Interactions of the antimalarial amodiaquine with lipid model membranes.

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CPL 4341 1–12 Chemistry and Physics of Lipids xxx (2015) xxx–xxx

Contents lists available at ScienceDirect

Chemistry and Physics of Lipids journal homepage: www.elsevier.com/locate/chemphyslip

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Interactions of the antimalarial amodiaquine with lipid model membranes

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Rafael P. Barroso, Luis G.M. Basso, Antonio J. Costa-Filho *

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Laboratório de Biofísica Molecular, Departamento de Física, Faculdade de Filosofia, Ciências e Letras de Ribeirão Preto. Av. Bandeirantes, 3900, 14040-901 Ribeirao Preto, SP, Brazil

A R T I C L E I N F O

A B S T R A C T

Article history: Received 12 March 2014 Received in revised form 24 December 2014 Accepted 26 December 2014 Available online xxx

A detailed molecular description of the mechanism of action of the antimalarial drug amodiaquine (AQ) is still an open issue. To gain further insights on that, we studied the interactions of AQ with lipid model membranes composed of dipalmitoylphosphatidylcholine (DPPC) and dipalmitoylphosphatidylserine (DPPS) by spin labeling electron spin resonance (ESR) and differential scanning calorimetry (DSC). Both techniques indicate a coexistence of an ordered DPPS-rich domain with a disordered DPPC-rich domain in the binary DPPC/DPPS system. We found that AQ slightly lowered the melting transition temperatures associated to both domains and significantly increased the enthalpy change of the whole DPPC/DPPS phase transition. DSC and ESR data also suggest that AQ increases the number of DPPC molecules in the DPPC-rich domains. AQ also causes opposing ordering effects on different regions of the bilayer: while the drug increases the ordering of the lipid acyl chains from carbon 7 to 16, it decreases the order parameter of the lipid head group and of carbon 5. The gel phase was mostly affected by the presence of AQ, suggesting that AQ is able to influence more organized lipid domains. Moreover, the effects of AQ and cholesterol on lipid acyl chain ordering and mobility were compared at physiological temperature and, in a general way, they are similar. Our results suggest that the quinoline ring of AQ is located completely inside the lipid bilayers with its phenol ring and the tertiary amine directed towards the head group region. The nonspecific interaction between AQ and DPPC/DPPS bilayers is a combination of electrostatic and hydrophobic interactions. ã 2014 Elsevier Ireland Ltd. All rights reserved.

Keywords: Amodiaquine DSC ESR Antimalarial Spin label

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1. Introduction

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Malaria is still one of the most infectious diseases affecting the developing world. According to the World Health Organization (WHO), 655,000 people die every year due to malaria, out of which 86% are children under five years of age (WHO, 2011). One of the major barriers on fighting malaria is the worldwide drug resistance. Malaria parasites have developed resistance to all current drugs. To overcome drug resistance, WHO recommends the use of combined therapies with two or more drugs, in particular, the combination of amodiaquine (AQ) and artesunate (WHO, 2010). AQ is a 4-aminoquinoline antimalarial drug widely used in the past and whose chemical structure is similar to that of chloroquine (Olliaro et al., 1996 O’Neill et al., 1998). Several countries chose AQ as the first-line drug in combination with artesunate (WHO, 2010). In areas where artesunate resistance is

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* Corresponding author. Tel.: +55 163315 3665; fax: +55 16 33715381. E-mail address: [email protected] (A.J. Costa-Filho).

low or absent, the therapeutic efficacy of artesunate–amodiaquine is highly correlated with the efficacy of AQ alone (Sá et al., 2009). The mechanism of action of 4-aminoquinolines is not completely understood, but it is believed that is related to hemoglobin degradation in the digestive vacuole of the parasite (Pussard and Verdier, 1994; O’Neill et al., 1998; Fitch, 2004). Hemoglobin is degraded by several enzymes of the parasite resulting in toxic heme metabolites. A process of detoxification converts heme into an insoluble crystal called hemozoin or “malaria pigment”, which participates in the parasite life cycle (Sherman, 1977). A complex formed by 4-aminoquinolines with heme (ferriprotoporphyrin IX) inhibits conversion of heme to hemozoin, thus killing the parasite. Although AQ forms complexes with ferriprotoporphyrin IX (Blauer et al., 1993), its complete mechanism of action, at a molecular level, remains unclear. In particular, there is not much information on how the drug overcomes the physical barrier represented by the cell membrane. Membrane lipids are associated with vital cell functions. Besides being the structural blocks of cell membranes and the sources of metabolic energy, lipids play a central role in the regulation of innumerous cellular processes (Escribá, 2006;

http://dx.doi.org/10.1016/j.chemphyslip.2014.12.003 0009-3084/ ã 2014 Elsevier Ireland Ltd. All rights reserved.

