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Colloids and Surfaces B: Biointerfaces 82 (2011) 550–561

Contents lists available at ScienceDirect

Colloids and Surfaces B: Biointerfaces

journa l homepage: www.e lsev ier .com/ locate /co lsur fb

esicle and bilayer formation of diphytanoylphosphatidylcholine (DPhPC) andiphytanoylphosphatidylethanolamine (DPhPE) mixtures and their bilayers’lectrical stability

artin Anderssona,b,∗, Joshua Jackmana, Danyell Wilsona, Patrik Jarvollb,iveka Alfredssonc, George Okeyoa, Randolph Durana

Department of Chemistry, University of Florida, Gainesville, FL 32611, USADepartment of Chemical and Biological Engineering, Chalmers University of Technology, SE-412 96 Göteborg, SwedenPhysical Chemistry, Center of Chemistry and Chemical Engineering, Lund University, SE-221 00 Lund, Sweden

r t i c l e i n f o

rticle history:eceived 15 June 2010eceived in revised form8 September 2010ccepted 11 October 2010vailable online 15 October 2010

eywords:PhPCPhPEesiclesipid bilayeresicle fusionCM-DMR diffusionFMryo-TEMLS

a b s t r a c t

Lipid bilayers are of interest in applications where a cell membrane mimicking environment is desired.The performance of the lipid bilayer is largely dependent on the physical and chemical properties of thecomponent lipids. Lipid bilayers consisting of phytanoyl lipids have proven to be appropriate choicessince they exhibit high mechanical and chemical stability. In addition, such bilayers have high electricalresistances. Two different phytanoyl lipids, 1,2-diphytanoyl-sn-glycero-3-phosphocholine (DPhPC) and1,2-diphytanoyl-sn-glycero-3-phosphoethanolamine (DPhPE), and various combinations of the two havebeen investigated with respect to their behavior in aqueous solutions, their interactions with solid sur-faces, and their electrical stability. Dynamic light scattering, nuclear magnetic resonance diffusion, andcryogenic transmission electron microscopy measurements showed that pure DPhPC as well as mixturesof DPhPC and DPhPE consisting of greater than 50% (mol%) DPhPC formed unilamellar vesicles. If thetotal lipid concentration was greater than 0.15 g/l, then the vesicles formed solid-supported bilayers onplasma-treated gold and silica surfaces by the process of spontaneous vesicle adsorption and rupture, asdetermined by quartz crystal microbalance with dissipation monitoring and atomic force microscopy. Thesolid-supported bilayers exhibited a high degree of viscoelasticity, probably an effect of relatively highamounts of imbibed water or incomplete vesicle fusion. Lipid compositions consisting of greater than50% DPhPE formed small flower-like vesicular structures along with discrete liquid crystalline structures,

ip-diplectrophysiologylectroporation

as evidenced by cryogenic transmission electron microscopy. Furthermore, electrophysiology measure-ments were performed on bilayers using the tip-dip methodology and the bilayers’ capacity to retain itselectrical resistance towards an applied potential across the bilayer was evaluated as a function of lipidcomposition. It was shown that the lipid ratio significantly affected the bilayer’s electrical stability, withpure DPhPE having the highest stability followed by 3DPhPC:7DPhPE and 7DPhPC:3DPhPE in decreasingorder. The bilayer consisting of 5DPhPC:5DPhPE had the lowest stability towards the applied electrical

potential.

. Introduction

Lipid bilayers are popular tools to mimic the cell membrane.

hey have been used in a wide range of applications such as mem-rane protein studies [1–3], biosensors [4–7], and pharmaceuticalrug discovery [8–10]. As a platform for electrophysiology studies,he lipid bilayers can be used either as a thin barrier separating

∗ Corresponding author at: Department of Chemical and Biological Engineering,halmers University of Technology, SE-412 96 Göteborg, Sweden. Tel.: +46 7722966;

ax: +46 31160062.E-mail address: martina@chalmers.se (M. Andersson).

927-7765/$ – see front matter © 2010 Elsevier B.V. All rights reserved.oi:10.1016/j.colsurfb.2010.10.017

© 2010 Elsevier B.V. All rights reserved.

two aqueous solutions across a small aperture [11] or supportedon a solid substrate, which provides increased mechanical stabil-ity [4]. In particular, supported tethered bilayers are useful forthe design of biosensors, as demonstrated earlier by our researchgroup [12–14]. Important considerations for using lipid bilayers asplatforms for electrophysiology measurements based on ion chan-nel conductance are membrane durability in terms of mechanicalstress and chemical stability. As well, high electrical resistance

across the bilayer is necessary because the signal associated withion channel conductance is in the picoampere range. One classof lipids having suitable properties for such measurements is thephytanoyl lipids. These lipids are naturally found in the cellularmembranes of extremophiles such as archaebacteria [15], which

M. Andersson et al. / Colloids and Surfaces

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ig. 1. The chemical structure of the two lipids investigated in this study, 1,2-iphytanoyl-sn-glycero-3-phosphoethanolamine (DPhPE) and 1,2-diphytanoyl-sn-lycero-3-phosphocholine (DPhPC).

re adapted to harsh environments, and are phase stable over aide range of temperatures (−120 to 120 ◦C) [16].

In this present paper, the focus has been on the behavior of twohytanoyl lipids, 1,2-diphytanoyl-sn-glycero-3-phosphocholineDPhPC) and 1,2-diphytanoyl-sn-glycero-3-phosphoethanolamineDPhPE), as well as specific mixtures of them. The chemical struc-ures of the two lipids are presented in Fig. 1. Their head groups,holine and ethanolamine, are two of the most abundant found inammalian cell membranes [17]. Particular emphasis was directed

owards the lipids’ aqueous behavior and their electrical stabilityt the liquid-air interface, as well as how their structures inter-ct with solid substrates, specifically the environmental conditionsnd lipid compositions that favor solid-supported bilayer forma-ion. Pure DPhPC, as well as DPhPC mixed with DPhPE in a 7:3

olar ratio, have previously been shown to form vesicles, whichpon fusion to modified self-assembled monolayer (SAM) surfacesormed tethered bilayers, creating a suitable environment for fullyunctional ion channels [18–20]. This 7:3 lipid mixture has beenarticularly useful for these sensing applications since it has beenemonstrated that it is possible to measure single ion channel activ-

ty using such tethered bilayers [12–14]. Furthermore, it has beenhown that bilayers of this particular composition can be furthertabilized using surface-layer proteins (S-layer proteins). M2ı ionhannels were functionally incorporated into such S-layer stabi-ized membranes and single ion channel recordings for as long as0 h were achieved [21]. Despite these promising results in terms ofeasuring single ion channel activity and stability over time, there

s a lack of fundamental studies on mixtures of these lipids, howhey form bilayers, both at the liquid–air interface and on solid sur-aces, and their bilayers’ electrical stabilities. In 2001, Gauger et al.ublished a paper comparing the hydration of DPhPC and DPhPE

ipids in order to understand their phase transition behavior, usingTIR spectroscopy [22]. They drew several conclusions: the lipids’ranched chains are extremely disordered, resulting in highly fluidembrane structures, and that DPhPC lipids have more hydrody-

amically coupled water than DPhPE lipids. To our knowledge, thiss the only published report that systematically evaluates these twoipids.

