Revisiting lipid – general anesthetic interactions(I): Thinned domain formation in supported planarbilayers induced by halothane and ethanol

Zoya V. Leonenko and David T. Cramb

Abstract: A long-standing question in anesthesia is that of the molecular mechanism. Do anesthetics target proteins orchange membrane properties or both? We used temperature-dependent magnetic A/C mode atomic force microscopy(AFM) to study interaction of the volatile anesthetics halothane and ethanol with model membranes made from sup-ported planar bilayers (SPBs) of 1,2-dioleoyl-sn-3-glycero-3-phosphocholine (DOPC), dioleoyltrimethylammonium pro-pane (DOTAP), or 1,2-dipalmitoyl-sn-3-glycero-3-phosphocholine (DPPC). We found that the incorporation of halothaneor ethanol induces structural changes in the bilayer. These compounds cause thickness reduction in Lα bilayers (eitherglobally or in domains) and the formation of domains with reduced thickness in Lβ phase bilayers. We propose that ananesthetic-induced increased area per lipid drives local chain disorder, thus promoting local phase change. The charac-teristics of SPBs with halothane or ethanol incorporated were compared with characteristics of the Lα and Lβ phases ofanesthetic-free SPBs.

Key words: atomic force microscopy, anesthesia, lipid bilayer domains, phase transition

Résumé : Le mécanisme moléculaire de l’anesthésie n’a toujours pas été résolu. On se demande encore est-ce que lesproduits anesthésiques s’attaquent aux protéines ou s’ils changent les propriétés des membranes ou agissent-ils sur lesdeux fronts. On a fait appel à la microscopie de forces atomiques (MFA) en mode A/C magnétique et dépendante dela température pour étudier l’interaction d’anesthésiques volatiles, tels l’halothane et l’éthanol, avec des membranesmodèles faites à partir de bicouches planaires supportées (BPS) de 1,2-dioléoyl-sn-3-glycéro-3-phosphocholine (DOPC),de dioléoyltriméthylammonium propane (DOTAP) ou de 1,2-dipalmitoyl-sn-3-glycéro-3-phosphocholine (DPPC). On aobservé que l’incorporation d’halothane ou d’éthanol induit des changements structuraux dans la bicouche. Ces compo-sés provoquent une réduction de l’épaisseur des bicouches Lα (globalement ou dans les domaines) accompagnée de laformation de domaines comportant des épaisseurs réduites dans les bicouches de la phase Lβ. On suggère qu’une sur-face par lipide accrue, induite par l’anesthésique, provoque un désordre de chaîne local qui induit un changement dephase local. On a comparé les caractéristiques des BPS dans lesquelles on a incorporé de l’halothane ou de l’éthanolavec les caractéristiques des phases Lα et Lβ de BPS ne contenant pas d’anesthésique.

Mots clés : microscopie de forces atomiques, anesthésie, domaines de bicouches lipidiques, transition de phase.

[Traduit par la Rédaction] Leonenko and Cramb 1138

Introduction

The phenomenon of domain formation in model mem-branes is of great interest, as membrane structural changesdefine changes in membrane functional properties. This mayalso be true for anesthesia. Indeed, there have been paradigmshifts in the molecular theory of anesthetic action over thepast century. The correlation found by Meyer and Overton(1, 2) between anesthetic potency and lipid solubility sug-gested that alteration of the lipid membrane structure waskey to anesthetic efficiency. As membrane protein studies

became possible at the molecular level, attention shifted tothe direct action of anesthetics on ligand-gated ion channels(3, 4). More recently, it has been suggested that volatile an-esthetics indirectly affect membrane protein by altering thelateral pressure within bilayers (5). Also, a multiple-sites-of-action hypothesis has been forwarded in which general anes-thetics may bind to many protein and lipid sites with rela-tively low affinity (1–10 mmol/L) (6). Despite considerablestudy, the exact molecular effect of volatile anesthetics onmembranes is only beginning to emerge. It has been demon-strated that volatile anesthetics can induce a change in lipidpacking in membranes (7–9). For example, several experi-mental and theoretical studies were devoted to investigationof changes in membrane properties due to the incorporationof alcohols. For example, Holte and Gawrish (10) found thatethanol partitions near the lipid–water interface close to thelipid glycerol backbone and upper segments of lipid hydro-carbon chains and increases the degree of motional disorderin lipid chains. The implication is that a contributing factorin the physiological effect of ethanol results from the mole-

Can. J. Chem. 82: 1128–1138 (2004) doi: 10.1139/V04-023 © 2004 NRC Canada

1128

Received 8 October 2003. Published on the NRC ResearchPress Web site at http://canjchem.nrc.ca on 27 May 2004.

Z.V. Leonenko and D.T. Cramb.1 Department of Chemistry,University of Calgary, 2500 University Drive NW, Calgary,AB T2N 1N4, Canada.

