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Properties of diphytanoyl phospholipids at the air-water interface Anthony Yasmann, and Sergei Sukharev Langmuir, Just Accepted Manuscript • DOI: 10.1021/la503800g • Publication Date (Web): 04 Dec 2014 Downloaded from http://pubs.acs.org on December 10, 2014
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Properties of diphytanoyl phospholipids at the air-water interface Anthony Yasmann and Sergei Sukharev Department of Biology, University of Maryland College Park, MD 20742 Correspondence: [email protected], 301-405-6923
ABSTRACT: Diphytanoyl phosphatidyl choline (DPhPC) is a synthetic ester lipid with methylated tails found in archaeal ether lipids. Due to stability of DPhPC bilayers and absence of phase transitions in a broad range of temperatures, the lipid is used as an artificial membrane matrix for reconstitution of channels, pumps, and membrane-active peptides. We characterized monomolecular films made of DPhPC and its natural ether analog DOPhPC at the air-water interface. We measured compression isotherms and dipole potentials of films made of DPhPC, DPhPE and DOPhPC. We determined that at 40 mN/m the molecular area of DPhPC is 81.2 Å2, consistent with x-ray and neutron scattering data obtained in liposomes. This indicates that 40 mN/m is the monolayer-bilayer equivalence pressure for this lipid. At this packing density, the compressibility modulus (Cs-1= 122 ± 7 mN/m) and interfacial dipole potential (V= 355 ± 16 mV) were near their maximums. The molecular dipole moment was estimated as 0.64 ± 0.02 D. The ether DOPhPC compacted to 70.4 Å2/lipid at 40 mN/m displaying similar peak compressibility to DPhPC. The maximal dipole potential of the ether lipid was about half of that for DPhPC at this density and the elemental dipole moment was about a quarter. Spreading of DPhPC and DOPhPC liposomes reduced surface tension of the aqueous phase by 46 and 49 mN/m, respectively. This corresponds well to the monolayer collapse pressure. The equilibration time shortened as temperature increased from 20 oC to 60 oC, but surface pressure at equilibrium did not change. The data illustrates the properties of branched chains and contributions of ester bonds in setting the mechanical and electrostatic parameters of diphytanoyl lipids. These properties determine an environment in which reconstituted voltage- or mechano-activated proteins may function. Electrostatic properties are important in preparation of asymmetric folded bilayers, whereas lateral compressibility defines tension in mechanically stimulated droplet interface bilayers.
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Introduction Archaeal branched phospholipids are a special adaptation to the extreme environment which thermophilic organisms tolerate 1. The branched phytanoyl hydrocarbon chains remain in a liquid crystalline state in a wide range of temperatures. At the same time, the lipids retain an ordered bilayer structure due to steric crowding and conformational restraints imposed by methyl branches. These chains are resistant to oxidation and attached through ether linkages, chemically more stable than ester bonds, to the glycerol backbone. The stability of ether lipids provide a robust and stable lipid matrix for many archaea. Diphytanoyl phosphocholine (DPhPC) was synthesized for the purpose of creating an oxidation-resistant and stable lipid bilayers for in-vitro experiments with artificial membranes and reconstituted membrane proteins 2. It contains archaeal hydrocarbon chains but ester linkages resembling bacterial and eukaryotic phospholipids. DPhPC has been reported to have no gel to liquid crystalline phase transition from -120 °C to 120 °C3. Methyl branches, evenly spaced along the chains (Figure 1), reduce the density of packing and inter-chain Van-der-Waals interactions, which suppresses phase transitions more than cis-unsaturation kinks in linear hydrocarbon chains. The absence of phase transitions and related instabilities is a desired trait for many practical uses of the lipid; this advantage is reflected by numerous publications using DPhPC as a matrix for reconstitution of channels, peptides, and transporters4, 5, 6. Despite many experimental uses such as preparation of liposomes 7, planar bilayers 8, supported bilayers 9, or droplet interface bilayer (DIBs)10, 11 the properties of DPhPC compared to its ether diphytanoyl DOPhPC analog and nonbranched phospholipids have not been exhaustively studied. Branched phytanoyl chains increase the in-plane cross-section of the molecule, possibly exposing more hydrocarbon to water. The polar carbonyl oxygens of DPhPC, substituting less polar ether oxygens of DOPhPC, reside in that boundary region, which may alter the character of solvation and hydrogen bonding in this interfacial zone. As a result, the mechanics and electrostatics of the interfacial layer may be altered. X-ray and neutron scattering experiments determined that the molecular area of DPhPC in liposomes is near 80 Å2, which is 16 Å2 larger than that of the non-branched dipalmitoylphosphatidylcholine (DPPC) of similar length 12, 13, but no comparison with DOPhPC was reported. There have been several comparative studies of ester and ether phospholipid analogs, showing that ester lipids exhibited an area 7-10 Å2/molecule larger at all surface pressures compared to analogs with one 14 or two ether bonds 15. A number of studies marked a notable decrease in surface potential and dipole moment of pure ether lipids 16, 17, 18 or etherester hybrid phospholipids 19 compared to their ester analogs. Use of diphytanoyl lipids for reconstitution warrants investigation into the equilibrium between their vesicular and monolayer forms. Vesicles open up at the air-water or oil-water interfaces forming monolayers, a property that is widely used to form folded bilayers 20 or DIBs 21, 22. On the other hand, at high lateral pressures monolayers collapse producing vesicles. Whether the same equilibrium between the vesicular and monolayer forms is achieved in both processes is 2 ACS Paragon Plus Environment
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unclear. This pressure resulted from spreading of liposomes made of saturated lipids at the airwater interface was shown to have strong temperature dependence 23. Although it is known that DPhPC monolayers collapse at high lateral pressures, the equilibria between the interfacial and vesicular forms have not been studied. In addition, the compression elasticity modulus of artificial monolayers is becoming an important parameter for tension estimations when droplet interface bilayers (DIBs) are subjected to mechanical distortion in experiments with reconstituted mechanosensitive channels.24 In the present paper, we compare interfacial properties of ester and ether diphytanoyl phosphocholines. We report area per head group at monolayer-bilayer equivalence pressures, compressibility, and dipolar properties of these lipids. We also present data on the liposome spreading kinetics and equilibria, illustrating an almost thermodynamically ‘ideal’ behavior of these lipids.
Figure 1. Chemical structures of DPhPE, DPhPC and its natural ether analog DOPhPC.