This present paper in part examines how the lipid compositionf DPhPC and DPhPE affects vesicle formation at room temperature.ynamic light scattering (DLS), diffusion nuclear magnetic reso-ance (NMR-D), and cryogenic transmission electron microscopycryo-TEM) were used to characterize the structures formed inqueous solution. Each technique revealed insight into the lipidtructure’s physical properties including size, phase, and diffusiv-ty. The lipid compositions that formed vesicles in aqueous solution

ere further investigated. The interactions of these vesicles witholid surfaces, specifically plasma-treated gold and silica, werexamined using quartz crystal microbalance with dissipation mon-toring (QCM-D). The effects of lipid concentration on the type ofnteractions (vesicle adsorption vs. fusion) that occurred were mon-

B: Biointerfaces 82 (2011) 550–561 551

itored. Atomic force microscopy (AFM) was used to visually analyzein liquid the resulting, solid-supported lipid structures (vesiclesor lipid bilayer). The images obtained corresponded well with theQCM-D data regarding vesicle fusion and vesicle adsorption. Fur-thermore, electrophysiology measurements were performed onbilayers formed at the liquid-air interface by the tip-dip method andthe bilayers’ capacity to retain their electrical resistance towards anapplied potential across the bilayers’ was evaluated as a functionof the lipid ratio.

2. Experimental

2.1. Materials

Archaea phospholipids, 1,2-diphytanoyl-sn-glycero-3-phosphocholine (DPhPC) and 1,2-diphytanoyl-sn-glycero-3-phosphoethanolamine (DPhPE), purity > 99%, were used aspurchased from Avanti Polar Lipids (Alabaster, AL). Buffer solu-tions were prepared with 5 mM 3-morpholinopropanesulfonicacid, MOPS, (>99.5%, ultra grade, from Fluka), 250 mM potassiumchloride (ACS certified grade), and 0.1 mM calcium chloride (ACScertified grade). The buffer was titrated to pH 7.4 using potassiumhydroxide (ACS certified grade). All of the salts were purchasedfrom Fisher Scientific and were used as received. Milli-Q filteredwater (>18 M� cm) was used for all sample preparations andstudies.

2.2. Vesicle preparation

The vesicles were prepared by mixing lipid solutions (2 mg/ml inchloroform) of DPhPC and DPhPE lipids in the desired molar ratios.The chloroform was evaporated under vacuum and the lipid mix-ture was hydrated in water or buffer at the desired concentrationand heated at 50 ◦C under vigorous stirring until a clear solutionwas obtained (ca. 1 h). After allowing the suspension to cool toroom temperature, it was sonicated in a bath sonicator for 5 min,followed by extrusion through an 80 nm polycarbonate membrane(11 times) using a mini-extruder from Avanti Polar Lipids. Extrusionwas performed no longer than 24 h prior to any investigation.

2.3. Gold surfaces

For the AFM investigations, ultra smooth gold surfaces were pre-pared on silicon wafers by evaporation of Ti (3 nm) followed by a60% Au/40% Pd alloy (50 nm). A 20 nm gold layer was depositedusing evaporation on top of the alloy. To clean the surface, it wasrinsed with hexane, acetone, ethanol, and water, respectively, thendried with a stream of nitrogen air and subsequently treated withUV/ozone for 20 min.

2.4. Cryo-TEM

Cryo-TEM was performed on a Phillips CM120 BioTWIN cryoelectron microscope operated at 120 kV using an Oxford CT3500cryo-holder. Specimens were kept in the microscope and imaged ata temperature of about 93.5 K using liquid nitrogen cooling. Imageswere recorded digitally with a CCD camera (Gatan MSC791), andunder focus conditions to improve the phase contrast. Specimensfor electron microscopy were prepared in a controlled environ-ment vitrification system (CEVS) [23] to ensure a fixed temperature(26–28 ◦C) and high humidity in order to minimize evaporation.

In brief, a drop of sample (0.2 g lipids/l buffer) was put on a glowdischarge pretreated Pelco grid (lacy carbon film, supported by acopper grid). Then, excess solution was removed by blotting withfilter paper, leaving a thin meniscus of the solution in the holes ofthe carbon film. The blotting was done first on the side opposite

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o the sample drop and then gently on the same side as the placedample. The grid was then rapidly plunged into liquid ethane at itselting temperature. The vitrified specimens were stored under

iquid nitrogen.

.5. DLS

DLS measurements were performed on a Precision DetectorsDDLS/CoolBatch+90T instrument and the data were analyzedsing the Precision Deconvolve32 Program. The measurementsere performed using a 683 nm laser source and a 90◦ scattering

ngle, set at 20 ◦C. When necessary, additional dilutions were per-ormed until the count rate was between the recommended 200nd 400 k counts per second. However, all presented DLS resultsave been obtained using 0.2 g/l. CONTIN analysis was used foralculating the size distributions [24,25].