1Corresponding author (e-mail: dcramb@ucalgary.ca).

cules dissolving in specific lipid hydrophobic sites, whichsignificantly modifies membrane structure and may lead tobilayer thinning. Thinned bilayers could contain both inter-digitated lipids (interpenetration of hydrocarbon chains ofthe upper leaflet to the lower leaflet of the bilayer, resultingin disappearance of the bilayer midplane) and lipids withvastly increased tilt angles. It has been reported that bilayerthinning occurs when the local concentration of the speciesin the bilayer is higher than a threshold value (11). Bilayerthickness changes may have a strong effect on the functionof membrane-associated proteins because of a mismatch ofthe protein–lipid hydrophobic interaction region. Changes inlipid packing in the presence of ethanol have been examinedby X-ray diffraction, and they have been suggested in fluo-rescence spectroscopic studies (12, 13). Interdigitated do-mains caused by ethanol were proposed by Mou et al. (14)to explain the thinned dipalmitoylphosphatidylcholine(DPPC) bilayer they observed using atomic force micros-copy (AFM). Recently, AFM studies by McClain and Breen(15) suggest that 2-propanol-induced DPPC bilayer thinningis strictly due to interdigitation.

Halothane is a clinical anesthetic that has received in-creasing attention as a model molecule for fundamentalstudies on anesthetic action. Molecular dynamics simula-tions (16) of halothane in a DPPC bilayer revealed that halo-thane self-associates and also is distributed nonuniformlyacross the membrane interior. This resulted in significantmodifications of the lipid bilayer, including thinning. A de-crease in lipid wobble and an increase in tail defects of thelipid system were also observed.

A temperature-dependent AFM study may provide valu-able information about the effect of anesthetics on thethinned-bilayer formation by monitoring the effect of anes-thetics on the chain-melting phase transition. Very recently,three temperature-dependent AFM studies involving bilayershave been reported (17–19). In these papers, increases intemperature were used to induce Lβ–Lα phase changes in thesupported planar bilayers (SPBs). In our studies of DNA –cationic bilayer interactions as a function of temperature, wehave shown that topology can be accurately measured usingmagnetic A/C (MAC) mode AFM to temperatures as high as70 °C in a liquid sample cell (19). Temperature-dependentAFM allows a novel examination of halothane-inducedchanges in lipid bilayer properties and permits powerful re-examination of some previous results (14) for anesthetic –DPPC bilayer interactions, because the characteristics ofboth the gel- and the liquid-phase SPB can be measured onthe same sample in the presence and absence of anesthetic.

In the present work, we have concentrated our efforts onstudying the incorporation of ethanol and halothane into SPBs.We employed MAC mode AFM to study anesthetic-inducedchanges in supported phospholipid bilayers of dioleoyl-phosphatidylcholine (DOPC), DPPC, or dioleoyltrimethyl-ammonium propane (DOTAP). The goal of this study was toextend the knowledge that exists in the literature for ethanoland to examine the incorporation of halothane into a bilayer.We were also able to measure the bilayer properties as afunction of temperature and to induce the Lβ–Lα phase tran-sition in DPPC, and we used a temperature increase to irre-versibly drive ethanol out of the SPBs. In the followingcompanion paper (20), the results of our AFM studies for

halothane are compared with those obtained using spectro-scopic techniques.

Materials and methods

Dioleoylphosphatidylcholine (DOPC, lyophilized or inchloroform solution), dioleoyltrimethylammonium propane(DOTAP, chloroform solution), and dipalmitoylphosphatidyl-choline (DPPC) were obtained from Avanti Polar Lipids Inc.(Alabaster, Ala.) and were used without further purification.Tris–EDTA (TE) buffer, pH 7.6, and distilled, ultrapurewater were used in the generation of all vesicles. Freshlycleaved ASTMV-2 quality, scratch-free ruby mica(Asheville-Schoonmaker Mica Co., Newport News, Va.) wasused throughout this study as a substrate. In some cases,mica was modified by addition of a dilute solution of 3-aminopropyltriethoxysilane (APTES) using standard proce-dures (21).

All vesicles were prepared using the “dry” method as pre-viously reported (22). An appropriate aliquot of phospho-lipid chloroform solution was measured into a small vessel,and the chloroform was removed using a stream of dry nitro-gen. The dry phospholipid was then resuspended in buffer toits final concentration and stirred for 30 min. The solutionwas sonicated (Branson 1200; Branson Ultrasonics,Danbury, Conn.) at room temperature for 10-min periods.Between each 10-min period, there was a 15-min “rest” in-terval where the solution was placed in an ice bath and thenstirred at room temperature. For this method, the solutionswere cycled an average of 10 times or until they were ob-served to clear.

Supported planar bilayers were prepared for AFM imag-ing by the method of vesicle fusion (22, 23). Aliquots ofliposome solution were deposited on modified or unmodifiedfreshly cleaved mica. After a controlled period of time, themica was gently rinsed with ultrapure water, and the surfacewas imaged under water in the liquid cell, at room tempera-ture and at various higher temperatures.