Methods and Materials 1,2-diphytanoyl-sn-glycero-3-phosphocholine (DPhPC), 1,2-diphytanoyl-sn-glycero-3phosphoethanolamine (DPhPE), and 1,2-di-O-phytanyl-sn-glycero-3-phosphocholine (DOPhPC) were purchased from Avanti Polar Lipids (Alabaster, AL, USA). Potassium Chloride, Trizma® Hydrochloride, Potassium Hydroxide, and Hexane were acquired from Sigma-Aldridge (St. Louis, MO, USA). All solutions were prepared with ultrapure 18.2 MΩ x cm water from a Barnstead 3 ACS Paragon Plus Environment
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Nanopure system. All monolayer and tensiometry experiments were performed in a dust-free laminar flow hood (AirClean® Systems, Raleigh, NC, USA). Lipid stocks. Each lipid was dried under stream of nitrogen gas in pre-weighed air tight vials. Lipids were placed overnight under deep vacuum (10-20 µmHg) with a solvent trap. DPhPC, DOPhPC, and DPhPE were reconstituted in hexane by weight to a stock concentration. Buffers. A 10X stock buffer was prepared to a concentration of 1.5 M KCl and 250 mM Tris in ultrapure water and titrated to a pH of 7.2 using KOH. The buffer was filtered through a 2 µm bottle top filter (Nalgene™ Rapid-Flow™) and stored in glass. All glassware was acidwashed and thoroughly rinsed. Preparations of Liposomes. Lipids were dried from hexane solutions in round-bottom disposable glass tubes first under stream of nitrogen and then under vacuum (4 hrs), rehydrated for an hour in buffer to a concentration of 6.25 mg/mL. The solution was then vigorously vortexed, and sonicated for a minute at room temperature in a Branson-type sonicator (Aquasonic 50t, VWR, West Chester, PA, USA). Liposomes were then diluted to 5 mg/mL. Pressure-area (π-A) isotherms. Surface tensions of lipids were observed using Wilhelmy method with a filter paper strip (Whatman, No. 1, 10.5 mm wide and 0.25 mm thick) as the plate. The pressure sensor (model 601, NIMA, Coventry, U.K.) was calibrated with a 100 mg weight, after calibration surface tension of pure water was determined to be -72 mN/m. The sensor was zeroed to provide negative measures for surface tension. A rectangular Teflon® monolayer trough (max area of ̴550 cm2) with a single barrier (NIMA) were used in the pressure-area isotherm experiments. The stock solutions of DPhPC, DOPhPC, and DPhPE were diluted by weight to final concentrations of 1.2 mg/mL, 1.2 mg/mL, and 1.4 mg/mL respectively. The 30 µL of lipids were spread onto the subphase using a gastight 50µL Hamilton syringe for all lipids. A 150 mM KCl and 25 mM Tris (pH 7.2) buffer was the subphase in all experiments. After allowing time for solvents to evaporate (15 min), the area compression isotherms were performed at room temperature (22 °C) at a speed of 150 cm2/min. The onset of the pressure change with film compression, the collapse pressure and the lateral compressibility modulus Cs1 = A(dπ/dA) were determined from the curves. Potentials-area (V-A) isotherms. Potentials were measured using potential sensor MicroSpot (Kibron, Helsinki, Finland) in a Kibron MicroTroughXS. A rectangular aluminum trough (max area of ̴104 cm2) with two mobile barriers (Kibron) were used in the potential experiments. The stock solutions of DPhPC, DOPhPC, and DPhPE were diluted by weight to final concentrations of 0.2 mg/mL each. Lipid spreading method and the buffer used were identical to the pressure area isotherm experiments. Area compression of the monolayer was repeated as in π-A isotherm experiments at a speed of 3 mm/min. Spreading Kinetics. Surface spreading kinetics were observed using Wilhelmy method with a Kibron MicroTroughXS sensor and flamed wire (Kibron, proprietary alloy, 0.51 mm diameter) as the Wilhelmy plate in a 2 x 3 array of 200 µL wells custom-made out of Teflon. The tensiometer was calibrated by contrasting voltage measurements of air and ultrapure water as per the Kibron manual. Control measurements of the buffer were taken at 20 °C. Buffer and 4 ACS Paragon Plus Environment
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liposomes were then heated by water bath to each experimental temperature. Buffer was placed into the well and recordings were then infused with liposomes to a final concentration of 1 mg/ml. The wells were cleaned with chloroform between trials. Dynamic Light Scattering. Liposome sizes were measured by dynamic light scattering with a 90Plus Particle Size Analyzer (Brookhaven Instruments Corporation, Holtsville, NY, USA). The analyzer was standardized with a 90 nm calibrating particles (Duke Scientific Corporation, Palo Alto, CA, USA). Liposomes were freshly sonicated and diluted from 6.25 mg/mL to 0.1 mg/mL in subphase buffer. Measurements were performed in polystyrene cuvettes (Brookhaven Instruments Corporation, square, 10mm, 4.5 mL) at room temperature (22 °C).
Results Pressure-area (π-A) isotherms. Compression isotherms for lipid monolayers formed from DPhPC, DOPhPC, and DPhPE are shown in Figure 2A. DPhPC and DPhPE begin showing changes in surface pressure at molecular area around 140 Å2 while DOPhPC change is observed near 130 Å2. The collapse pressure of DPhPC and DOPhPC films was observed as roughly 45 mN/m with DPhPE collapsing around 47 mN/m. At 40 mN/m considered to be the monolayer-bilayer equivalence point25 the area per molecule is 78.3 Å2 ± 5.1 Å2 (n=7 for DPhPC, 70.4 Å2 ± 1.5 Å2 (n=7) for DOPhPC, and 81.3 Å2 ± 0.3 Å2 (n=7) for DPhPE. Compressibility modulus. Isotherms of each lipid were converted to compressibility modulus curves (Figure 2B) as described in 26. DPhPE is the stiffest lipid of the group having the highest compressibility modulus near 160 mN/m. DOPhPC indicates tighter packing than both DPhPC and DPhPE with a left shift of the compression modulus curve and a lower peak of about 115 mN/m. DPhPC exhibited compressibility modulus peaking near 125 mN/m. Areas per lipid and compressibility moduli for the three lipids are presented together with electrostatic parameters in Table 1.
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Figure 2. Monolayer pressure-area (π-A) isotherms of DPhPC, DPhPE, and DOPhPC (A). Monolayer-bilayer equivalence seen at 78.3 Å2 ± 5.1 Å2 for DPhPC, 70.4 Å2 ± 1.5 Å2 for DOPhPC, and 81.3 Å2 ± 0.3 Å2 for DPhPE (n=7 for each lipid). Area compressibility modulus (B) over molecular area calculated from pressure-area (π-A) isotherms Cs-1 = A(dπ/dA) for DPhPC, DPhPE, and DOPhPC.
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Surface Potential and its area dependence. We consider the measured surface (volta) potential as purely dipole under the assumption that all lipids studied here are zwitterionic near physiological pH with no net charge, and that possible specific adsorption of ions at the interface are small. Figure 3 shows a group of surface potential –area curves as the lipid monolayers were compressed. The DPhPC curve displays the typical initial potential buildup to a plateau as area per molecule decreases, followed by a gradual increase to the second plateau near the collapse point. DOPhPC and DPhPE show a similar but, less defined behavior. DPhPC and DPhPE showed a similar surface potential profile during monolayer compression, with DPhPE surface potential curve being shifted to the left. DPhPC initial potential buildup appears at ̴220 Å per molecule and DPhPE potential build up is seen at ̴180 Å per molecule. DPhPC and DPhPE show a gradual change in potential as the lipid density increases that is greater than that of DOPhPC. DPhPC consistently had the largest change of potential during the monolayer compression peaking at ̴390 mV near the point of collapse ( ̴80 Å per molecule). DPhPE potential reached ̴350 mV at collapse, and DOPhPC peaked at ̴200 mV. DOPhPC consistently showed the smallest potential change, after the initial propagation of potential ( ̴180 Å per molecule), as the monolayer density increased.
Figure 3. Selected surface potential monolayers of DPhPC, DPhPE, and DOPhPC on aqueous subphase. Subphase consisted of 150 mM KCl and 25 mM Tris at ph 7.2 in every measurement (N=5 minimum for each lipid).
As previously described 27, the surface potential-area curves were converted to surface potential-density curves shown in Figure 4. The associated slopes of each curve and the calculated elemental dipole for each lipid are displayed in Table 1. Calculations were done using the Helmholtz equation describing voltage between the plates of a planar capacitor ∆V = 4πρd/ε, where ρ is the charge density, d is distance between the plates and ε is the dielectric 7 ACS Paragon Plus Environment
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constant. Defining the capacitor as an array of dipoles and assuming ε = 1, Gaines28 determined the normal component of the elemental dipole moment in the array as: µ⊥ = A∆V/12π, where µ⊥ is expressed in mD, A in Å2, and ∆V in mV. The assumption of ε = 1 is adequate because here µ⊥ is the effective dipole including all contributions. According experiments16 and MD simulations29, water oriented around polar atoms makes a decisive contribution to the surface dipole at the bilayer-water interface, and for this reason it is logical to consider it as a part of the total dipole, but not as a passive high-dielectric solvent. This view justifies ε = 1 as a framework for atomistic representation of lipid dipoles including structural water with its own dipole. Table 1. Spatial, elastic and electric properties of DPhPC, DPhPE, and DOPhPC derived from pressure-area and surface potential-area traces. A40 and A35 are the molecular areas measured at lateral pressures of 40 and 35 mN/m, respectively. The unitary dipole moment µ⊥ was calculated from linear slopes of V-A-1 curves according to equation ∆V=∆V0+12πµ⊥A-1 (ref 27). Lipid DPhPC DPhPE DOPhPC
A40 (Å2) 81.2 ± 2.6 81.7 ± 0.3 70.4 ± 1.5
A35 (Å2) 84.8 ± 3.0 84.3 ± 0.4 73.6 ± 1.7
Cs-1 (mN/m) 122 ± 7 156 ± 6 114 ± 8
Vo (mV) 95 ± 12 38 ± 14 136 ± 25
V·A-1 (mV/Å-2) 23670 ± 1550 26430 ± 510 6710 ± 260
µ˔ (D) 0.63 ± 0.04 0.70 ± 0.01 0.17 ± 0.04
Figure 4. Change of surface potential with varying density of DPhPC, DPhPE, and DOPhPC derived from potential isotherms (V-A-1 curves). Black lines indicated fitting of linear sections of measured traces for dipole moment calculations. Characterization of liposomes made out of diphytanoyl lipids. We used dynamic light scattering (DLS) to measure size distributions of DPhPC and DOPhPC liposomes prepared via a 1-min sonication (Figure 5A). Ignoring small fraction of aggregates, the size of the liposomes 8 ACS Paragon Plus Environment
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ranged from 30 nm to 44 nm. The average size of DPhPC liposomes was 34.3 ± 2.0 nm. DOPhPC liposomes populations, Figure 5B, showed a slight shift of the distribution towards a larger liposome size profile compared to DPhPC liposomes. DOPhPC liposomes ranged from 35 nm to 76 nm. Average DOPhPC liposomes size was 39.7 ± 5.4 nm. Additional sonication did not change these distributions. Attempts to prepare liposomes from DPhPE by sonication were unsuccessful, apparently, due to strong negative intrinsic curvature.