.6. NMR diffusion

The NMR diffusion experiments were performed on a Bruker50 MHz (17.6 T) equipped with a diffusion probe with 24 T/mctively shielded gradients (Diff60). A Bruker 10 mm diffusionrobe was used but with 5 mm NMR tubes. All NMR experimentsere performed at 25 ◦C using a lipid concentration of 0.2 g/l.

timulated Echo with Pulsed Field Gradients (PFG-STE) was used.iffusion time (�) of 100 ms and gradient time (ı) of 1.1 ms weresed and the gradient strength (g) was arrayed in 32 steps with6 scans in each step. To monitor the diffusion of the vesicles, amall amount (less than 1% of the lipids) of hexamethyl-disiloxaneHMDS) was added. HMDS is essentially insoluble in water, so theharacteristic HMDS signal (0 ppm) will arise from HMDS that isissolved into the lipid membrane. The echo-decays obtained fromhe vesicles gave curved plots in a Log Y-axis of signal intensity vs., k = �2g2ı2(� − ı/3), where � is the gyromagnetic constant. Echo-ecays were fitted to a Log Normal distributed function, which cane described by:

(D) = 1

D�ln√

2�exp

(− [ln(D) − ln(Dm)]2

2�2ln

)(1)

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D〉 = Dm exp

(�2

ln2

)(2)

nd the normal standard deviation, �D, of the diffusion coefficientsith

2D = 〈D〉2

(exp(�2

ln) − 1)

(3)

The diameters of the vesicles were obtained through thetokes–Einstein relation [26].

.7. QCM-D

The QCM-D measurements were performed on a QCM-Z500rom KSV Instruments (Helsinki, Finland) equipped with a temper-ture control unit from Oven Instruments. AT-cut crystals, coatedith either an evaporated silica or gold layer, had a resonance fre-

uency of 5 MHz and were also purchased from KSV. Prior to theeasurements, the crystals were cleaned either using piranha solu-

ion for the gold-coated crystals (3:1 concentrated sulphuric acid to0% hydrogen peroxide solution) or an SDS solution for the silica-oated crystals, followed by rinsing with water and ethanol, and

B: Biointerfaces 82 (2011) 550–561

drying with a gentle stream of nitrogen gas. All rinse cleaning pro-cedures were followed by UV-oxygen plasma cleaning for 20 min.This sequence of pre-treatment led to clean and oxidized surfacesthat had contact angles of less than 6◦. The instrument was cali-brated using the standard procedure as described by the instrumentprovider [19]. The fluids were introduced batch-wise via a reservoir,and then proceeded to a bypass chamber (500 �l volume), situatedabove the crystal. After loading the instrument with the crystal,it was allowed to stabilize in the used solvent for approximately30 min until a stable baseline was obtained. The measurementswere recorded at the fundamental frequency, the 3rd overtone, andthe 5th overtone. The measured resonant frequency, f, depends onthe oscillating mass of the crystal including any adsorbed species.For rigid films there is a relation between the change in frequency�f and the adsorbed mass, m, according to the Sauerbrey equation

�m = −C

n�f (4)

where C is the mass sensitivity constant (17.7 ng cm−2 Hz−1) atf = 5 MHz and n is the overtone number. The dissipation factor, D, isrelated to the energy lost (Elost) to the surroundings in relation tothe stored energy (Estored) upon oscillation and is defined as

D = Elost

2�Estored�f = R

ωL= ωRC (5)

where ω is the angular frequency, R is the resistance, L is the induc-tance, and C is the capacitance of the crystal.

2.8. AFM

AFM analyses were performed in buffer solution using aNanoscope III MultiMode Scope (Veeco) equipped with a 13 �mE-scanner. The instrument was calibrated in the z-direction usinga silicon grating (TGZ01, Mikromash), with a step height of 20 nm(accuracy 1 nm). A fluid cell was used, which was cleaned in ethanoland water prior to use. The cell was used without o-ring. Imageswere obtained using silicon cantilevers (Nanosensors, Neuchatel,Switzerland with dimensions: T = 3.8–4.5 �m, W = 26–27 �m, andL = 128 �m). The cantilevers were mounted in the fluid cell and adrop of the solvent solution was placed on the tip to ensure wetting.The solution was placed on the sample surface, followed by attach-ment of the fluid cell. No addition or removal of liquid through thehoses that are attached to the sample holder was performed. Priorto the measurements, the sample was left undisturbed for ∼20 minto let the solution equilibrate. AFM analyses were performed byscanning in contact mode and the images were processed using asecond-order parameter flattening.

2.9. Electrophysiology

Electrophysiology investigations were performed using thetip-dip methodology, by which a bilayer is formed at the tip ofa microelectrode [27]. First, a lipid monolayer was formed byspreading 15 �l of a 5 g/l lipid chloroform solution using a Hamil-ton syringe on the surface of the buffer solution in a round Teflonbath having a diameter of 0.8 cm. Based on the bath’s surface areaand the cross sectional area of DPhPC, the amount of used lipid wasin orders of magnitude in excess compared to the number of lipidsnecessary to form a monolayer [28]. The lipids were allowed toself-assemble into a monolayer for 10 min prior to the lowering ofa buffer-filled glass patch pipette into the solution. The pipette was

withdrawn from the solution followed by re-lowering, forming abilayer at the tip of the pipette. The patch pipettes were producedusing a P97 Flaming/Brown Micropipette Puller from Sutter Instru-ment Company (Novato, CA). The borosilicate glass capillary tubesused were obtained from Warner Instruments Inc. (Hamden, CT).

rfaces

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M. Andersson et al. / Colloids and Su

he obtained pipette tips had an approximate diameter of 1 �m, aseasured using scanning electron microscopy (SEM). Ag/AgCl elec-

rodes were used; prior to use, they were submerged in industrialtrength Chlorox for 30 min at the beginning of each day before use,

ig. 2. TEM micrographs of samples composed of varying lipid composition: (A) pure DPhite squares in images (D) and (E) were Fast Fourier Transformed (FFT), resulting in thell scale bars correspond to 100 nm.

B: Biointerfaces 82 (2011) 550–561 553

and then rinsed with water and ethanol. Patch clamp recordingswere performed using an Axon Patch-Clamp 200B amplifier fromMolecular Devices Corporation (Union City, CA). The signals werepassed through a low-pass 5 kHz filter and digitized at a sampling

hPC, (B) 7DPhPC:3DPhPE, (C, D, E) 5DPhPC:5DPhPE, and (F) 3DPhPC:7DPhPE. TheFFT patterns inserted in the respective image. The lipid concentration was 0.2 g/l.

5 rfaces B: Biointerfaces 82 (2011) 550–561

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300

350

400

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Vesicle composition DPhPC:DPhPE

NMR (Dm) DLS (Intensity) NMR ( ⟨D⟩) DLS (Number)

Fig. 3. Average effective diameter as a function of lipid ratio. The sizes wereobtained using DLS and NMR-diffusion and the lipid concentration was 0.2 g/l. Forthe results obtained by NMR-D, the standard deviation of the logarithmical distribu-

54 M. Andersson et al. / Colloids and Su

ate of 20 kHz using a Digidata 1322A digitizer from Molecularevices Corporation. The presented unitary conductances are

he mean values obtained from Gaussian fits of the measuredonductances. Opening and closing dwell time histograms, plotteds logarithmic binned data, were fitted by a sum of exponentialrobability density functions using maximum likelihood [29].