Please cite this article in press as: Barroso, R.P., et al., Interactions of the antimalarial amodiaquine with lipid model membranes. Chem. Phys. Lipids (2015), http://dx.doi.org/10.1016/j.chemphyslip.2014.12.003

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Bittman, 2004). Changes in the structure and lipid composition of membranes seem to be related to the development of several diseases (Lee, 2004). Although drug targets are usually proteins (Cohen, 2002; Overington et al., 2006), drugs can also bind to membrane lipids. Non-specific interactions of antiarrhythmics with myocardial membranes, for instance, are related to their mode of action (Gourley, 1971). Amlodipine, an antihypertensive drug, also interacts non-specifically with membranes (Mason et al., 1991). In a previous work, the non-specific interaction of the antimalarial drug primaquine with biological membranes was suggested to represent an additional route in its overall mode of action and/or could be associated to its adverse effects (Basso et al., 2011). The importance of the non-specific drug-membrane interactions as a molecular event can be enhanced by a new therapeutic approach called membrane-lipid therapy. It is based on the binding of drugs to lipids, thus modulating the structure of membranes (Escribá, 2006). To understand the overall mechanism of action of AQ, studies of AQ-membrane interaction are important since the molecular targets of this drug, either proteins or lipids from membranes, are not completely known yet. A combination of techniques is frequently used to study drugmembrane interactions (Seydel and Wiese, 2002). In particular, electron spin resonance (ESR) and differential scanning calorimetry (DSC), among several available techniques, have provided important contributions (Zhao et al., 2007; Budai et al., 2003; Könczöl et al., 2005; Pentak et al., 2008; Hamilton et al., 1991; Basso et al., 2011). Changes in membrane properties induced by drugs or other molecules are detected by lineshape alterations in the ESR spectra or by changes in heat exchange in DSC experiments (Duarte et al., 2008; Basso et al., 2011; Couto et al., 2011; Dyszy et al., 2013). In this paper, interactions of the antimalarial drug AQ with lipid membrane models were investigated by ESR and DSC. The membrane models were constituted by an equimolar mixture of dipalmitoylphosphatidylcholine (DPPC) and dipalmitoylphosphatidylserine (DPPS), which are representative lipids of the parasite membrane (Hsiao et al., 1991). The effects of AQ on the melting behavior of DPPC/DPPS bilayers were monitored by DSC. Using spin labels attached to different positions along the lipid acyl chains and to the head group, we investigated the alterations induced by AQ on the ordering and dynamics of different regions of lipid bilayers by ESR. We also compared the effects caused by AQ with the effects caused by cholesterol on the structural dynamics of the binary system at physiological temperature. A possible drug location in the lipid bilayer was proposed.

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2. Material and methods

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2.1. Materials The lipids 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC), 1,2-dipalmitoyl-sn-glycero-3-phospho-L-serine (DPPS), cholesterol, and the spin labels 1-palmitoyl-2-stearoyl-(ndoxyl)-sn-glycero-3-phosphocholine (n-PCSL, where n = 5, 7, 10, 12, 14, and 16) and 1,2-dioleoyl-sn-glycero-3-phospho(tempo) choline (DOPTC) were obtained from Avanti Polar Lipids, Inc. (Alabaster, AL). The antimalarial drug AQ and the Hepes buffer (4-(2-hydroxyethyl)-1-piperizineethanesulfonic) were purchased from Sigma Chem., Co. (St. Louis). All reagents were used without further purification. 2.2. Methods 2.2.1. Lipid dispersion preparation A lipid film was formed from a chloroform: methanol (2:1) solution of DPPC:DPPS (1:1) or DPPC:DPPS:cholesterol (10:10:1),