Figure 5. Example measurements of liposome size of DPhPC (A), and DOPhPC (B). Dynamic Light Scattering was done at room temperature. All measurements N=5.
Liposome Spreading. The kinetics of surface tension change observed upon introduction of DPhPC and DOPhPC liposomes were highly variable. In a typical case, an injection of a 28 µL aliquot of a liposome suspension into the well at room temperature leads to an almost instantaneous drop of surface tension by 10-20 mN/m, followed by a considerably slower decrease to an equilibrium value of 26-27 mN/m over a course of several minutes (Figure 6). As temperature increased, an increase in the rate of spreading and the ‘instantaneous’ fraction of tension drop were generally observed for both lipids. Figure 6A represents DPhPC spreading kinetics showing a decrease of initial surface tension as temperatures increased. At 60 °C, DPhPC lipids almost instantaneously spread to the surface. DOPhPC at 60 °C, as shown in Figure 6, did not exhibit instantaneous spreading to the surface. DOPhPC surface tension after the initial addition of liposomes to the buffer were observed to be lower than DPhPC with the exception of measurements at 60 °C.
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Figure 6. Representative traces of spreading kinetics of DPhPC (A), and DOPhPC (B). Arrows indicate time points when liposomes where injected into the buffer. Figure 7 illustrates the statistics of surface tension changes measured at different temperatures for sonicated DPhPC and DOPhPC liposomes. DPhPC lipids reduced surface tension by 45.9 ± 0.1 mN/m at 20 °C. Spreading at higher temperatures consistently showed surface tension identical to measurements at 20 °C. DOPhPC liposomes decreased surface tension by 48.9 ± 1.0 mN/m at 20 °C. Further increases in temperature did not significantly change the surface tension. Both values were close to the monolayer collapse pressures (Figure 2A).
Figure 7. Surface tension measurements of DPhPC and DOPhPC at varying temperatures. DPhPC values measured as 26.9 ± 0.1 mN/m at 20 °C, 26.9 ± 0.1 mN/m at 40 °C, and 26.8 ± 1.0 mN/m at 60 °C (N=5 for all temperatures). DOPhPC values were 23.9 ± 1.0 mN/m at 20 °C, 25.0 ± 0.8 mN/m at 40 °C, and 24.3 ± 1.4 mN/ m at 60 °C (N= 5 for all temperatures).
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Discussion In nature, phytanoyl (3,7,11,15-tetramethylhexadecyl) chains are found throughout Archaea, specifically in halophiles, methanogens, and thermophiles 30. Phytanoyl chains provide higher stability for the lipid bilayer in a wide range of temperatures as measured by decreased proton permeability of DPhPC versus egg PC31 and decreased permeability to protons, urea, and glycerol compared to E.coli 32, 33. The stablity of diphynatoyl lipids membranes stems from the 4-methylations in the alkyl chains sterically hindering tight interactions with neighboring tails. Due to a greater separation of chains, the mebrane remains liquid and at the same time more odered due to restricted rotational chain dynamics 34. Diphytanoyl lipids found in nature are mostly diether. Ether bonds are more chemically stable and resistant to certain lipases compared to ester bonds 35, adding to archaeal lipid stability. Synthetic DPhPC with ester linkages has been a lipid of choice for formation of artificial bilayers and reconstitution of various membrane proteins reported in more than 600 papers. Yet, there have been only a few systematic studies where the intrincic properties of DPhPC have been compared to its native archaeal diether analog (DOPhPC) or other phospholipids. The scope of the above experiments was the comparison of basic mechanical and electrostatic (diople) properties of these two lipids. The monolayer-bilayer equivalence pressure (πB) is the surface pressure at which the lipids in a monlayer are packed at the same density (area per lipid) as in a bilayer. Other criteria for choosing equivalence pressure may include compressibility modulus, phase transition temperature, or aliphatic chain tilt angle36. Each can produce a different value of πB. At the lateral pressure of 40 mN/m the molecular area of DPhPC is 78.3 ± 5.1 Å2/molecule, whereas at 35 mN/m, an earlier estimation of the monolayer-bilayer equivalence pressure 37, DPhPC molecular area takes 84.4 ± 3.0 Å2/molecule. The former is in good agreement with the bilayer molecular area of 80.5 ± 1.5 Å2/molecule obtained from x-ray and neutron scattering data in liposomes 12, as well as from MD simulations38. This points to 40 mN/m as the πB for this particularly lipid. Generally, the monolayer-bilayer equivalence pressure does not have to be the same for all lipids36, 39. However, it is interesting that this pressure is the same as the value derived from fluorescent measurements reported by Brockman and coworkers for stearoyloleoyl phosphocholine (SOPC) monolayers and liposomes 25. The pressure is ~12% below the typical collapse pressure for DPhPC monolayers (45-47 mN/m). The area of DPhPC molecules at the equivalence pressure exceeds that of a regular non-branched dipalmitoyl (47.4 Å2/molecule)40 or dipalmitoleoyl (65.8 Å2/molecule)41 phosphatidylcholines with similar length of hydrocarbon chains. Branching increases effective in-plane cross-section of the molecule. At this packing density the compressibility modulus (Cs-1) of DPhPC reaches 122 ± 7 mN/m, which is considerably softer than films made of the saturated linear-chain DPPC with a Cs-1 of 294 mN/m42. The unsaturated analog of dioleoylphosphatidylcholine has a Cs-1 of 265 ± 18 mN/m43 which lies between DPhPC and DOPhC. Looking past the tail structures to the replacement of the choline head group with ethanolamine in DPhPE, the effective molecular area (81 Å2/molecule) changed only slightly, 11 ACS Paragon Plus Environment
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but the compressibility modulus increased to 160 mN/m (Table 1). An increased capacity to form a network of hydrogen bonds between the ethanolamine headgroups seems to make the compressed film more rigid. The ether version of the lipid with the PC headgroup (DOPhPC) compacted to 70.4 Å2/molecule at 40 mN/m and it’s compressibility peaked at 115 mN/m, lower than the esther analog. The presence of ester carbonyls increases area per lipid, and measurably increases stiffness (reciprocal compressibility modulus). This can be due to both hydrogen-bonding ability and solvation with extra water 14, but the ‘stiffening’ effect of ester groups at bilayer densities is surprizingly minimal. Keeping in mind that monolayers are halfbilayers, the expected elastic modulus for DOPhPC or DPhPC liposome membranes should be in the range of 220-250 mN/m, i.e. well corresponding to elastic stretch moduli measured in pipette aspiration experiments for liposomes made of C18 unsaturated lipids43. However, MD simulations obviously overestimated the elastic modulus of DPhPC bilayeras reported as 605 ± 40 mN/m38 and 670 mN/m44. The experimental values of expansion modulus for DPhPC monolayers have become important since the use of physical stimulation of mechanosensative channels reconstituted in DIBs24. In this system, tension which gates the channel is proportional to the increase in area of compressed droplets and the elasticity (compressibility) modulus. Although, the air-water interface is not the same as the oil-water interface in DIBs systems, it is a reasonable approximation of the interface.