. Results

.1. Cryo-TEM

To visualize the lipid structures, cryo-TEM was utilized. In Fig. 2,ypical micrographs of the lipid structures are presented for theifferent lipid compositions. In the sample containing pure DPhPC,nilamellar vesicles were observed, which are shown in Fig. 2A.he mean diameter of these vesicles was 92 nm (n = 8) and theyere often found adhered to each other. In Fig. 2B, a micrograph of

esicles composed of 7DPhPC:3DPhPE is presented. These vesiclesere also unilamellar, though slightly smaller in size compared to

he pure DPhPC, with a diameter of 60 nm (n = 7). Micrographs of theample consisting of 5DPhPC:5DPhPE are shown in Fig. 2C–E. Theample contained both unilamellar vesicles (C) and more denselyacked structures, which appear to be composed of close packed

ipid bilayers, likely discrete lamellar liquid crystalline phasesD and E). The repeat distance was calculated from the Fourierransform of selected parts of the image (Fig. 2E) and was foundo be around 6.5 nm, which corresponds to the thickness of theipid bilayer together with the separation distance between theilayers. However, the specific phase of the liquid crystal cannote determined solely from the micrographs. In Fig. 2F, a micro-raph obtained from the 3DPhPC:7DPhPE sample is presented.n these samples, small flower-like, vesicles were observed andoth the larger unilamellar vesicles and the densely packed lipidilayer aggregates were absent. For the sample consisting of purePhPE, no objects could be observed with the cryo-TEM tech-ique.

.2. DLS and NMR diffusion

DLS and NMR diffusion were utilized to investigate the size of theipid structures in aqueous solution. For DLS, the time-dependentuctuation in the scattering intensity is observed. The scattering

ntensity is highly dependent on the size of the scattering entityhich can cause larger objects to be overrepresented in the mea-

ured size distribution of a polydisperse sample. However, thisverrepresentation can be corrected for by using a weighting func-ion that transforms the mean intensity value to the mean numberalue. In Fig. 3, both the intensity data and the adjusted data areresented as the sample’s average effective diameter vs. the lipidomposition. The calculated diameters of the lipid aggregates’ arendependent of the lipid compositions ranging from pure DPhPCo 30% DPhPE (additional data not included). The samples’ aver-ge effective diameters were approximately 100 nm (mean valueeighted by the total number of object). Increasing DPhPE’s rel-

tive molar concentration from 30% to 70%, the sample diameterinearly increased to approximately 200 nm. The pure DPhPE sam-le’s diameter was somewhat smaller. In Fig. 3, the aggregate sizesbtained by NMR diffusion using Dm and 〈D 〉 extracted from Eq. (1)re also presented. The sizes measured by the NMR diffusion tech-ique follow the same trend as the sizes measured by DLS. For the

ure DPhPC and 7DPhPC:3DPhPE samples, smaller diameters werebtained by NMR diffusion in comparison to the results obtainedy DLS. For the sample composed of 5DPhPC:5DPhPE, the twoechniques gave sizes that were in good agreement. For higher con-entrations of DPhPE, including pure DPhPE, the sample diameters

tion, �log, were 0.71 (pure DPhPC), 1.17 (7DPhPC:3DPhPE), 0.91 (5DPhPC:5DPhPE),0.72 (3DPhPC:7DPhPE) and 0.74 (pure DPhPE). The lines in the figure are only aguide to the eye.

obtained by NMR diffusion were larger than the number-averagedDLS sizes.

3.3. QCM-D

Fig. 4 shows the measured change in frequency and dissipationas a function of time that was recorded when two different concen-trations of pure DPhPC vesicles interacted with a silica substrate.When the lipid concentration was relatively low, 0.1 g/l, a decreasein the measured frequency together with an increase in dissipa-tion over the course of time was observed (A). The correspondingfrequency shift indicates that the vesicles adsorbed on the silicasurface, forming a highly viscoelastic film as evidenced by the highenergy dissipation. When the lipid concentration was increasedto 0.15 g/l, the frequency change is initially similar as for lowerconcentrations, but after ∼1150 s the frequency starts to increase,indicating a decrease in adsorbed mass, followed by frequency sta-bilization (B). This adsorption behavior is typical of spontaneousvesicle adsorption and rupture, where vesicles first adsorb on thesurface until reaching a critical concentration and then ruptureto form a solid-supported lipid bilayer [30]. When an adsorbedvesicle ruptures, hydrodynamically coupled solvent in the vesicle’sinterior is released, resulting in an increased frequency (decreasedmass) and a decreased dissipation (lower degree of viscoelasticity).The total change in frequency was ∼37 Hz, which corresponds toan adsorbed mass of 655 ng/cm2 (according to Eq. (4)). Assumingthat a bilayer is formed, based on characteristic two-step vesiclefusion kinetics, this results in a surface area per molecule of 43 A2

(Mw = 846 D). However, this area is smaller than 77 A2, which hasbeen previously reported for DPhPC [31].

Fig. 5 shows the measured change in frequency and dissipationas a function of time that was recorded when various concen-trations of pure DPhPC vesicles at various lipid concentrationsinteracted with a plasma-treated gold substrate. When the lipidconcentration was low, 0.02 g/l, a decrease in the measured fre-

quency together with an increase in dissipation over the course oftime was observed (A). The corresponding frequency shift indicatesthat vesicles adsorbed on the gold surface. When the lipid con-centration was increased to 0.1 g/l, a more pronounced frequencydecrease and dissipation increase as a function of time were

M. Andersson et al. / Colloids and Surfaces B: Biointerfaces 82 (2011) 550–561 555

-250

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-150

-100

-50

0A B

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-60

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ig. 4. Changes in resonant frequency and dissipation versus time for the adsorptiohe presented QCM-D data was recorded at the fifth overtone and renormalized su

bserved (B). This indicates a higher concentration of adsorbedesicles and a higher degree of film viscoelasticity (likely due tohe increased vesicle surface coverage and corresponding increasen hydrodynamically coupled solvent) in comparison to the 0.02 g/l

-220

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ig. 5. Changes in resonant frequency and dissipation versus time for the adsorption of p.1 g/l, and (C) 0.15 g/l. The presented QCM-D data was recorded at the fifth overtone andn an identical scale.