Addition of ethanol and halothane was performed in twoways: (i) by addition of pure anesthetic into the vesicle solu-tion with an incubation time of 1.5 h and (ii) by addition ofan ethanol/water mixture (50% by volume) or pure liquidhalothane into the AFM liquid cell with the established sup-ported bilayer and incubation for the same period of time. Atime of 1.5 h was shown by fluorescence experiments to beoptimal for halothane distribution between buffer and mem-brane (20).

We employed MAC mode AFM, where a magneticallycoated probe oscillates near its resonant frequency driven byan alternating magnetic field. This technique has proven tobe advantageous for measuring supported planar bilayers inliquid media (22, 24, 25). All images were taken using aPico SPM microscope with AFMS-165 scanner (MolecularImaging Inc., Phoenix, Ariz.). Magnetically coated type IMaclevers® (Molecular Imaging Inc.) were used for MACmode imaging. Their specifications include a length of85 µm, a force constant of 0.06–0.1 N/m, a resonant fre-quency of 20–40 kHz in water, and a tip radius of curvaturequoted as less than 10 nm. The standard MAC mode fluidcell (Molecular Imaging Inc.) was used throughout. Thescanning speed was 1–2 lines per second. The height scale

© 2004 NRC Canada

Leonenko and Cramb 1129

was calibrated using colloidal gold spheres of 5 nm and14 nm in diameter (26). For elevated-temperature experi-ments, the AFM Temperature Controller and Hot Mac ModeStage from Molecular Imaging Inc. were used. Heating andcooling were performed at ∆T = 1 °C/min, with the possibil-ity to hold at any given temperature. The mean value(s) ofthe bilayer thickness was calculated using between 50 and70 measurements over several scanned regions, where ap-propriate. The standard deviation is quoted at the 99% confi-dence level.

Results and discussion

DPPCThe changes in a Lβ DPPC bilayer structure as a function

of incorporation of halothane and ethanol were examinedand compared with the effect of temperature. The reason forusing ethanol was to compare our data, obtained by MACmode AFM, with the results of previous studies (14, 15).There are several studies in the literature that have examinedincorporation of ethanol and other alcohols into phospholi-pid bilayers using various methods, including fluorescence(27) and AFM (14, 15). It has been suggested that a highconcentration of ethanol in a lipid-containing aqueous solu-tion can induce transformation of a lipid bilayer from (Lβ)gel phase to the fully interdigitated (LβI) phase of mono-unsaturated phosphatidylcholine (PC) lipids, which alters theLβ–Lα melting transition temperature of the phospholipidbilayer (28). In our study, ethanol was added to a supportedDPPC bilayer on mica. After 1.5 h, the cell was then rinsedand imaged in water. In the absence of ethanol, we measuredthe thickness of a pure supported DPPC bilayer on mica as5.5 nm, from natural defects, which is in good agreementwith the value of 5.7 nm reported earlier (14). We observedthe formation of thin domains in the presence of ethanol(Fig. 1). The thickness of the higher domains was 5.6 nm,and that of the lower domains was 3.6 nm (Table 1). Thedifference in thickness of the two domains was 2.0 nm,compared with 1.9 nm previously reported (14). Theethanol/SPB system remained stable at room temperature forup to 2 days. Changing the relative concentration of ethanolonly changed the extent of thin domains, but not their thick-ness. The stability of the thin domains suggests that ethanolremains preferentially in the bilayer at room temperature.When the bilayer was heated to 50 °C (i.e., above the melt-ing transition point, Tm = 41 °C) while imaging was per-formed and then slowly cooled back to room temperature,the lower domains disappeared and the original bilayerthickness was restored. This reversibility of the ethanol ef-fect is in agreement with data of Mou et al. (14), althoughthe latter authors were not able to simultaneously image andscan temperature. These findings suggest that ethanol

repartitions into water when DPPC is at elevated tempera-ture and that this repartitioning depends on the phase of thebilayer. Recently, we investigated the temperature depend-ence of a pure DPPC SPB (data not shown). As the tempera-

© 2004 NRC Canada

1130 Can. J. Chem. Vol. 82, 2004

Fig. 1. Topography image (a), cross section (b), and phase image(c) of a supported dipalmitoylphosphatidylcholine (DPPC) bilayerin the presence of ethamol obtained by AFM. An EtOH–watermixture (50:50 by volume) was added to a supported DPPCbilayer, incubated for 1.5 h, and then rinsed with pure waterprior to the AFM measurements. The scale bar in panel (c) is500 nm long.

ture approached the Lβ–Lα phase transition, thinned domainsappeared. The heights of the domains were exactly the sameas those induced by ethanol, described above. When holdingat a temperature of 50 °C, the lower domains coalesced andgrew in extent. There is every reason to believe that thesethinner domains represent a DPPC SPB in its Lα state. Thevalue of 3.6 nm for Lα DPPC thickness is in the range previ-ously determined using X-ray scattering (Xhead-to-head = 3.6–4.0 nm) (29). A similar range was also calculated by Felleret al. (30) by molecular dynamics simulation. Interestingly,the simulation illustrated that the DPPC lipid chains in theLα phase displayed both some interdigitation and significanttilting. However, there was no indication of the completelyinterdigitated LβI phase, where methyl end groups would beexposed to water.