Regarding lipid electrostatics, the surface potential measured on lipid films at the air-water interface consists of a constant Vo, and a lipid density-dependent term 19. Dipole potential appears at a density corresponding to the formation of an expanded but continuous monolayer 19 and increases as the films gets compressed. Consistent with previous studies 16, 17, our experiments showed a lower diople potential and the dipole moment of the ether analog relative to ester lipid. Ester carbonyls coordinate water molecules differently, which in turn contribute to the lipid dipole. Comparatively, when no carbonyl is present in the lipid the surface potential drops by 30 mV - 200 mV 17. This phenomena is seen through various methods including simulations on diphytanoyl lipids which suggest a membrane dipole potential of 1002 mV for DPhPC and 567 mV for DOPhPC 45 and Cryo-Electron microscopy phase contrast studies which have shown DPhPC membrane dipole potential as 510 mV and 260 mV for DOPhPC46. The overal polarity of the interface is lower in DOPhPC since the estimated partial charges on ether oxygens are -0.445 compared to the combined charges of the carbonyl (-0.63) and bridging ester (-0.31) oxygens in DPhPC47. The DPhPC interface is thus characterized with a stronger water-polarizing ability, which is the major contribution to the interfacial dipole potential16, 29. Although monolayers reperesent only half of the membrane, our data is more consistent with cryo-electron studies, whereas simulations, seem to overestimate the absolute values of potentials and the difference between DOPhPC and DPhPC. Knowledge of dipole potential associated with the lipid interface may predict the character of foreign substance interaction with the bilayer, driving forces for permeation, and will allow engineering of asymmetric bilayers with a permanent intrinsic electrostaic bias. Equilibration between the monolayer and vesicular forms is a critical property of lipids forming lung surfactant 48 and a useful property for formation of artificial bilayers 49. Droplet interface 12 ACS Paragon Plus Environment
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bilayers (DIBs) are usually formed by introducing liposomes to an aqueous phase50. The phase state of lipids is known to strongly affect the spreading capacity 51. We show that both the ether and ester forms of diphytanoyl PC spread to the density that corresponds to the collapse pressure of monolayers, independent of temperature. This is consistent with absence of phase transitions in DPhPC. With this regard, this behavior is almost ideal. Liposomes sonicated under identical conditions show a slight shift of distribution towards larger size for DOPhPC suggesting that membranes made of this lipid might have a slightly higher bending modulus. Spreading kinetics were variable, but there was a general tendency for the equilibration time to shorten with temperature and occur with two-stage kinetics. The lateral pressure almost instantaneously increased by 10-20 mN/m, which was followed by a slower (1-6 min) nonexponential approach to equilibrium pressure. The reason for two-stage behavior is unknown, but it appears that the character of liposome interaction with the bare air-water interface is different from that with partially formed monolayer. Independent of temperature, the final value of surface pressure was ~45 mN/m for DPhPC and between 46 and 48 mN/m for DOPhPC. The obtained lateral pressure was within 4% of collapse pressure for the respective monomolecular films formed from chloroform solutions. This shows that both DPhPC and DOPhPC behave almost ‘ideally’ at the air-water interface reaching the same equilibrium pressure from either side: forming vesicles as a result of buckling of the overcompressed monolayer or forming monolayers from excess lipid presented to the interface in the form of liposomes. Conclusions Here we addressed two basic questions: what are the consequences of branched chains for the mechanics of membranes and how do introduced ester linkages change the properties of archaeal lipids. Methylated chains undoubtedly increase molecular area but, exhibit lateral compressibility/expansion moduli similar to common unsaturated phospholipids. Branching, which removes phase transitions, permits ‘ideal’ spreading in a broad range of temperatures. The presence of ester carbonyls in the synthetic DPhPC form increases molecular area, most likely with aid from hydration. Coordination of water with ester carbonyls doubles the dipole potential. This difference can be utilized in biomolecular engineering to create permanent intrinsic electrical bias inside the folded planar membranes or droplet interface bilayers 22, 50.
Acknowledgements The authors thank Drs. Don Leo and Jeff Klauda for critical reading of the manuscript. This program is funded by the Air Force Office of Scientific Research Basic Research Initiative grant FA9550-12-1-0464.
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References (1) Ulrih, N. P.; Gmajner, D.; Raspor, P. Structural and physicochemical properties of polar lipids from thermophilic archaea. Appl. Microbiol. Biotechnol. 2009, 84, 249-60. (2) Redwood, W. R.; Pfeiffer, F. R.; Weisbach, J. A.; Thompson, T. E. Physical properties of bilayer membranes formed from a synthetic saturated phospholipid in n-decane. Biochim. Biophys. Acta 1971, 233, 1-6. (3) Lindsey, H.; Petersen, N. O.; Chan, S. I. Physicochemical characterization of 1,2-diphytanoyl-snglycero-3-phosphocholine in model membrane systems. Biochim. Biophys. Acta 1979, 555, 147-67. (4) Langecker, M.; Arnaut, V.; Martin, T. G.; List, J.; Renner, S.; Mayer, M.; Dietz, H.; Simmel, F. C. Synthetic Lipid Membrane Channels Formed by Designed DNA Nanostructures. Science 2012, 338, 932-936. (5) Astier, Y.; Braha, O.; Bayley, H. Toward single molecule DNA sequencing: direct identification of ribonucleoside and deoxyribonucleoside 5'-monophosphates by using an engineered protein nanopore equipped with a molecular adapter. J. Am. Chem. Soc. 2006, 128, 1705-10. (6) Oiki, S.; Danho, W.; Montal, M. Channel protein engineering: synthetic 22-mer peptide from the primary structure of the voltage-sensitive sodium channel forms ionic channels in lipid bilayers. Proc. Natl. Acad. Sci. USA 1988, 85, 2393-7. (7) Kitano, T.; Onoue, T.; Yamauchi, K. Archaeal lipids forming a low energy-surface on air-water interface. Chem. Phys. Lipids 2003, 126, 225-32. (8) Mirzabekov, T.; Lin, M. C.; Yuan, W. L.; Marshall, P. J.; Carman, M.; Tomaselli, K.; Lieberburg, I.; Kagan, B. L. Channel formation in planar lipid bilayers by a neurotoxic fragment of the beta-amyloid peptide. Biochem. Biophys. Res. Commun. 1994, 202, 1142-8. (9) Shenoy, D. K.; Barger, W. R.; Singh, A.; Panchal, R. G.; Misakian, M.; Stanford, V. M.; Kasianowicz, J. J. Functional reconstitution of protein ion channels into planar polymerizable phospholipid membranes. Nano Lett. 2005, 5, 1181-5. (10) Gross, L. C.; Heron, A. J.; Baca, S. C.; Wallace, M. I. Determining membrane capacitance by dynamic control of droplet interface bilayer area. Langmuir 2011, 27, 14335-42. (11) Sarles, S. A.; Leo, D. J. Regulated attachment method for reconstituting lipid bilayers of prescribed size within flexible substrates. Anal. Chem. 2010, 82, 959-66. (12) Tristram-Nagle, S.; Kim, D. J.; Akhunzada, N.; Kucerka, N.; Mathai, J. C.; Katsaras, J.; Zeidel, M.; Nagle, J. F. Structure and water permeability of fully hydrated diphytanoylPC. Chem. Phys. Lipids 2010, 163, 630-7. (13) Kucerka, N.; Nagle, J. F.; Sachs, J. N.; Feller, S. E.; Pencer, J.; Jackson, A.; Katsaras, J. Lipid bilayer structure determined by the simultaneous analysis of neutron and X-ray scattering data. Biophys. J. 2008, 95, 2356-67.
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(14) Smaby, J. M.; Hermetter, A.; Schmid, P. C.; Paltauf, F.; Brockman, H. L. Packing of Ether and Ester Phospholipids in Monolayers. Evidence for Hydrogen-Bonded Water at the sn- 1 Acyl Group of Phosphatidylcholines. Biochemistry (Wash.) 1983, 22, 5808-5813. (15) Guler, S. D.; Ghosh, D. D.; Pan, J.; Mathai, J. C.; Zeidel, M. L.; Nagle, J. F.; Tristram-Nagle, S. Effects of ether vs. ester linkage on lipid bilayer structure and water permeability. Chem. Phys. Lipids 2009, 160, 33-44. (16) Gawrisch, K.; Ruston, D.; Zimmerberg, J.; Parsegian, V. A.; Rand, R. P.; Fuller, N. Membrane dipole potentials, hydration forces, and the ordering of water at membrane surfaces. Biophys. J. 1992, 61, 1213-23. (17) Paltauf, F.; Hauser, H.; Phillips, M. C. Monolayer characteristics of some 1,2-diacyl, I-alkyl-2-acyl and 1,2-dialkyl phospholipids at the air-water interface. Biochim. Biophys. Acta 1971, 249, 539-47. (18) Alakoskela, J. I.; Kinnunen, P. K. Control of a redox reaction on lipid bilayer surfaces by membrane dipole potential. Biophys. J. 2001, 80, 294-304. (19) Smaby, J. M.; Brockman, H. L. Surface dipole moments of lipids at the argon-water interface. Similarities among glycerol-ester-based lipids. Biophys. J. 1990, 58, 195-204. (20) Schindler, H. Formation of planar bilayers from artificial or native membrane vesicles. FEBS Lett. 1980, 122, 77-9. (21) Stanley, C. E.; Elvira, K. S.; Niu, X. Z.; Gee, A. D.; Ces, O.; Edel, J. B.; Demello, A. J. A microfluidic approach for high-throughput droplet interface bilayer (DIB) formation. Chem. Commun. (Cambridge, U.K.) 2010, 46, 1620-2. (22) Bayley, H.; Cronin, B.; Heron, A.; Holden, M. A.; Hwang, W. L.; Syeda, R.; Thompson, J.; Wallace, M. Droplet interface bilayers. Mol. Biosyst. 2008, 4, 1191-208. (23) MacDonald, R. C.; Simon, S. A. Lipid monolayer states and their relationships to bilayers. Proc. Natl. Acad. Sci. USA 1987, 84, 4089-93. (24) Najem, J.; Dunlap, M.; Sukharev, S.; Leo, D. J. Mechanosensitive Channels Activity in a Droplet Interface Bilayer System. MRS Online Proc. Libr. 2014, 1621, 171-176. (25) Dahim, M.; Mizuno, N. K.; Li, X. M.; Momsen, W. E.; Momsen, M. M.; Brockman, H. L. Physical and photophysical characterization of a BODIPY phosphatidylcholine as a membrane probe. Biophys. J. 2002, 83, 1511-24. (26) Smaby, J. M.; Momsen, M. M.; Brockman, H. L.; Brown, R. E. Phosphatidylcholine acyl unsaturation modulates the decrease in interfacial elasticity induced by cholesterol. Biophys. J. 1997, 73, 1492505. (27) Brockman, H. Dipole potential of lipid membranes. Chem. Phys. Lipids 1994, 73, 57-79. (28) Gaines, G. L. Insoluble Monolayers At Liquid-Gas Interfaces; John Wiley & Sons Inc.: New York, 1966.