Time (S)

re DPhPC vesicles onto silica with lipid concentrations of (A) 0.1 g/l and (B) 0.15 g/l.t �f = �fn = 5/5.

sample. In Fig. 5C, the measured data obtained from a 0.15 g/lsample is shown. Here, the frequency decreases initially followedby a frequency increase after ∼1100 s, indicating a decrease inadsorbed mass. The total change in frequency was ∼59 Hz, which

60004000 6000400020000

1 g/l 0.15 g/l

Time(s)e(s)

C

ure DPhPC vesicles onto oxidized gold with lipid concentrations of (A) 0.02 g/l, (B)renormalized such that �f = �fn = 5/5. For each concentration, the data are plotted

556 M. Andersson et al. / Colloids and Surfaces B: Biointerfaces 82 (2011) 550–561

F and sw

c(tms

cfidteffvbrcpttDw

Fu

ig. 6. (A) AFM of the gold surface and (B) the same surface after addition of vesiclesere obtained in contact mode using a liquid cell.

orresponds to an adsorbed mass of 1044 ng/cm2 (according to Eq.4)). Assuming that a bilayer is formed, based on the characteristicwo-step spontaneous vesicle adsorption and rupture kinetics, this

ass results in a surface area per molecule of 27 A2, which is evenmaller than the one obtained using the silica surface.

It is important to note that the Sauerbrey equation used to cal-ulate this molecular surface areas is only valid for non-viscoelasticlms, which is not the case in this situation. This can be seen in theissipation curves (Figs. 4B and 5C), which shows that the dissipa-ion is relatively high even after the bilayer has formed, indicatingither a highly viscoelastic bilayer due to the unique structuraleatures of phytanoyl lipids (see Section 4) or incomplete bilayerormation such that intact, adsorbed vesicles coexist with rupturedesicles. The presented QCM-D experiments were all performed inuffer at pH 7.4. Experiments were also done in pure water, whichesulted in no vesicle adsorption or fusion independent of lipid con-entration. Furthermore, additional QCM-D measurements wereerformed on all of the aforementioned lipid compositions. Mix-

ures that had DPhPE lipid concentrations of less than 50% showedhe same behavior as for pure DPhPC. However, with increasingPhPE concentration, no adsorption, independent of composition,as observed.

ig. 7. AFM images of pure DPhPC vesicles deposited on an ultra flat gold surface. The lipsing a liquid cell.

ubsequent fusion, leading to solid-supported bilayer formation. The measurements

3.4. AFM

AFM was performed on ultra smooth gold surfaces in the samebuffer solution as used in the QCM-D measurements. In Fig. 6A,an image of a pure gold surface is shown. As seen, the surface issmooth even on the nanometer length scale. In Fig. 6B, the samesurface is shown 2 h after a 0.2 g/l DPhPC vesicle solution was added.The surface is still smooth, however much less sharper imageswere obtained. This decreased sharpness is most probably due tothe presence of a lipid bilayer on the gold surface, which wouldresult in a viscoelastic layer that can be disturbed by the scanningprobe.

Fig. 7 shows AFM images, both 2D and 3D renderings, as well ascross-section analysis. These images were obtained 2 h after addi-tion of a 0.1 g/l vesicle solution. In the bottom left image, the surfaceis viewed from above and there are round objects that have diam-eters between 100 and 250 nm, which is within expected rangefor adsorbed vesicles. Due to substrate–vesicle interactions, the

adsorbed vesicles are no longer spherical but rather pancake-like,supporting the larger measured diameters of these entities. These3D features can be seen in the right figure, where a topographicalimage of the surface is presented. On the upper left, a cross section

id concentration was ∼0.1 g/l. The measurements were performed in contact mode

rfaces B: Biointerfaces 82 (2011) 550–561 557

od

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M. Andersson et al. / Colloids and Su

f the lower left figure is presented together with some relevantistances.

.5. Electrophysiology

Bilayers composed of five different lipid ratios of DPhPC andPhPE were examined using the tip-dip technique. The experi-ents began by obtaining a gigaseal corresponding to the electrical

esistance of the bilayer being in the G� range, followed bypplying potentials between ±200 mV. Recordings obtained fromach evaluated lipid ratio are presented as current measured asfunction of applied potential at a lipid concentration of 5 g/l in

hloroform. The purpose of the measurements was to examine thelectrical stability of the formed bilayer and its ability to with-tand current leakage and pore formation, which relates to randomransitions in current amplitude. As a consequence, the presentedecordings were obtained at a potential where instability in theilayers was observed. For pure DPhPC lipid bilayers, gigaseals wereormed at a success rate of 56% (n = 22) and the bilayer remainedtable for potentials up to 80 mV. At higher potentials, pore for-ation occurred. The pore formation was heterogeneous as seen

n Fig. 8, which was obtained at an applied potential of 120 mV.he lifetimes of the bilayers were usually between 2 and 10 min.nce electroporation occurred, it continued at both high and lowotentials. In addition, the baseline was unstable and would driftetween 1 and 3 pA during the experiments. Quantitatively shortpiky events lasting less than two seconds appeared throughouthe lifetime of the bilayer. It was concluded that pores were formedndependent of the applied potential, and that large and small pores

ith conductances of 0.4–5 pS were formed randomly.For the bilayer consisting of pure DPhPE, the success rate for

igaseal formation was 75% (n = 15). The threshold potential for

ore formation was 120 mV with electroporation observed 36% ofhe time (n = 6). Pore formation was consistent and more uniform asompared to pores formed with pure DPhPC bilayers. The baselinef the bilayer remained stable, having a high recovery rate, withlectroporation conductance decreasing with decreasing potential

ig. 9. A electrophysiology recording obtained from a pure DPhPE bilayer as a function ohe pore threshold at 200 mV first was applied.

Fig. 10. A electrophysiology recording obtained from a bilayer cons

Fig. 8. A electrophysiology recording obtained from a bilayer consisting of pureDPhPC lipids at an applied potential of 120 mV.

as seen in Fig. 9. There was also more electroporation activity atnegative potentials for this lipid composition, despite the bufferbeing symmetrical (results not shown). Bilayer lifetimes were aslong as 30 min to 1 h before electrical breakdown occurred duringwhich time the seal strength decreased from the giga-ohm to eitherthe mega- or kilo-ohm range.