We have also examined the incorporation of halothaneinto a DPPC bilayer to compare the effects of ethanol andhalothane. The halothane was added into the vesicle solution

or into the liquid cell with a preformed supported bilayer. InFig. 2 we illustrate the effects of incorporating halothaneinto vesicles prior to SPB formation. In the presence ofhalothane, we observed similar domain formation in a DPPCbilayer to that induced by ethanol. The thickness of thehigher and lower domains was measured as 5.5 and 3.5 nm,respectively, giving the same difference of 2.0 nm as we ob-served in the case of ethanol (Table 1). The surface coverageof the thin domains was 2–3% at lower halothane concentra-tions (Fig. 2b) and comparable to that naturally formed inpure DPPC SPBs (1–2%). At higher halothane content, weobserved a considerable amount of lower domain surfacecoverage (25–50%, Fig. 2c). We were able to image the sur-face during slow heating of the bilayer above the meltingtransition point (Tm = 41 °C). Heating the system to 42 °Cdid not induce considerable changes in the domain structure.After 1 h of scanning at 42 °C, lower domains in the bilayerwere still present. A further increase in temperature to 50 °C

© 2004 NRC Canada

Leonenko and Cramb 1131

Thickness after addition of ethanol Thickness after addition of halothane

SPBaThickness of purelipid bilayer

CEtOH in water =75 mmol/L

CEtOH in water =1.4 mol/L MR = 3.6:1 MR = 7.3:1

DOTAP 3.7 ± 0.2 3.6 ± 0.2 3.5 ± 0.22.9 ± 0.2

DOPC 5.4 ± 0.1 5.3 ± 0.3 3.5 ± 0.2 5.0 ± 0.5 3.6 ± 0.23.5 ± 0.2 3.6 ± 0.3

DPPC 5.5 ± 0.1 5.6 ± 0.2 5.5 ± 0.1 5.5 ± 0.13.6 ± 0.3 3.5 ± 0.1

Note: MR is the ratio of the number of moles of halothane in the membrane to the number of moles of lipids.aDOTAP, dioleoyltrimethylammonium propane; DOPC, dioleoylphosphatidylcholine; DPPC, dipalmitoylphosphatidylcholine.

Table 1. Dependence of thickness (nm) of supported planar bilayers (SPBs) on the incorporation of halothane and ethanol.

Fig. 2. Topography images and cross sections showing the effects of incorporation of halothane into a DPPC bilayer. (a) Pure DPPCbilayer; (b) DPPC with halothane, at an estimated molar ratio of halothane in the membrane to lipid of 3.4:1; (c) DPPC withhalothane, molar ratio = 14.6:1. In these experiments, halothane was added into the vesicle solution prior to SPB formation.

demonstrated that the remaining thicker domains melted at ahigher temperature than the pure DPPC free-standing bilayerand the phase transition was broad. This is consistent withthe melting of a pure DPPC SPB (data not shown).

In the alternate preparation, halothane was added into theliquid cell containing the supported DPPC bilayer coveredwith 1 mL of water. After the addition of halothane, no rins-ing was performed. In Fig. 3, we follow the SPB containing

© 2004 NRC Canada

1132 Can. J. Chem. Vol. 82, 2004

Fig. 3. Topography images and cross sections of supported DPPC bilayer with halothane recorded as a function of temperature andtime. Halothane was added into the liquid cell containing a supported DPPC bilayer covered with 1 mL of water; no rinsing was per-formed for this sample. Halothane remained in the water and the sample was scanned during heating. (a) Room temperature; (b) 30-min scanning at room temperature; (c) heated to 50 °C; (d) heated to 58 °C; (e) heated to 62 °C; (f) cooled to 55 °C; (g) cooled to50 °C; (h) cooled to room temperature. The estimated molar ratio of halothane in the membrane to lipid was 3.4:1.

halothane as a function of temperature and time. In this casewe observed a very interesting transition of the bilayer con-taining halothane at room temperature during the first30 min of scanning. Domains (thickness of higher domainswas 4.5 ± 0.2 nm) initially present (Fig. 3a) disappeared andthe bilayer became generally thin (3.3 ± 0.3 nm, Fig. 3b).Because the bilayer was supported on mica, we consider thathalothane first causes structural changes in the upper leaflet,

and scanning at room temperature might induce equilibra-tion of halothane between the two leaflets. After this thinnerbilayer is formed, the system is stable during scanning atroom temperature and becomes thinner yet at a temperaturehigher than that of the Lβ–Lα transition for a pure DPPCbilayer. Further thinning was observed to start at 50 °C andwas complete at 62 °C (Figs. 3c–e). After the system wascooled back to room temperature (Figs. 3f–h), bilayer thick-

© 2004 NRC Canada

Leonenko and Cramb 1133

Fig. 3 (concluded).

ness was 3.9 ± 0.2 nm, which is lower than that of pure LβDPPC bilayer. The initial thickness of pure DPPC withouthalothane was restored during a similar cooling process(data not shown). The molar ratio of the amount of halo-thane in the membrane to that of lipid was estimated asnHal/nL = 3.4 at room temperature and has been shown to in-crease by a factor of 4 above the chain-melting transitiontemperature (31). The amount of halothane in the bilayerwas estimated using the partition coefficient of halothane inDPPC provided by Simon et al. (31). This increase inhalothane partitioning during heating is probably responsiblefor a saturation of the bilayer with halothane and stabiliza-tion of thinned domains when the system was cooled back toroom temperature.