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(29) Zhou, F.; Schulten, K. Molecular Dynamics Study of a Membrane-Water Interface. J. Phys. Chem. 1995, 99, 2194-2207. (30) De Rosa, M.; Gambacorta, A. The lipids of archaebacteria. Prog. Lipid Res. 1988, 27, 153-75. (31) Komatsu, H.; Chong, P. L. Low permeability of liposomal membranes composed of bipolar tetraether lipids from thermoacidophilic archaebacterium Sulfolobus acidocaldarius. Biochemistry 1998, 37, 107-15. (32) Mathai, J. C.; Sprott, G. D.; Zeidel, M. L. Molecular mechanisms of water and solute transport across archaebacterial lipid membranes. J. Biol. Chem. 2001, 276, 27266-71. (33) Yamauchi, K.; Doi, K.; Kinoshita, M.; Kii, F.; Fukuda, H. Archaebacterial lipid models: highly salttolerant membranes from 1,2-diphytanylglycero-3-phosphocholine. Biochim. Biophys. Acta 1992, 1110, 171-7. (34) Koga, Y.; Morii, H. Recent advances in structural research on ether lipids from archaea including comparative and physiological aspects. Biosci., Biotechnol., Biochem. 2005, 69, 2019-34. (35) Choquet, C. G.; Patel, G. B.; Beveridge, T. J.; Sprott, G. D. Stability of pressure-extruded liposomes made from archaeobacterial ether lipids. Appl. Microbiol. Biotechnol. 1994, 42, 375-84. (36) Nagle, J. F.; Tristram-Nagle, S. Structure of lipid bilayers. Biochim. Biophys. Acta 2000, 1469, 159-95. (37) Marsh, D. Lateral pressure in membranes. Biochim. Biophys. Acta 1996, 1286, 183-223. (38) Lim, J. B.; Klauda, J. B. Lipid chain branching at the iso- and anteiso-positions in complex Chlamydia membranes: a molecular dynamics study. Biochim. Biophys. Acta 2011, 1808, 323-31. (39) Kucerka, N.; Nieh, M. P.; Katsaras, J. Fluid phase lipid areas and bilayer thicknesses of commonly used phosphatidylcholines as a function of temperature. Biochim. Biophys. Acta 2011, 1808, 276171. (40) Sun, W. J.; Tristram-Nagle, S.; Suter, R. M.; Nagle, J. F. Structure of gel phase saturated lecithin bilayers: temperature and chain length dependence. Biophys. J. 1996, 71, 885-91. (41) Kucerka, N.; Gallova, J.; Uhrikova, D.; Balgavy, P.; Bulacu, M.; Marrink, S. J.; Katsaras, J. Areas of monounsaturated diacylphosphatidylcholines. Biophys. J. 2009, 97, 1926-32. (42) Barbara Gzyl , M. P. Mixed monolayers of dipalmitoyl phosphatidylcholine and ethyl palmitate at the air/water interface. Appl. Surf. Sci. 2005, 246, 356-361. (43) Rawicz, W.; Olbrich, K. C.; McIntosh, T.; Needham, D.; Evans, E. Effect of chain length and unsaturation on elasticity of lipid bilayers. Biophys. J. 2000, 79, 328-39. (44) Shinoda, W.; Shinoda, K.; Baba, T.; Mikami, M. Molecular dynamics study of bipolar tetraether lipid membranes. Biophys. J. 2005, 89, 3195-202.
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(45) Shinoda, K.; Shinoda, W.; Baba, T.; Mikami, M. Comparative molecular dynamics study of ether- and ester-linked phospholipid bilayers. J. Chem. Phys. 2004, 121, 9648-54. (46) Wang, L.; Bose, P. S.; Sigworth, F. J. Using cryo-EM to measure the dipole potential of a lipid membrane. Proc. Natl. Acad. Sci. USA 2006, 103, 18528-33. (47) 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. (48) Baoukina, S.; Monticelli, L.; Amrein, M.; Tieleman, D. P. The molecular mechanism of monolayerbilayer transformations of lung surfactant from molecular dynamics simulations. Biophys. J. 2007, 93, 3775-82. (49) Nelson, N.; Anholt, R.; Lindstrom, J.; Montal, M. Reconstitution of purified acetylcholine receptors with functional ion channels in planar lipid bilayers. Proc. Natl. Acad. Sci. USA 1980, 77, 3057-61. (50) Sarles, S. A.; Leo, D. J. Physical encapsulation of droplet interface bilayers for durable, portable biomolecular networks. Lab Chip 2010, 10, 710-7. (51) Heurtault, B.; Saulnier, P.; Pech, B.; Proust, J. E.; Benoit, J. P. Physico-chemical stability of colloidal lipid particles. Biomaterials 2003, 24, 4283-300.
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Table of Contents Figure
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Article
Properties of diphytanoyl phospholipids at the air-water interface Anthony Yasmann, and Sergei Sukharev Langmuir, Just Accepted Manuscript • DOI: 10.1021/la503800g • Publication Date (Web): 04 Dec 2014 Downloaded from http://pubs.acs.org on December 10, 2014
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Properties of diphytanoyl phospholipids at the air-water interface Anthony Yasmann and Sergei Sukharev Department of Biology, University of Maryland College Park, MD 20742 Correspondence: [email protected], 301-405-6923
ABSTRACT: Diphytanoyl phosphatidyl choline (DPhPC) is a synthetic ester lipid with methylated tails found in archaeal ether lipids. Due to stability of DPhPC bilayers and absence of phase transitions in a broad range of temperatures, the lipid is used as an artificial membrane matrix for reconstitution of channels, pumps, and membrane-active peptides. We characterized monomolecular films made of DPhPC and its natural ether analog DOPhPC at the air-water interface. We measured compression isotherms and dipole potentials of films made of DPhPC, DPhPE and DOPhPC. We determined that at 40 mN/m the molecular area of DPhPC is 81.2 Å2, consistent with x-ray and neutron scattering data obtained in liposomes. This indicates that 40 mN/m is the monolayer-bilayer equivalence pressure for this lipid. At this packing density, the compressibility modulus (Cs-1= 122 ± 7 mN/m) and interfacial dipole potential (V= 355 ± 16 mV) were near their maximums. The molecular dipole moment was estimated as 0.64 ± 0.02 D. The ether DOPhPC compacted to 70.4 Å2/lipid at 40 mN/m displaying similar peak compressibility to DPhPC. The maximal dipole potential of the ether lipid was about half of that for DPhPC at this density and the elemental dipole moment was about a quarter. Spreading of DPhPC and DOPhPC liposomes reduced surface tension of the aqueous phase by 46 and 49 mN/m, respectively. This corresponds well to the monolayer collapse pressure. The equilibration time shortened as temperature increased from 20 oC to 60 oC, but surface pressure at equilibrium did not change. The data illustrates the properties of branched chains and contributions of ester bonds in setting the mechanical and electrostatic parameters of diphytanoyl lipids. These properties determine an environment in which reconstituted voltage- or mechano-activated proteins may function. Electrostatic properties are important in preparation of asymmetric folded bilayers, whereas lateral compressibility defines tension in mechanically stimulated droplet interface bilayers.