Results from both 3DPhPC:7DPhPE and 7DPhPC:3DPhPE weresimilar and representative recordings of bilayers having these com-positions are presented in Fig. 10. Gigaseals having resistancesbetween 2 and 10 G� were formed at a success rate of 75% for bothratios (n = 19). Threshold potentials for electroporation were above120 mV for both mixtures. Pores formed with 7DPhPC:3DPhPEwere random with current changes ranging from 0 pA to 5 pAand they possessed a drifting baseline, a behavior resembling thatof pure DPhPC bilayers. Pores formed with the 3DPhPC:7DPhPElipid composition remained uniform and a decrease in conduc-

tance with decreasing applied potentials was observed. Bilayerlifetimes were also relatively long for both the 3DPhPC:7DPhPEand 7DPhPC:3DPhPE lipid mixtures, lasting on the order of severalminutes to 1 h.

f increased applied potential, 100–180 mV. The recording was obtained after that

isting of a 7DPhPC:3DPhPE lipid mixture obtained at 200 mV.

558 M. Andersson et al. / Colloids and Surfaces B: Biointerfaces 82 (2011) 550–561

Fig. 11. Current trace obtained from a 5DPhPC:5DPhPE bilayer s

Fig. 12. Typical I/V curve for a 7DPhPC:3DPhPE lipid ratio. The I/V relationshipwas determined by applying a series of voltages, each for the duration of 2.5 s. Allpea

ubpbac

7ts

4

l1wasDrpvttt∼opI

oxygen and nitrogen contacts due to the presence of the methyl

oints on the IV curve were the average of n ≥ 3 different experiments. Standardrror bars were plotted; if bars are not readily visible, then the standard error waspproximately the height of the dot or less.

Bilayers composed of 5DPhPC:5DPhPE were exceptionallynstable. Formation of gigaseals with this ratio was rare and theaseline was unstable, even without any applied potential. Electro-oration that did occur was closely followed by disruption of theilayer. The bilayers were short-lived, with lifetimes of less thanminute. Fig. 11 shows instability of the bilayer at 40 mV with

urrent fluctuations between 0 and 5 pA.In Fig. 12, a current vs. potential plot is shown for a

DPhPC:3DPhPE bilayer. As can be seen, no linearity exists betweenhe current and potential; hence, the bilayer’s resistance is not con-tant with applied potential.

. Discussion

In the present study, structures consisting of two phytanoylipids, 1,2-diphytanoyl-sn-glycero-3-phosphocholine (DPhPC) and,2-diphytanoyl-sn-glycero-3-phosphoethanolamine (DPhPE),ere examined. From the results obtained by Cryo-TEM, DLS

nd NMR diffusion, it was determined that lipid compositions ofingle-component DPhPC and mixtures containing up to ∼50%PhPE form unilamellar vesicles, having sizes that are directly

elated, though not identical, to the membrane pore size of theolycarbonate membrane that was used for extrusion (∼100 nmesicle diameters vs. 80 nm membrane pore size). However,he measured vesicle sizes varied depending on the analyticalechnique used. For example, in the case of pure DPhPC vesicles,he diameters obtained were 122 nm (DLS), 43 nm (NMR-D), and

92 nm (Cryo-TEM). These differences in size are characteristicf using these different techniques and are due to the differenthysical properties that are being measured with each technique.

n the cases of DLS and NMR-D, the hydrodynamic radius is

howing electroporation. The applied potential was 40 mV.

measured since they measure the free diffusion of particles. Eventhough the techniques gave rise to different sizes for the samesample, the sizes follow an identical trend when the lipid ratiowas varied. Cryo-TEM is a more direct method to measure vesiclesize. The technique, however, suffers from artifacts arising fromthe blotting process, which induces shearing on the sample. Thisshearing might affect the size and structure of the observed objects[32]. The specimen preparation technique might also result inthat objects are forced closer to each other resulting in that theyappear adhered to each other. This could for example be thereason for why the pure DPhPC vesicles seem to be adhered toeach other, even though DLS and NMR-D results show that thevesicles diffuse totally separate from one another. The Cryo-TEMtechnique, as in all microscopy, also suffers from poor statisticsdue to the low number of observed objects. At higher DPhPEconcentrations (>50%), both DLS and NMR revealed that the sizeof the measured objects increased together with an increasedsize distribution, indicating a change in the obtained structures.This was further confirmed by cryo-TEM. In solutions containing5DPhPC:5DPhPE, several new structures appeared in addition tothe unilamellar vesicles. These new structures were composedof lipid bilayers, which were less solvated in comparison to thevesicles. For the 3DPhPC:7DPhPE lipid composition, only smallflower-like objects were observed, although much larger objectswere expected to be seen based on the results from the DLS andNMR-D measurements. Likely, this is due to the cryo-TEM samplepreparation process during which these larger objects may havebeen removed. These larger structures are likely similar to thedispersed liquid crystalline structures that were observed in the5DPhPC:5DPhPE sample. The observed flower-like objects are verysimilar to those presented by Gustavsson et al. [33] In their workthe coexistence of flower-like, or as they mention it, conglomeratesof partially fused vesicles and cubosomes, which are dispersedparticles having a cubic liquid crystalline structure, was observed.These results strongly suggest that amphiphiles having a curvaturepromoting reversed structures might result in such flower-likeobjects. That no unilamellar vesicles might be present in the sam-ples containing predominantly DPhPE, could also be the reasonsfor why our QCM-D results (data not shown) showed no vesicleadsorption with these lipid compositions. The large effect of thelipids’ propensity to form or not form vesicles might at first seem abit non-intuitive, since the only difference between the two lipidsis a relatively small variation of the hydrophilic part, as seen inFig. 1. However, this difference will significantly affect the criticalpacking parameter, resulting in a change of the lipid’s spontaneouscurvature. Also, the electrostatic energies are significantly differentbetween the two head groups. The amine head group forms astrong electrostatic interaction and hydrogen bonds between thephosphate oxygens and nitrogen in the ammonium group. Thecholine head group on the other hand does not allow for such close

groups. It is instead water molecules that are closely incorporatedlinking the phosphate groups [34]. As a consequence, the crystalto liquid crystalline transition temperature is lower for lipidswith choline head-groups as compared to the amine head-groups.