and 0.5 mol% of spin label for ESR experiments, by drying under a stream of N2 and left under reduced pressure for 2 h to remove all traces of organic solvent. The film was then hydrated with 10 mM Hepes, at pH 7.4, to a lipid concentration of 20 mM, heated at 60 C for 5 min, and vortexed. Part of the lipid dispersion was diluted with Hepes buffer to a final lipid concentration of 10 mM and another part was diluted with a 2 mM drug buffered solution to final lipid and drug concentrations of 10 and 1 mM, respectively. For ESR measurements with cholesterol-containing samples, a lipid film was formed from stock solutions of pure DPPC, DPPS, and cholesterol and the dried film was hydrated with Hepes to a final concentration of 10 mM of DPPC and DPPS and 1 mM of cholesterol.

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2.2.2. Electron spin resonance ESR measurements were performed with a JEOL FA-200 X-band spectrometer (JEOL Ltd., Tokyo, Japan). Temperature was controlled by a circulator bath that pumps fluid through an aluminum radiator attached to the ESR resonant cavity as described by Alaouie and Smirnov (2006). The samples were placed into 1 mm inner diameter glass capillaries and then concentrated by centrifugation. The glass capillary was accommodated within a quartz tube. The acquisition conditions were: field modulation frequency, 100 kHz; field modulation amplitude, from 1 to 2 G; sweep width, 100 G; microwave power, 10 mW. Data presented here are an average of duplicate experiments and the uncertainties are the respective standard deviations.

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2.2.3. Nonlinear least-squares (NLLS) simulations of ESR spectra NLLS analyses of the ESR spectra were performed using the program Multicomponent (https://sites.google.com/site/altenbach/labview-programs/epr-programs/), a LabVIEW-based version (Altenbach, 2014) of the traditional scheme of Freed et al. for fitting ESR spectra of nitroxide spin probes (Schneider and Freed, 1989; Freed, 1976). The software calculates an ESR spectrum by solving the stochastic Liouville equation, allowing for the quantification of the structural dynamics of spin-labeled biomolecules in terms of motional rates and orientational ordering imposed by the local environment around the nitroxide radical. The dynamics of spin-labeled lipids in lipid bilayers are better described by an axially symmetric rotational diffusion tensor, whose principal components, R// and R?, represent the rotational diffusion rates around axes parallel and perpendicular, respectively, to the mean symmetry axis for the rotation, zR (Schneider and Freed, 1989). For n-PC spin labels (n = 5, 7, 10, 12, 14, and 16), zR is defined along the symmetry axis of the lipid, i.e., the fully extended direction of the acyl chains (see Fig. S1B of the Supporting information). For DOPTC, zR is defined to be collinear to both the N-C4 and the N—O bonds of the tempo-choline group (Ge and Freed, 1998; Ge and Freed, 2003), which makes it also parallel to the magnetic xm axis, in which the magnetic frame is defined (xm points along the N—O bond, zm lies along the 2pz axis of the nitrogen atom, and ym is perpendicular to both – see Fig. S1A). We found in our simulations that the spectra are not very sensitive to R//. Thus, to reduce the number of fitting parameters, N R== =R? qffiffiffiffiffiffiffiffiffiffiffiffiffi 3 was fixed to 1 and R ¼ R2? R== was varied instead. The mobility of

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the lipid acyl chains and the head groups was then reported as R?. The calculated ESR spectra of the spin-labeled lipids are the result of the so-called MOMD (microscopic order-macroscopic disorder) model that takes into account the tendency of the spin labels to become partially ordered with respect to a local director zD (defined as the normal vector to the bilayer surface), but these local directors are randomly oriented (macroscopically disordered) in the sample (Meirovitch et al., 1984). The microscopic molecular ordering of the spin labels is characterized by the order parameters S0 and S2, which are manifested through an orienting potential on