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Introduction Archaeal branched phospholipids are a special adaptation to the extreme environment which thermophilic organisms tolerate 1. The branched phytanoyl hydrocarbon chains remain in a liquid crystalline state in a wide range of temperatures. At the same time, the lipids retain an ordered bilayer structure due to steric crowding and conformational restraints imposed by methyl branches. These chains are resistant to oxidation and attached through ether linkages, chemically more stable than ester bonds, to the glycerol backbone. The stability of ether lipids provide a robust and stable lipid matrix for many archaea. Diphytanoyl phosphocholine (DPhPC) was synthesized for the purpose of creating an oxidation-resistant and stable lipid bilayers for in-vitro experiments with artificial membranes and reconstituted membrane proteins 2. It contains archaeal hydrocarbon chains but ester linkages resembling bacterial and eukaryotic phospholipids. DPhPC has been reported to have no gel to liquid crystalline phase transition from -120 °C to 120 °C3. Methyl branches, evenly spaced along the chains (Figure 1), reduce the density of packing and inter-chain Van-der-Waals interactions, which suppresses phase transitions more than cis-unsaturation kinks in linear hydrocarbon chains. The absence of phase transitions and related instabilities is a desired trait for many practical uses of the lipid; this advantage is reflected by numerous publications using DPhPC as a matrix for reconstitution of channels, peptides, and transporters4, 5, 6. Despite many experimental uses such as preparation of liposomes 7, planar bilayers 8, supported bilayers 9, or droplet interface bilayer (DIBs)10, 11 the properties of DPhPC compared to its ether diphytanoyl DOPhPC analog and nonbranched phospholipids have not been exhaustively studied. Branched phytanoyl chains increase the in-plane cross-section of the molecule, possibly exposing more hydrocarbon to water. The polar carbonyl oxygens of DPhPC, substituting less polar ether oxygens of DOPhPC, reside in that boundary region, which may alter the character of solvation and hydrogen bonding in this interfacial zone. As a result, the mechanics and electrostatics of the interfacial layer may be altered. X-ray and neutron scattering experiments determined that the molecular area of DPhPC in liposomes is near 80 Å2, which is 16 Å2 larger than that of the non-branched dipalmitoylphosphatidylcholine (DPPC) of similar length 12, 13, but no comparison with DOPhPC was reported. There have been several comparative studies of ester and ether phospholipid analogs, showing that ester lipids exhibited an area 7-10 Å2/molecule larger at all surface pressures compared to analogs with one 14 or two ether bonds 15. A number of studies marked a notable decrease in surface potential and dipole moment of pure ether lipids 16, 17, 18 or etherester hybrid phospholipids 19 compared to their ester analogs. Use of diphytanoyl lipids for reconstitution warrants investigation into the equilibrium between their vesicular and monolayer forms. Vesicles open up at the air-water or oil-water interfaces forming monolayers, a property that is widely used to form folded bilayers 20 or DIBs 21, 22. On the other hand, at high lateral pressures monolayers collapse producing vesicles. Whether the same equilibrium between the vesicular and monolayer forms is achieved in both processes is 2 ACS Paragon Plus Environment
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unclear. This pressure resulted from spreading of liposomes made of saturated lipids at the airwater interface was shown to have strong temperature dependence 23. Although it is known that DPhPC monolayers collapse at high lateral pressures, the equilibria between the interfacial and vesicular forms have not been studied. In addition, the compression elasticity modulus of artificial monolayers is becoming an important parameter for tension estimations when droplet interface bilayers (DIBs) are subjected to mechanical distortion in experiments with reconstituted mechanosensitive channels.24 In the present paper, we compare interfacial properties of ester and ether diphytanoyl phosphocholines. We report area per head group at monolayer-bilayer equivalence pressures, compressibility, and dipolar properties of these lipids. We also present data on the liposome spreading kinetics and equilibria, illustrating an almost thermodynamically ‘ideal’ behavior of these lipids.
Figure 1. Chemical structures of DPhPE, DPhPC and its natural ether analog DOPhPC.
Methods and Materials 1,2-diphytanoyl-sn-glycero-3-phosphocholine (DPhPC), 1,2-diphytanoyl-sn-glycero-3phosphoethanolamine (DPhPE), and 1,2-di-O-phytanyl-sn-glycero-3-phosphocholine (DOPhPC) were purchased from Avanti Polar Lipids (Alabaster, AL, USA). Potassium Chloride, Trizma® Hydrochloride, Potassium Hydroxide, and Hexane were acquired from Sigma-Aldridge (St. Louis, MO, USA). All solutions were prepared with ultrapure 18.2 MΩ x cm water from a Barnstead 3 ACS Paragon Plus Environment
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Nanopure system. All monolayer and tensiometry experiments were performed in a dust-free laminar flow hood (AirClean® Systems, Raleigh, NC, USA). Lipid stocks. Each lipid was dried under stream of nitrogen gas in pre-weighed air tight vials. Lipids were placed overnight under deep vacuum (10-20 µmHg) with a solvent trap. DPhPC, DOPhPC, and DPhPE were reconstituted in hexane by weight to a stock concentration. Buffers. A 10X stock buffer was prepared to a concentration of 1.5 M KCl and 250 mM Tris in ultrapure water and titrated to a pH of 7.2 using KOH. The buffer was filtered through a 2 µm bottle top filter (Nalgene™ Rapid-Flow™) and stored in glass. All glassware was acidwashed and thoroughly rinsed. Preparations of Liposomes. Lipids were dried from hexane solutions in round-bottom disposable glass tubes first under stream of nitrogen and then under vacuum (4 hrs), rehydrated for an hour in buffer to a concentration of 6.25 mg/mL. The solution was then vigorously vortexed, and sonicated for a minute at room temperature in a Branson-type sonicator (Aquasonic 50t, VWR, West Chester, PA, USA). Liposomes were then diluted to 5 mg/mL. Pressure-area (π-A) isotherms. Surface tensions of lipids were observed using Wilhelmy method with a filter paper strip (Whatman, No. 1, 10.5 mm wide and 0.25 mm thick) as the plate. The pressure sensor (model 601, NIMA, Coventry, U.K.) was calibrated with a 100 mg weight, after calibration surface tension of pure water was determined to be -72 mN/m. The sensor was zeroed to provide negative measures for surface tension. A rectangular Teflon® monolayer trough (max area of ̴550 cm2) with a single barrier (NIMA) were used in the pressure-area isotherm experiments. The stock solutions of DPhPC, DOPhPC, and DPhPE were diluted by weight to final concentrations of 1.2 mg/mL, 1.2 mg/mL, and 1.4 mg/mL respectively. The 30 µL of lipids were spread onto the subphase using a gastight 50µL Hamilton syringe for all lipids. A 150 mM KCl and 25 mM Tris (pH 7.2) buffer was the subphase in all experiments. After allowing time for solvents to evaporate (15 min), the area compression isotherms were performed at room temperature (22 °C) at a speed of 150 cm2/min. The onset of the pressure change with film compression, the collapse pressure and the lateral compressibility modulus Cs1 = A(dπ/dA) were determined from the curves. Potentials-area (V-A) isotherms. Potentials were measured using potential sensor MicroSpot (Kibron, Helsinki, Finland) in a Kibron MicroTroughXS. A rectangular aluminum trough (max area of ̴104 cm2) with two mobile barriers (Kibron) were used in the potential experiments. The stock solutions of DPhPC, DOPhPC, and DPhPE were diluted by weight to final concentrations of 0.2 mg/mL each. Lipid spreading method and the buffer used were identical to the pressure area isotherm experiments. Area compression of the monolayer was repeated as in π-A isotherm experiments at a speed of 3 mm/min. Spreading Kinetics. Surface spreading kinetics were observed using Wilhelmy method with a Kibron MicroTroughXS sensor and flamed wire (Kibron, proprietary alloy, 0.51 mm diameter) as the Wilhelmy plate in a 2 x 3 array of 200 µL wells custom-made out of Teflon. The tensiometer was calibrated by contrasting voltage measurements of air and ultrapure water as per the Kibron manual. Control measurements of the buffer were taken at 20 °C. Buffer and 4 ACS Paragon Plus Environment
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liposomes were then heated by water bath to each experimental temperature. Buffer was placed into the well and recordings were then infused with liposomes to a final concentration of 1 mg/ml. The wells were cleaned with chloroform between trials. Dynamic Light Scattering. Liposome sizes were measured by dynamic light scattering with a 90Plus Particle Size Analyzer (Brookhaven Instruments Corporation, Holtsville, NY, USA). The analyzer was standardized with a 90 nm calibrating particles (Duke Scientific Corporation, Palo Alto, CA, USA). Liposomes were freshly sonicated and diluted from 6.25 mg/mL to 0.1 mg/mL in subphase buffer. Measurements were performed in polystyrene cuvettes (Brookhaven Instruments Corporation, square, 10mm, 4.5 mL) at room temperature (22 °C).