The persistence of thinner domains and the complete thin-ning of the bilayer show that interaction of halothane withthe lipids is very strong and increases at elevated tempera-ture. These results are consistent with those reported in thecompanion paper (20) as well as with those obtained by Si-mon et al. (31) and by Hill (32).

When compared with ethanol, halothane shows differentbehavior during heating. Whereas ethanol leaves the bilayer,thus restoring its initial thickness, halothane partitioning in-creases with temperature, leading to more extensive domainformation. The reason for this difference in behavior likelyoriginates from the higher hydrophobicity of halothane andits lower solubility in water (17 mmol/L) (20) comparedwith that of ethanol. Ethanol can escape during heatingwhen the mobility of phospholipids and ethanol is high,whereas for solubility reasons halothane remains in thebilayer.

The transitions observed upon heating the bilayer withand without halothane were compared. For a pure DPPCbilayer, we observed a broad main transition between 42 °Cand 52 °C and further change between 52 °C and 60 °C;upon cooling back to room temperature, a transition between35 °C and 36 °C occurred. When the bilayer with halothaneincorporated was heated, the lower temperature transitionwas not observed, and only one transition at 52–60 °Coccurred. The presence of halothane likely changes struc-tural properties of the bilayer in such a way that the othertransitions observed for a pure bilayer are not possible. Thetransition below Tm likely involves reorganization of watermolecules at the interface. In the presence of halothane, wa-ter molecules are exchanged for halothane molecules, whicheliminates the low-temperature transition.

The inhomogeneous domain formation observed with bothethanol and halothane incorporation is presumably due to acooperative mechanism of anesthetic penetration and parti-tioning into the bilayer. This mechanism may include local-ized dehydration of headgroups by disruption of thecoordinated hydration shell. The existence of such ahydration shell has been suggested by molecular dynamicssimulations (33). Additionally, the initial rupture of thehydration shell may promote inhomogeneous anesthetic in-corporation into the bilayer at the exposed location. Afterwater shell rupture, further incorporation of anesthetic intothe bilayer increases the headgroup area and motional disor-der in lipid chains and leads to formation of the thinnedstate. The possible role of water in the mechanism of anes-thesia was emphasized in previous studies (33, 34). Ueda

and co-workers (34, 35) have obtained evidence, throughproton NMR measurements and FTIR spectroscopy, for therelease of bound interfacial water by inhalation anestheticsin a water-in-oil emulsion. This suggested that some anes-thetics compete with water for binding sites. Both ethanoland halothane are able to form hydrogen bonds. Halothanecontains a so-called acidic hydrogen, which is considered tobe involved in a “hydrogen bond breaking effect”. It hasbeen shown that anesthetics with “acidic hydrogen” andtherefore with high “hydrogen bond breaking” ability havehigher anesthetic potency than halogenated molecules con-taining no acidic hydrogen (36). An NMR study (10) hasshown that ethanol incorporates into the lipid–water inter-face near the lipid glycerol backbone and upper segments oflipid hydrocarbon chains and increases the degree ofmotional disorder in the lipid chains, which promotes forma-tion of a thinned bilayer.

Molecular dynamics simulations (16) indicate thathalothane induces lateral expansion and contraction of theLα DPPC bilayer. Koubi et al. (16) reported a thickness re-duction of 8 Å (from 42 to 34 Å; 1 Å = 0.1 nm) and an in-crease in area per lipid of 16% (from 63.6 to 72.0 Å2). Weobserved a thickness reduction from 5.5 to 3.5 nm (2.0 nm)in the Lβ DPPC bilayer due to incorporation of halothane.We also have observed a thinning of a pure DPPC bilayer to3.6 nm upon heating above 42 °C. Thus, incorporation ofhalothane produces a similar thinning effect as heating thebilayer above the chain melting transition temperature. Thedifference in an Lα DPPC SPB with and without halothaneis small (0.1 nm) compared with that predicted (0.8 nm(16)). This might reflect the differences between supportedplanar bilayers and free-standing bilayers.