M. Andersson et al. / Colloids and Surfaces B: Biointerfaces 82 (2011) 550–561 559

Table 1

Shtt

acsfiobccvIhlTvdAt[pecbrcodDtbsmdcmchvttwtlvtifcbfelDvc

Lipid mixture DPhPC 7DPhPC:3DPhPE

Success rate (%) 56 75Threshold (mV) 80 >120

tudies performed on non-phytanoyl lipids that have the sameead-groups as investigated here, choline and ethanolamine, showhat a lamellar to reverse hexagonal phase transition occurs whenhe mixture contains more than 60% DOPE [35,36].

The QCM-D technique is an excellent tool to study vesicledsorption and vesicle fusion since it reveals binding kinetics, cal-ulates adsorbed film mass including hydrodynamically coupledolvent, and provides viscoelastic information about the adsorbedlm. In comparison to QCM-D studies performed on vesicles madef straight chained lipids, phytanoyl lipids show slightly differentehavior, especially when analyzing the dissipation. Usually, in thease of lipid bilayer formation, the dissipation increases when vesi-les first adsorb to the surface but then decreases to near zero asesicles rupture and the non-viscoelastic lipid bilayer forms [30].n the case of phytanoyl lipids, the dissipation remains relativelyigh throughout the entire process, which has been observed ear-

ier for these types of lipids, but on tethered supports [19,37].his result indicates that the bilayer formed has relatively highiscoelastic character as compared to other bilayers, though theissipation is still much less than that of an adsorbed vesicle film.lso, the final change in frequency after the vesicle fusion is larger

han what has been obtained for DHPE, 37 Hz compared to 26 Hz30]. A possible explanation for this finding could be due to thereviously discussed high viscoelasticity of phytanoyl lipid bilay-rs, caused by the relatively high amounts of hydrodynamicallyoupled solvent, together with incomplete bilayer formation. Theranched chain structure of the lipids’ hydrophobic tails leads toeduction of the conformational and wobbling motions of the alkylhains but, in contrast, the head groups experience a higher degreef freedom and as a consequence, more solvent will be hydro-ynamically coupled around the lipid head groups, especially forPhPC [22]. This increased bound water content together with

he decreased packing of the hydrophobic tails will increase theilayer’s viscoelasticity, which could partly explain the QCM-D dis-ipation response. Despite the fact that phytanoyl membranes haveore coupled solvent, excess chemical potential calculations have

emonstrated that the lipid chain branching does not significantlyhange the partition of water through the hydrophobic part of theembrane [38]. Also, the alkyl branching inhibits the formation of

avities in the membrane, which could explain why the membranesave higher electrical resistances and low ion conduction [38]. Theiscoelasticity of the formed bilayer could also be an explanation ofhe observations made using the AFM. When the lipid concentra-ion was high enough to result in lipid bilayer formation, no sharp,ell-resolved AFM micrographs could be obtained. This is some-

hing which is typical for AFM investigations on low surfaces havingow viscosity [39]. The QCM-D results showed that spontaneousesicle adsorption and rupture was dependent on the lipid concen-ration and the presence of ions, since no adsorption was observedn pure water. Even though the vesicle concentration is sufficient forull surface coverage, the concentration may not be large enough toause spontaneous vesicle adsorption and rupture, which results inilayer formation. An excess of vesicles is needed to induce bilayerormation, indicating that the interaction between the vesicles is

ssential. This phenomenon has been observed earlier for otheripid vesicle systems [40]. The AFM studies also confirm the QCM-

results such that a relatively low lipid concentration results inesicle adsorption and that there is a threshold in the lipid con-entration when a bilayer starts to form at approximately 0.15 g/l.

5DPhPC:5DPhPE 3DPhPC: 7DPhPE DPhPE

<10 75 750 >120 120

The fact that the vesicles fused independently on both tested sur-faces, plasma-treated silica and plasma-treated gold, is somewhatsurprising. Vesicle fusion resulting in bilayer formation is oftenobserved on silica surfaces but rarely on pure gold [30]. It is believedthat the used plasma treatment, which reduced both of the surfaces’contact angles to ∼6◦, played a major role in causing vesicle fusionto occur on the gold surface. In order to evaluate the possibility thatthe plasma treatment induced surface contaminations such as, forexample, silica originating from the interior of the plasma chamber,X-ray photoelectron spectroscopy (XPS) was used. The XPS results(data not shown) showed that no other elements except gold, oxy-gen and carbon were present; hence, no contamination due to theplasma treatment was observed that might have explained howvesicle fusion could occur on a gold surface.

Analysis of the AFM results and the resulting cross sectional cal-culations presented in Fig. 7 indicate that the adsorbed vesiclesappear to be flattened out and deformed. This vesicle deformationhas been observed earlier as a result of vesicle–surface interactionsand depends on both the surface and vesicle composition, amongmany other factors [41].

From the electrophysiology measurements it was shown thatpure DPhPC, pure DPhPE, and mixtures of them formed lipidbilayers, when the tip-dip method was utilized. The membranesstability, however, when the electrical field was applied, varieddepending on the lipid composition. The success rate of the formedgigaseals and the threshold potential for electroporation for all theinvestigated lipid mixtures are presented in Table 1. At relativelylow applied potentials, for example 80 mV for pure DPhPC, unitarycurrents in the picoampere started to appear, that is, the mem-brane opened up allowing for ions to pass through the membrane.Such trans-membrane transport due to applied potential is oftenreferred to as electroporation, a phenomenon often observed atpotentials between 0.2 V and 1 V and for potential pulses appliedfor shorter periods of time. Even though the theory of electropo-ration is still under development, several models exist explaininggreat parts of the process [42]. It is the water molecules along withthe natural dielectric potential of the bilayer that creates an overallbilayer dipole potential which becomes polarized when an externalpotential is applied [42,43]. Using molecular dynamic simulationsto atomically map this process, Tieleman et al. found that poreformation during electroporation is driven by local electric fieldgradients at the water/lipid interface due to this polarization [44].The movement of the water molecules due to the electric fieldgradient increases the probability of water “defects” penetratinginto the bilayer [44,45]. These defects are the molecular basis ofelectroporation, which occurs in the pico-nanosecond time range.Once inside the bilayer, water molecules hydrogen bond withother water molecules already present within the hydrophobicpart forming what is now termed “water fingers” that span thebilayer forming “water pipes” [42,44,45]. Both the lipid chainsand the head-groups are then exposed to water resulting in adeformation of the head-groups whereby aligning the interface ofthe pores, which results in a change in capacitance or conductancethat is observed as a current peak. When applying these theories

to the results obtained in this study, certain characteristics of thedifferent lipid mixtures can be better understood. As mentionedabove, one important aspect that is greatly affecting the stability ofthe lipid bilayer, and the formation of pores, is the hydration stateof the membrane. In this respect, the lipid head-group properties