Results Pressure-area (π-A) isotherms. Compression isotherms for lipid monolayers formed from DPhPC, DOPhPC, and DPhPE are shown in Figure 2A. DPhPC and DPhPE begin showing changes in surface pressure at molecular area around 140 Å2 while DOPhPC change is observed near 130 Å2. The collapse pressure of DPhPC and DOPhPC films was observed as roughly 45 mN/m with DPhPE collapsing around 47 mN/m. At 40 mN/m considered to be the monolayer-bilayer equivalence point25 the area per molecule is 78.3 Å2 ± 5.1 Å2 (n=7 for DPhPC, 70.4 Å2 ± 1.5 Å2 (n=7) for DOPhPC, and 81.3 Å2 ± 0.3 Å2 (n=7) for DPhPE. Compressibility modulus. Isotherms of each lipid were converted to compressibility modulus curves (Figure 2B) as described in 26. DPhPE is the stiffest lipid of the group having the highest compressibility modulus near 160 mN/m. DOPhPC indicates tighter packing than both DPhPC and DPhPE with a left shift of the compression modulus curve and a lower peak of about 115 mN/m. DPhPC exhibited compressibility modulus peaking near 125 mN/m. Areas per lipid and compressibility moduli for the three lipids are presented together with electrostatic parameters in Table 1.
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Figure 2. Monolayer pressure-area (π-A) isotherms of DPhPC, DPhPE, and DOPhPC (A). Monolayer-bilayer equivalence seen at 78.3 Å2 ± 5.1 Å2 for DPhPC, 70.4 Å2 ± 1.5 Å2 for DOPhPC, and 81.3 Å2 ± 0.3 Å2 for DPhPE (n=7 for each lipid). Area compressibility modulus (B) over molecular area calculated from pressure-area (π-A) isotherms Cs-1 = A(dπ/dA) for DPhPC, DPhPE, and DOPhPC.
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Surface Potential and its area dependence. We consider the measured surface (volta) potential as purely dipole under the assumption that all lipids studied here are zwitterionic near physiological pH with no net charge, and that possible specific adsorption of ions at the interface are small. Figure 3 shows a group of surface potential –area curves as the lipid monolayers were compressed. The DPhPC curve displays the typical initial potential buildup to a plateau as area per molecule decreases, followed by a gradual increase to the second plateau near the collapse point. DOPhPC and DPhPE show a similar but, less defined behavior. DPhPC and DPhPE showed a similar surface potential profile during monolayer compression, with DPhPE surface potential curve being shifted to the left. DPhPC initial potential buildup appears at ̴220 Å per molecule and DPhPE potential build up is seen at ̴180 Å per molecule. DPhPC and DPhPE show a gradual change in potential as the lipid density increases that is greater than that of DOPhPC. DPhPC consistently had the largest change of potential during the monolayer compression peaking at ̴390 mV near the point of collapse ( ̴80 Å per molecule). DPhPE potential reached ̴350 mV at collapse, and DOPhPC peaked at ̴200 mV. DOPhPC consistently showed the smallest potential change, after the initial propagation of potential ( ̴180 Å per molecule), as the monolayer density increased.
Figure 3. Selected surface potential monolayers of DPhPC, DPhPE, and DOPhPC on aqueous subphase. Subphase consisted of 150 mM KCl and 25 mM Tris at ph 7.2 in every measurement (N=5 minimum for each lipid).
As previously described 27, the surface potential-area curves were converted to surface potential-density curves shown in Figure 4. The associated slopes of each curve and the calculated elemental dipole for each lipid are displayed in Table 1. Calculations were done using the Helmholtz equation describing voltage between the plates of a planar capacitor ∆V = 4πρd/ε, where ρ is the charge density, d is distance between the plates and ε is the dielectric 7 ACS Paragon Plus Environment
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constant. Defining the capacitor as an array of dipoles and assuming ε = 1, Gaines28 determined the normal component of the elemental dipole moment in the array as: µ⊥ = A∆V/12π, where µ⊥ is expressed in mD, A in Å2, and ∆V in mV. The assumption of ε = 1 is adequate because here µ⊥ is the effective dipole including all contributions. According experiments16 and MD simulations29, water oriented around polar atoms makes a decisive contribution to the surface dipole at the bilayer-water interface, and for this reason it is logical to consider it as a part of the total dipole, but not as a passive high-dielectric solvent. This view justifies ε = 1 as a framework for atomistic representation of lipid dipoles including structural water with its own dipole. Table 1. Spatial, elastic and electric properties of DPhPC, DPhPE, and DOPhPC derived from pressure-area and surface potential-area traces. A40 and A35 are the molecular areas measured at lateral pressures of 40 and 35 mN/m, respectively. The unitary dipole moment µ⊥ was calculated from linear slopes of V-A-1 curves according to equation ∆V=∆V0+12πµ⊥A-1 (ref 27). Lipid DPhPC DPhPE DOPhPC
A40 (Å2) 81.2 ± 2.6 81.7 ± 0.3 70.4 ± 1.5
A35 (Å2) 84.8 ± 3.0 84.3 ± 0.4 73.6 ± 1.7
Cs-1 (mN/m) 122 ± 7 156 ± 6 114 ± 8
Vo (mV) 95 ± 12 38 ± 14 136 ± 25
V·A-1 (mV/Å-2) 23670 ± 1550 26430 ± 510 6710 ± 260
µ˔ (D) 0.63 ± 0.04 0.70 ± 0.01 0.17 ± 0.04
Figure 4. Change of surface potential with varying density of DPhPC, DPhPE, and DOPhPC derived from potential isotherms (V-A-1 curves). Black lines indicated fitting of linear sections of measured traces for dipole moment calculations. Characterization of liposomes made out of diphytanoyl lipids. We used dynamic light scattering (DLS) to measure size distributions of DPhPC and DOPhPC liposomes prepared via a 1-min sonication (Figure 5A). Ignoring small fraction of aggregates, the size of the liposomes 8 ACS Paragon Plus Environment
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ranged from 30 nm to 44 nm. The average size of DPhPC liposomes was 34.3 ± 2.0 nm. DOPhPC liposomes populations, Figure 5B, showed a slight shift of the distribution towards a larger liposome size profile compared to DPhPC liposomes. DOPhPC liposomes ranged from 35 nm to 76 nm. Average DOPhPC liposomes size was 39.7 ± 5.4 nm. Additional sonication did not change these distributions. Attempts to prepare liposomes from DPhPE by sonication were unsuccessful, apparently, due to strong negative intrinsic curvature.
Figure 5. Example measurements of liposome size of DPhPC (A), and DOPhPC (B). Dynamic Light Scattering was done at room temperature. All measurements N=5.
Liposome Spreading. The kinetics of surface tension change observed upon introduction of DPhPC and DOPhPC liposomes were highly variable. In a typical case, an injection of a 28 µL aliquot of a liposome suspension into the well at room temperature leads to an almost instantaneous drop of surface tension by 10-20 mN/m, followed by a considerably slower decrease to an equilibrium value of 26-27 mN/m over a course of several minutes (Figure 6). As temperature increased, an increase in the rate of spreading and the ‘instantaneous’ fraction of tension drop were generally observed for both lipids. Figure 6A represents DPhPC spreading kinetics showing a decrease of initial surface tension as temperatures increased. At 60 °C, DPhPC lipids almost instantaneously spread to the surface. DOPhPC at 60 °C, as shown in Figure 6, did not exhibit instantaneous spreading to the surface. DOPhPC surface tension after the initial addition of liposomes to the buffer were observed to be lower than DPhPC with the exception of measurements at 60 °C.
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Figure 6. Representative traces of spreading kinetics of DPhPC (A), and DOPhPC (B). Arrows indicate time points when liposomes where injected into the buffer. Figure 7 illustrates the statistics of surface tension changes measured at different temperatures for sonicated DPhPC and DOPhPC liposomes. DPhPC lipids reduced surface tension by 45.9 ± 0.1 mN/m at 20 °C. Spreading at higher temperatures consistently showed surface tension identical to measurements at 20 °C. DOPhPC liposomes decreased surface tension by 48.9 ± 1.0 mN/m at 20 °C. Further increases in temperature did not significantly change the surface tension. Both values were close to the monolayer collapse pressures (Figure 2A).
Figure 7. Surface tension measurements of DPhPC and DOPhPC at varying temperatures. DPhPC values measured as 26.9 ± 0.1 mN/m at 20 °C, 26.9 ± 0.1 mN/m at 40 °C, and 26.8 ± 1.0 mN/m at 60 °C (N=5 for all temperatures). DOPhPC values were 23.9 ± 1.0 mN/m at 20 °C, 25.0 ± 0.8 mN/m at 40 °C, and 24.3 ± 1.4 mN/ m at 60 °C (N= 5 for all temperatures).
