method: a preliminary investigation. Cell. Mol. Biol. Lett. 2002, 7, 923-927. (15) Krisovitch, S. M.; Regen, S. L. Nearest-neighbor recognition in phospholipid membranes: a molecular-level approach to the study of membrane suprastructure. J. Am. Chem. Soc. 1992, 114, 9828-9835. (16) Silvius, J. R. Solid- and liquid-phase equilibria in phosphatidylcholine / phosphatidylethanolamine mixtures. A calorimetric study. Biochim. Biophys. Acta - Biomembr. 1986, 857, 217-228. (17) Larsen, J.; Hatzakis, N. S.; Stamou, D. Observation of Inhomogeneity in the Lipid Composition of Individual Nanoscale Liposomes. J. Am. Chem. Soc. 2011, 133, 10685-10687. (18) Elizondo, E.; Larsen, J.; Hatzakis, N. S.; Cabrera, I.; Bjørnholm, T.; Veciana, J.; Stamou, D.; Ventosa, N. Influence of the Preparation Route on the Supramolecular Organization of Lipids in a Vesicular System. J. Am. Chem. Soc. 2012, 134, 1918-1921. (19) Marsh, D. Handbook of Lipid Bilayers, 2nd ed.; CRC Press: Boca Raton, FL, USA, 2013. (20) Helm, C.; Israelachvili, J.; McGuiggan, P. Molecular mechanisms and forces involved in the adhesion and fusion of amphiphilic bilayers. Science. 1989, 246, 919-922. (21) Ohta-Iino, S.; Pasenkiewicz-Gierula, M.; Takaoka, Y.; Miyagawa, H.; Kitamura, K.; Kusumi, A. Fast Lipid Disorientation at the Onset of Membrane Fusion Revealed by Molecular Dynamics Simulations. Biophys. J. 2001, 81, 217-224.


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(22) Bhatia, T.; Husen, P.; Brewer, J.; Bagatolli, L. A.; Hansen, P. L.; Ipsen, J. H.; Mouritsen, O. G. Preparing giant unilamellar vesicles (GUVs) of complex lipid mixtures on demand: Mixing small unilamellar vesicles of compositionally heterogeneous mixtures. Biochim. Biophys. Acta Biomembr. 2015, 1848, 3175-3180. (23) MacDonald, R. I.; MacDonald, R. C. Lipid mixing during freeze-thawing of liposomal membranes as monitored by fluorescence energy transfer. Biochim. Biophys. Acta - Biomembr. 1983, 735, 243-251. (24) Lindblom, G.; Orädd, G.; Filippov, A. Lipid lateral diffusion in bilayers with phosphatidylcholine, sphingomyelin and cholesterol. Chem. Phys. Lipids 2006, 141, 179-184. (25) Vaz, W. L. C.; Clegg, R. M.; Hallmann, D. Translational diffusion of lipids in liquid crystalline phase phosphatidylcholine multibilayers. A comparison of experiment with theory. Biochemistry 1985, 24, 781-786. (26) Almeida, P. F. F.; Vaz, W. L. C.; Thompson, T. E. Lateral diffusion in the liquid phases of dimyristoylphosphatidylcholine/cholesterol lipid bilayers: a free volume analysis. Biochemistry 1992, 31, 6739-6747. (27) Marrink, S. J.; Risselada, J.; Mark, A. E. Simulation of gel phase formation and melting in lipid bilayers using a coarse grained model. Chem. Phys. Lipids 2005, 135, 223-244. (28) Wydro, P. The interactions between cholesterol and phospholipids located in the inner leaflet of humane erythrocytes membrane (DPPE and DPPS) in binary and ternary films – The effect of sodium and calcium ions. Colloids Surfaces B Biointerfaces 2011, 82, 209-216.


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(29) Haberland, M. E.; Reynolds, J. a. Self-Association of Cholesterol in Aqueous Solution. Proc. Natl. Acad. Sci. 1973, 70, 2313-2316. (30) Almeida, P. F. A Simple Thermodynamic Model of the Liquid-Ordered State and the Interactions between Phospholipids and Cholesterol. Biophys. J. 2011, 100, 420-429. (31) Huang, J.; Buboltz, J. T.; Feigenson, G. W. Maximum Solubility of Cholesterol in Phosphatidylcholine and Phosphatidylethanolamine Bilayers. Biochim. Biophys. Acta – Biomembr. 1999, 1417, 89-100. (32) Horváth, I. T. Introduction: Sustainable Chemistry. Chem. Rev. 2018, 118, 369-371.


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