The fact that very similar DPPC bilayer changes are ob-served for both ethanol and halothane incorporation suggestsa common mechanism. Moreover, because the formation ofthin bilayer domains surrounded by thick domains is analo-gous to what we observe for a DPPC SPB near the Lβ–Lαphase transition temperature, it is likely that ethanol andhalothane induce something similar to a local phase changein the SPB. The mechanism may include initially smallerLα-like domains coalescing into larger thin domains. Thestability of these domains for periods of days would be dueto a cage effect by the surrounding thicker and less mobileLβ regions. Fluorescence results from our companion paper(20) also suggest that increased disorder of the lipid packingand penetration of water near the headgroup region of DPPCsmall unilamellar vesicles is induced by the incorporation ofhalothane.

DOPCTo gain insight into how the lipid phase might affect the

interaction of ethanol and halothane with SPBs, we usedDOPC. DOPC exists in a fluid phase (Lα) at room tempera-ture and serves as a fluid-phase analog of the DPPC bilayer.A pure DOPC SPB (Fig. 4a) has a thickness of 5.4 nm, aswe reported earlier (22). In the presence of ethanol (Fig. 4b)or halothane (Fig. 4c) at a molar ratio of 7.2:1, the DOPCSPB transforms to a uniform thickness of 3.5 and 3.6 nm,respectively (Table 1). We assume that the higher lipid disor-der of a DOPC bilayer results in much less cooperativity in

© 2004 NRC Canada

1134 Can. J. Chem. Vol. 82, 2004

© 2004 NRC Canada

Leonenko and Cramb 1135

Fig. 4. Topography images and cross sections showing the effects of incorporation of ethanol or halothane into a dioleoylphos-phatidylcholine (DOPC) bilayer. (a) Pure DOPC bilayer; (b) DOPC bilayer with ethanol, CEtOH in water = 1.4 mol/L; (c) DOPC bilayerwith halothane, at an estimated molar ratio of halothane in the membrane to lipid of 7.5:1. Ethanol and halothane were added into thevesicle solution prior to SPB formation. The scale bars in all panels are 500 nm long. The arrows in all cross sections represent4.0 nm in height.

Fig. 5. Topography images and cross sections of DOPC bilayer with halothane, at an estimated molar ratio of halothane in the mem-brane to lipid of 3.7:1. (a) Initial scan; (b) after 40 min of scanning. Halothane was added into the vesicle solution prior to SPB for-mation. The scale bars in all panels are 500 nm long. The arrows in all cross sections represent 4.0 nm in height.

the incorporation of anesthetic at these higher molar ratios,and thus no domains are formed.

Interestingly, at a lower concentration of halothane (molarratio = 3.6:1), domain formation was observed (Fig. 5a).These domains were 5.0 and 3.6 nm thick (Table 1), and thedomain shape and size changed during 40 min of scanning(Fig. 5b). The fact that DOPC domains were not well re-solved and more dynamic compared to DPPC bilayer maybe explained by higher mobility of anesthetic in DOPCbilayer than in DPPC Lβ bilayer. Thinned DOPC domainswere observed in the presence of ethanol as well (Fig. 6a).

Although the ethanol-induced domains were more stableat room temperature than those induced by halothane, theystarted to disappear at 35 °C as the sample was heated andsimultaneously imaged. We found that heating the DOPCbilayer with ethanol incorporated and then cooling it back toroom temperature revealed changes in SPB surface coverage(Figs. 6a–c) and bilayer thickness. Initially, we observed abilayer with domains of 5.5 and 3.5 nm. Upon heating andslow cooling, the bilayer was restored to 5.3 nm. This im-plies that the release of ethanol from DPPC is largely due toelevated temperature rather than the lipid phase, since theethanol–DOPC system is stable at room temperature but re-leases ethanol at elevated temperature.

For an SPB with anesthetic incorporated, changes in sur-face coverage as a function of temperature can be used toprovide insight into the amount of lateral expansion the SPBundergoes during incorporation. For the ethanol-containingbilayer, surface coverage increased by 42.3% (compared tothe surface coverage at room temperature) upon heating to43 °C (observed after 50 min of scanning at 43 °C). After

© 2004 NRC Canada

1136 Can. J. Chem. Vol. 82, 2004

Fig. 6. Topography images and cross sections obtained during heating and cooling of DOPC bilayer with ethanol incorporated. (a) Atroom temperature; (b) at 42 °C; (c) after cooling back to room temperature. Ethanol (CEtOH in water = 75 mmol/L) was added into thevesicle solution prior to SPB formation. The scale bars in all panels are 500 nm long. The arrows in all cross sections represent4.0 nm in height.