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Discussion In nature, phytanoyl (3,7,11,15-tetramethylhexadecyl) chains are found throughout Archaea, specifically in halophiles, methanogens, and thermophiles 30. Phytanoyl chains provide higher stability for the lipid bilayer in a wide range of temperatures as measured by decreased proton permeability of DPhPC versus egg PC31 and decreased permeability to protons, urea, and glycerol compared to E.coli 32, 33. The stablity of diphynatoyl lipids membranes stems from the 4-methylations in the alkyl chains sterically hindering tight interactions with neighboring tails. Due to a greater separation of chains, the mebrane remains liquid and at the same time more odered due to restricted rotational chain dynamics 34. Diphytanoyl lipids found in nature are mostly diether. Ether bonds are more chemically stable and resistant to certain lipases compared to ester bonds 35, adding to archaeal lipid stability. Synthetic DPhPC with ester linkages has been a lipid of choice for formation of artificial bilayers and reconstitution of various membrane proteins reported in more than 600 papers. Yet, there have been only a few systematic studies where the intrincic properties of DPhPC have been compared to its native archaeal diether analog (DOPhPC) or other phospholipids. The scope of the above experiments was the comparison of basic mechanical and electrostatic (diople) properties of these two lipids. The monolayer-bilayer equivalence pressure (πB) is the surface pressure at which the lipids in a monlayer are packed at the same density (area per lipid) as in a bilayer. Other criteria for choosing equivalence pressure may include compressibility modulus, phase transition temperature, or aliphatic chain tilt angle36. Each can produce a different value of πB. At the lateral pressure of 40 mN/m the molecular area of DPhPC is 78.3 ± 5.1 Å2/molecule, whereas at 35 mN/m, an earlier estimation of the monolayer-bilayer equivalence pressure 37, DPhPC molecular area takes 84.4 ± 3.0 Å2/molecule. The former is in good agreement with the bilayer molecular area of 80.5 ± 1.5 Å2/molecule obtained from x-ray and neutron scattering data in liposomes 12, as well as from MD simulations38. This points to 40 mN/m as the πB for this particularly lipid. Generally, the monolayer-bilayer equivalence pressure does not have to be the same for all lipids36, 39. However, it is interesting that this pressure is the same as the value derived from fluorescent measurements reported by Brockman and coworkers for stearoyloleoyl phosphocholine (SOPC) monolayers and liposomes 25. The pressure is ~12% below the typical collapse pressure for DPhPC monolayers (45-47 mN/m). The area of DPhPC molecules at the equivalence pressure exceeds that of a regular non-branched dipalmitoyl (47.4 Å2/molecule)40 or dipalmitoleoyl (65.8 Å2/molecule)41 phosphatidylcholines with similar length of hydrocarbon chains. Branching increases effective in-plane cross-section of the molecule. At this packing density the compressibility modulus (Cs-1) of DPhPC reaches 122 ± 7 mN/m, which is considerably softer than films made of the saturated linear-chain DPPC with a Cs-1 of 294 mN/m42. The unsaturated analog of dioleoylphosphatidylcholine has a Cs-1 of 265 ± 18 mN/m43 which lies between DPhPC and DOPhC. Looking past the tail structures to the replacement of the choline head group with ethanolamine in DPhPE, the effective molecular area (81 Å2/molecule) changed only slightly, 11 ACS Paragon Plus Environment
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but the compressibility modulus increased to 160 mN/m (Table 1). An increased capacity to form a network of hydrogen bonds between the ethanolamine headgroups seems to make the compressed film more rigid. The ether version of the lipid with the PC headgroup (DOPhPC) compacted to 70.4 Å2/molecule at 40 mN/m and it’s compressibility peaked at 115 mN/m, lower than the esther analog. The presence of ester carbonyls increases area per lipid, and measurably increases stiffness (reciprocal compressibility modulus). This can be due to both hydrogen-bonding ability and solvation with extra water 14, but the ‘stiffening’ effect of ester groups at bilayer densities is surprizingly minimal. Keeping in mind that monolayers are halfbilayers, the expected elastic modulus for DOPhPC or DPhPC liposome membranes should be in the range of 220-250 mN/m, i.e. well corresponding to elastic stretch moduli measured in pipette aspiration experiments for liposomes made of C18 unsaturated lipids43. However, MD simulations obviously overestimated the elastic modulus of DPhPC bilayeras reported as 605 ± 40 mN/m38 and 670 mN/m44. The experimental values of expansion modulus for DPhPC monolayers have become important since the use of physical stimulation of mechanosensative channels reconstituted in DIBs24. In this system, tension which gates the channel is proportional to the increase in area of compressed droplets and the elasticity (compressibility) modulus. Although, the air-water interface is not the same as the oil-water interface in DIBs systems, it is a reasonable approximation of the interface.
Regarding lipid electrostatics, the surface potential measured on lipid films at the air-water interface consists of a constant Vo, and a lipid density-dependent term 19. Dipole potential appears at a density corresponding to the formation of an expanded but continuous monolayer 19 and increases as the films gets compressed. Consistent with previous studies 16, 17, our experiments showed a lower diople potential and the dipole moment of the ether analog relative to ester lipid. Ester carbonyls coordinate water molecules differently, which in turn contribute to the lipid dipole. Comparatively, when no carbonyl is present in the lipid the surface potential drops by 30 mV - 200 mV 17. This phenomena is seen through various methods including simulations on diphytanoyl lipids which suggest a membrane dipole potential of 1002 mV for DPhPC and 567 mV for DOPhPC 45 and Cryo-Electron microscopy phase contrast studies which have shown DPhPC membrane dipole potential as 510 mV and 260 mV for DOPhPC46. The overal polarity of the interface is lower in DOPhPC since the estimated partial charges on ether oxygens are -0.445 compared to the combined charges of the carbonyl (-0.63) and bridging ester (-0.31) oxygens in DPhPC47. The DPhPC interface is thus characterized with a stronger water-polarizing ability, which is the major contribution to the interfacial dipole potential16, 29. Although monolayers reperesent only half of the membrane, our data is more consistent with cryo-electron studies, whereas simulations, seem to overestimate the absolute values of potentials and the difference between DOPhPC and DPhPC. Knowledge of dipole potential associated with the lipid interface may predict the character of foreign substance interaction with the bilayer, driving forces for permeation, and will allow engineering of asymmetric bilayers with a permanent intrinsic electrostaic bias. Equilibration between the monolayer and vesicular forms is a critical property of lipids forming lung surfactant 48 and a useful property for formation of artificial bilayers 49. Droplet interface 12 ACS Paragon Plus Environment
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bilayers (DIBs) are usually formed by introducing liposomes to an aqueous phase50. The phase state of lipids is known to strongly affect the spreading capacity 51. We show that both the ether and ester forms of diphytanoyl PC spread to the density that corresponds to the collapse pressure of monolayers, independent of temperature. This is consistent with absence of phase transitions in DPhPC. With this regard, this behavior is almost ideal. Liposomes sonicated under identical conditions show a slight shift of distribution towards larger size for DOPhPC suggesting that membranes made of this lipid might have a slightly higher bending modulus. Spreading kinetics were variable, but there was a general tendency for the equilibration time to shorten with temperature and occur with two-stage kinetics. The lateral pressure almost instantaneously increased by 10-20 mN/m, which was followed by a slower (1-6 min) nonexponential approach to equilibrium pressure. The reason for two-stage behavior is unknown, but it appears that the character of liposome interaction with the bare air-water interface is different from that with partially formed monolayer. Independent of temperature, the final value of surface pressure was ~45 mN/m for DPhPC and between 46 and 48 mN/m for DOPhPC. The obtained lateral pressure was within 4% of collapse pressure for the respective monomolecular films formed from chloroform solutions. This shows that both DPhPC and DOPhPC behave almost ‘ideally’ at the air-water interface reaching the same equilibrium pressure from either side: forming vesicles as a result of buckling of the overcompressed monolayer or forming monolayers from excess lipid presented to the interface in the form of liposomes. Conclusions Here we addressed two basic questions: what are the consequences of branched chains for the mechanics of membranes and how do introduced ester linkages change the properties of archaeal lipids. Methylated chains undoubtedly increase molecular area but, exhibit lateral compressibility/expansion moduli similar to common unsaturated phospholipids. Branching, which removes phase transitions, permits ‘ideal’ spreading in a broad range of temperatures. The presence of ester carbonyls in the synthetic DPhPC form increases molecular area, most likely with aid from hydration. Coordination of water with ester carbonyls doubles the dipole potential. This difference can be utilized in biomolecular engineering to create permanent intrinsic electrical bias inside the folded planar membranes or droplet interface bilayers 22, 50.
Acknowledgements The authors thank Drs. Don Leo and Jeff Klauda for critical reading of the manuscript. This program is funded by the Air Force Office of Scientific Research Basic Research Initiative grant FA9550-12-1-0464.
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Table of Contents Figure
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