5 rfaces

sitgmhmtlmaolarhirrocs

tidtfaraatcifF5tb

lObnsmmstbrOonttvtlmaptlt

[[[

[

60 M. Andersson et al. / Colloids and Su

uch as size, polarity, charge, and hydrophobicity have a strongnfluence since they determine the exposure of the lipids towardshe bulk water phase surrounding the bilayer [43,45]. The cholineroup of the DPhPC head-group is hydrophobic due to the threeethyl groups attached to the nitrogen. As a consequence, the PC

ead-group is pulled inwards towards the hydrophobic part of theembrane. This exposes the phosphate group and its charges to

he aqueous phase allowing 18–20 water molecules to bind peripid in the liquid state [34,43]. This is in strong contrast to the

ore hydrophilic amine group present on the DPhPE, which formsmore compact layer above the carbonyl hindering the mobility

f the phosphate backbone and its exposure to water. This amineinkage via hydrogen bonding to adjacent phosphates produces

compact, rigid head-group network at the bilayer surface withoughly 4–6 water molecules bonded per lipid [43]. The relativelyigh amount of water retained by the DPhPC lipid could theoret-

cally be the explanation for the observed unstable baseline andandom pore formation, which is seen in Fig. 8. With the sameeasoning, the uniformity and relatively high potential thresholds,ver 120 mV, for bilayers where DPhPE is the most abundant lipidould be explained by considering the lower hydration state, themaller size, and hydrophobicity of the PE head-group.

Experiments showed that the 5DPhPC:5DPhPE lipid composi-ion proved to be extremely unstable. The explanation for thiss still unknown, but possible raft formation resulting in phaseomains of non-fully integrated lipids is a possibility. This forma-ion occurs when different lipids do not fully integrate and insteadorm isolated islands leading to the instability of the bilayer undern applied electric field. Wong et al., used the coarse-gained Mar-ink model to study mixtures of dipalmitoyl phosphatidylcholinend ethanolamine DPPC and DPPE and found that DPPE lipids havehigher density at the center of the bilayer [46]. They also found

hat areas enriched with DPPC bilayers were slightly thicker butould not fully explain this phenomenon. Even though this theorys not yet fully understood, it is however, a possible explanationor the unstable bilayers that were formed with 5DPhPC:5DPhPE.urthermore, the fact that the cryo-TEM results showed that theDPhPC:5DPhPE lipid ratio contained at least two totally differentypes of aggregates supports the above conclusion that the formedilayer is heterogeneous.

The electrophysiology results clearly show that the phytanoylipid membranes have a surprisingly low electroporation threshold.ne should be aware that these observations are highly influencedy the fact that the tip-dip methodology was used. In other pla-ar bilayer methods, the bilayer is either painted on a small holeeparating two buffers containing compartments or formed on aoveable aperture at the air/water interface. Using the paintingethod, diphytanoyl lipids have been studied where the results

hows threshold potentials of around 250 mV [47]. Also, as men-ioned earlier, 7DPhPC:3DPhPE lipid mixtures forming tetheredilayers have successfully been utilized. In this tethered configu-ation, the electroporation threshold was higher than 400 mV [13].ne property inherent with the tip-dip technique is the presencef a meniscus at the pipette tip giving rise to a curvature that isot present in the other used techniques. This curvature might behe reason for why the electroporation threshold is dependent onhe applied potential direction, as observed here. A similar obser-ation, that is, a difference in membrane behavior dependent onhe sign of the potential, was also observed in a study where S-ayer proteins were used to increase the life times of phytanoyl lipid

embranes [21]. A change in curvature has been shown to strongly

ffect the electroporation threshold with increased electroporationrobability upon increased curvature [48]. Another effect due tohe curvature is that the composition between the two lipid mono-ayers might differ. This is something that has been observed inhe case of smaller vesicles where a mixture containing PC and PE

[

[

[

B: Biointerfaces 82 (2011) 550–561

based lipids resulted in a higher concentration of PE based lipids inthe inner vesicle leaflet [49].

5. Conclusions

In this paper, it was demonstrated that the phytanoyl lipids,1,2-diphytanoyl-sn-glycero-3-phosphocholine (DPhPC) and 1,2-diphytanoyl-sn-glycero-3-phosphoethanolamine (DPhPE), can beused to form supported lipid bilayers on hydrophilic surfaces aswell as at the liquid–air interface. DLS, NMR diffusion, and cryo-TEM showed that pure DPhPC and mixtures of DPhPC and DPhPEcontaining less than 50% of DPhPE formed unilamellar vesicles.These vesicles have the ability to fuse on hydrophilic surfaces ofboth plasma-treated gold and silica, forming solid-supported lipidbilayers as monitored by QCM-D and AFM. The bilayer formation,however, was concentration dependent and only initiated whenthe lipid concentration was above 0.15 g/l and only in buffer solu-tion. The formed lipid bilayers had high viscoelasticities, based onthe QCM-D dissipation measurement. This viscoelasticity is prob-ably because of that phytanoyl lipids’ ability to imbibe relativelyhigh amounts of water or due to incomplete vesicle rupture andbilayer formation. No supported bilayers could be formed fromsolutions containing more than 50% DPhPE. Electrophysiology mea-surements performed using the tip-dip method showed that theelectroporation threshold was surprisingly low, especially for thebilayers consisting of pure DPhPC lipids. Results obtained from3DPhPC:7DPhPE and 7DPhPC:3DPhPE were similar and had thresh-old potentials for electroporation above 120 mV. Gigaseals havingresistances between 2 and 10 G� were formed for both of theseratios. Bilayers composed of 5DPhPC:5DPhPE were exceptionallyunstable. Taken as a whole, the results demonstrate the ability toform lipid bilayers of certain compositions of DPhPC and DPhPE atthe solid–liquid and liquid–air interfaces.

Acknowledgements

S. Blackband at the Advanced Magnetic Resonance Imagingand Spectroscopy (AMRIS) facility at the University of Florida isacknowledged for providing NMR access and for general assis-tance regarding the NMR analysis. The Swedish NMR-Centrum isacknowledged for granting NMR spectrometer time. Financial sup-port was from the Knut and Alice Wallenberg Foundation (MA), theSwedish Research Council, VR (MA), the Beckman Scholars program(JJ), and the NHMFL internal research program (RD and PJ).

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