Fig. 7. Topography image and cross section of DOPC aggregatesdeposited on mica. Vesicles were prepared in an aqueous solu-tion of ethanol (3 mol/L). The scale bar is 500 nm long. The ar-row in the cross section represents 12.5 nm in height.

the bilayer was slowly cooled back to room temperature, thesurface coverage was decreased by 6.2% compared to theinitial value before heating. The increase in surface area at43 °C (with a concomitant decrease in thickness to 3.6 nm)indicates an increase in lipid disorder. Restoring of thebilayer full thickness and a decrease of surface area indicateethanol escape and formation of the Lα DOPC bilayer. Forour conditions, we estimate that incorporation of ethanolproduces a 10% increase in DOPC area per lipid comparedto that of a pure DOPC bilayer at room temperature. This es-timation has been made on the basis of surface changes weobserved and known DOPC area per lipid in pure DOPCbilayer without ethanol (ADOPC = 64 Å2) (37). The increasein DOPC area per lipid can be compared to that of an LαDPPC bilayer (16%) in the presence of halothane at highconcentrations (16) and to that of DPPC in the presence of2-propanol (15) (57.8% — from 52 Å2 to 90 Å2). Thesmaller increase in the case of DOPC can be explained bytaking into account the liquid phase of DOPC bilayer, whichis more disordered compared to gel-phase DPPC bilayer,and also to differences in the volume of incorporated mole-cules.

At a higher concentration of ethanol (3 mol/L) in the vesi-cle incubation solution, we observed formation of smalldisk-shaped aggregates and clusters of these aggregates uponexposure of the solution to a mica surface (Fig. 7). Nobilayers were observed for this preparation. The height ofthe circular aggregates was in the range 10–15 nm. We as-sume that these aggregates could be flattened vesicles,which were stabilized by the partitioning of more ethanolmolecules in the upper leaflet. This seems to make vesicles

more susceptible to a pre-fusion type of aggregation but re-sistant to rupture upon adsorption to the mica surface. Mouet al. (14) mentioned previously the existence of such smallglobular structures in the presence of high concentrations ofethanol.

DOTAPTo help elucidate the influence of the headgroup on the

anesthetic-induced changes in SPB structure, we performedanalogous experiments using DOTAP. DOTAP is Lα at roomtemperature, and its headgroup carries a positive charge atneutral pH. A pure DOTAP supported bilayer was describedin our earlier publication (18). Halothane was incorporatedinto the DOTAP bilayer via incubation in the vesicle solu-tion at a 7.3:1 molar ratio of halothane to lipid. An AFM im-age of the resulting SPB is presented in Fig. 8. Although aDOTAP SPB is already very thin, we again observed forma-tion of thinner domains. The thickness of the higher domainwas 3.5 nm and that of the lower domain was 2.9 nm (Ta-ble 1). The difference in thickness of the two domains was0.5–0.6 nm. This difference is close to that calculated(0.4 nm) by Koubi et al. (16) for halothane in Lα DPPC us-ing molecular dynamics. We assume that high lateral mobil-ity of lipid in a DOTAP bilayer, which exists in Lα phase atroom temperature, explains the higher mobility of the do-mains that we observed (Fig. 8).

Conclusions

We have demonstrated by AFM that the thinned domainsin DOPC, DPPC, and DOTAP supported planar bilayers in-

© 2004 NRC Canada

Leonenko and Cramb 1137

Fig. 8. Topography images and cross sections of a dioleoyltrimethylammonium propane (DOTAP) bilayer with halothane, at an esti-mated molar ratio of halothane in the membrane to lipid of 7.3:1. (a) At room temperature; (b) at 30 °C, 30-min scanning. An aque-ous solution of halothane (41 mmol/L) was added into the vesicle solution prior to SPB formation. The scale bars in all panels are500 nm long. The arrows in all cross sections represent 4.0 nm in height.

duced by halothane and ethanol are similar, regardless ofwhether the headgroup is the zwitterionic PC or the cationicTAP. This is consistent with the molecular dynamics simula-tions of Koubi et al. (16) for halothane in the Lα phase ofDPPC, where halothane was observed to partition preferen-tially on the lipid side of the headgroup region. A room tem-perature DPPC bilayer showed stable domains when eitherethanol or halothane was present in the bilayer. Upon heat-ing above the phase transition temperature, ethanol was re-leased whereas halothane was retained. This indicates thatthe effects we observe are solely due to anesthetic within thebilayer and not an effect of anesthetic content in aqueous so-lution. In a fluid-phase DOPC bilayer, inhomogeneous, dy-namic domains were formed at a ratio of incorporatedanesthetic to lipid of less than 7:1. DOPC SPBs completelythinned when this ratio was 7.3:1. The fact that domain for-mation was almost universally observed implies that anes-thetic-induced bilayer thinning is a cooperative process. Forgel-phase bilayers, it seems that volatile anesthetics induce alocalized phase transformation to Lα, whereas for lipids al-ready in Lα, even thinner domains are generated. Heating ofthe DPPC bilayer with halothane showed that halothane af-fects the transition in the bilayer. For the anesthetics studiedhere, the mechanism likely involves partitioning of the anes-thetic preferentially near the headgroups. This is in agree-ment with the findings of the following companion paperand with previous theoretical and experimental studies. Thefact that thinning seems to be universal means that this ef-fect cannot be ignored when considering the physiologicalmechanism of anesthesia, since membrane thinning may af-fect the function of some membrane-associated proteins.

Acknowledgements

We thank Professor Y. Tarahovsky for helpful discussionsand Molecular Imaging Inc. for continuous support. This re-search is supported by the Natural Sciences and EngineeringResearch Council of Canada.

References

1. H.H. Meyer. Arch. Exp. Pathol. Pharmakol. 46, 338 (1901).2. C.E. Overton. Studies of narcosis. Chapman and Hall, London.

1990.3. J.W. Tanner, J.S. Johansson, P.A. Liebman, and R.G.

Eckenhoff. Biochemistry, 40, 5075 (2001).4. J.E. Sirois, Q. Lei, E.M. Talley, C. Lynch, and D.A. Bayliss. J.

Neurosci. 20, 6347 (2000).5. R.S. Cantor. Biophys. J. 80, 2284 (1994).6. G.E. Morgan, Jr., and M.S. Mikhail. In Clinical anesthesiol-

ogy. Appleton and Lange, Stamford, Conn. 1996. pp. 109–120.7. E.S. Rowe. Biochemistry 26, 46 (1987).8. S.A. Simon and T.J. McIntosh. Biochim. Biophys. Acta, 773,

169 (1984).

9. P. Nambi, E.S. Rowe, and T.J. McIntosh. Biochemistry, 27,9175 (1988).

10. L.L. Holte and K. Gawrish. Biochemistry, 36, 4669 (1997).11. E.S. Rowe and J.M. Champion. Biophys. J. 67, 1888 (1994).12. H. Komatsu, P.T. Guy, and E.S. Rowe. Chem. Phys. Lipids, 65,

11 (1993).13. L.T. Boni, S.R. Minchey, W.R. Perkins, P.L. Ahl, J.L. Slater,

M.W. Tate, S.M. Gruner, and A.S. Janoff. Biochim. Biophys.Acta, 1146, 247 (1993).

14. J. Mou, J. Yang, C. Huang, and Z. Shao. Biochemistry. 33,9981 (1994).

15. R.L. McClain and J.J. Breen. Langmuir, 17, 5121 (2001).16. L. Koubi, M. Tarek, M. Klein, and D. Scharf. Biophys. J. 78,

800 (2000).17. M.C. Giocondi, V. Vie, E. Lesniewska, P.E. Milhiet, M. Zinke-

Allmang, and C. Le Grimellec. Langmuir, 17, 1653 (2001).18. Z. Leonenko, D. Merkle, S.P. Lees-Miller, and D.T. Cramb.

Langmuir, 18, 4873 (2002).19. Z. Leonenko and D. Cramb. D. Nanoletters, 2, 305 (2002).20. A. Carnini, H.A. Phillips, L.G. Shamrakov, and D.T. Cramb.

Can J. Chem. 82 (2004). This issue.21. S.M. Lindsay, Y.L. Lyubchenko, A.A. Gall, L.S. Shlyakhtenko,

and R.E. Harrington. Proc. SPIE (Scanning ProbeMicroscopies), 1639, 84 (1992).

22. Z.V. Leonenko; A. Carnini, and D.T. Cramb. Biochim.Biophys. Acta, 1509, 131 (2000).

23. A.A. Brian and H.A. McConnell. Proc. Natl. Acad. Sci. U.S.A.81, 6159 (1984).

24. W. Han, S.M. Lindsay, and T. Jing. Appl. Phys. Lett. 69, 1(1996)

25. W. Han and S.M. Lindsay. Appl. Phys. Lett. 72, 1656 (1998).26. J. Vesenka, S. Manne, T. Giberson, E. Marsh, and E.

Henderson. Biophys J. 65, 992 (1993).27. Q.-T. Li and W.K. Kam. J. Biochem. Biophys. Methods, 35,

11 (1997).28. T.J. McIntosh, H. Lin, S. Li, and C. Huang. Biochim. Biophys.

Acta, 1510, 219 (2001).29. J.F. Nagle, R. Zhang, S. Tristram-Nagle, W. Sun, H.I.

Petrache, and R.M. Suter. Biophys. J. 70, 1419 (1996).30. S.E. Feller, R.M. Venable, and R.W. Pastor. Langmuir, 13,

6555 (1997).31. S.A. Simon, T.J. McIntosh, P.B. Bennet, and B.B. Shrivastav.

Mol. Pharmacol. 16, 163 (1979).32. M.W. Hill. Biochim. Biophys. Acta, 356, 117 (1974).33. L. Perera, U. Essman, and M.L. Berkowirz. Langmuir, 12,

2625 (1996).34. T. Yoshida, H. Okabayashi, K. Takahashi, and I. Ueda.

Biochim. Biophys. Acta, 772, 102 (1984).35. Y.S. Tsai, S.M. Ma, H. Kamaya, and I. Ueda. Mol. Pharmacol.

31, 623 (1987).36. G. Trudeau, K.C. Cole, R. Massuda, and C. Sandorfy. Can. J.

Chem. 56, 1681 (1978).37. N. Cuvillier, F. Millet, V. Petkova, M. Nedyalkov, and J.-J.

Benattar. Langmuir, 16, 5029 (2000).

© 2004 NRC Canada

1138 Can. J. Chem. Vol. 82, 2004