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the lipid head groups and the acyl chains that restricts the range of orientations of the spin probes (Schneider and Freed, 1989; Budil et al., 1996). S0 is a measure of the inclination of zR with respect to the local director, zD, whereas the nonsymmetric order parameter S2 gives the deviation from cylindrical symmetry of the molecular alignment of zR relative to zD. The larger the S0, the larger is the alignment, and the more restricted is the spin label motion. The parameters involved in the simulations were: g-tensor (gxx, gyy, gzz) and hyperfine-tensor (Axx, Ayy, Azz) components, rotational diffusion rates (R?, R//), orienting potential coefficients (c20, c22), and whenever needed, homogeneous lorentzian or Gaussian linewidths. Seed values of the magnetic tensors were obtained from Earle et al. (1994), but small variations in their values were allowed during the fitting process. The strategy of the simulation follows that of Basso et al. (2011). As for the uncertainties of the parameters from the NLLS analyses, we found an estimated error of 0.01 for S0 and 5% for R? of n-PC spin labels, and of 0.006 for S0 and 3% for R? of DOPTC. This means that we found good solutions for the stochastic Liouville equation with different sets of c20, c22 (only used in the simulations of DOPTC and 16-PCSL), and R?. In general, good agreement between the experimental and theoretical spectra is obtained whenever S0 and R? are both simultaneously increased within the uncertainties. Overall, the general trends observed in the ordering and dynamics of the spin labels for different sets of (S0, R?) values and for different lipid sample preparations were all consistent, even though the differences in S0 and mostly in R? might be within the uncertainty of the corresponding parameters. 2.2.4. Differential scanning calorimetry DSC traces were obtained by heating the samples from 5 to 70 C at a scan rate of 20 C/h with the Microcalorimeter VP-DSC (Microcal, Northampton, MA) or the calorimeter Nano-DSC-II (Calorimetry Sciences Corporation, Lindon, Utah, USA). Baseline

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Table 1 Calorimetric parameters of the phase transition of the binary DPPC/DPPS membranes obtained from the DSC traces.

DPPC/DPPS DPPC/DPPS with AQ

Sample

DH (kcal/mol)

T1 ( C)

T2 ( C)

1 2 1 2

17.4 18.9 21.3 22.2

43.23 43.19 43.00 43.00

44.39 44.40 44.20 44.12

subtractions and peak integrals were carried out using MicroCal Origin software.

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3. Results and discussions

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3.1. Calorimetric results show AQ-bilayer interaction

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The effect of AQ on lipid membranes was investigated by determining the thermal behavior of a DPPC/DPPS mixture. Fig. 1 shows the DSC traces of DPPC/DPPS dispersions with and without AQ. Upon heating, the AQ-free samples exhibited a nonideal lipid mixing, as also reported by other authors (Sevcsik et al., 2008), characterized by a broad melting transition. Nonideal lipid mixing means that the lipids are not randomly distributed in the membrane but organized in clusters or domains. In this situation the bilayer displays lateral heterogeneity. This lateral phase separation was previously observed in DPPC/DPPS mixtures (Luna and McConnell, 1977; Gordon et al., 2006). DSC traces of AQ-free and AQ-containing samples presented two calorimetric peaks at temperatures T1 and T2 (T1 < T2, see the inset of Fig. 1). We suggest that these peaks could be attributed to the melting of lipids in two different domains, probably DPPC-rich and DPPS-rich domains. The thermograms also displayed very low enthalpic pre-transition peaks that are neither well defined nor reproducible and thus will not be discussed. The enthalpy change (DH) of the whole transition and the values of T1 and T2 are presented in Table 1. As observed in

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Fig. 1. DSC traces of samples with 10 mM DPPC/DPPS in Hepes buffer with (gray) and without (black) AQ. Inset: zoom of the calorimetric DSC peaks.


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Fig. 2. Experimental (black) and best-fit (gray) ESR spectra of the head group spin label DOPTC and the acyl chain n-PCSL (n = 5, 7, 10, 12, 14, and 16) in DPPC/DPPS membranes in the absence (left) and in the presence (right) of AQ. (a) One-component nonlinear least squares fits of DOPTC, 5-, 7-, 10-, and 12-PCSL ESR spectra. (b) Spectra of 14- and 16PC spin labels were simulated with two components. The arrows show the second, more mobile, and less populated component. The spectra were acquired at 36 C.


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Fig. 1, incorporation of AQ into the DPPC/DPPS lipid mixture increases the peak intensity at T1, slightly lowers both T1 and T2, and significantly increases DH by (20 2)% on average (Table 1). In a previous work, a decrease of the transition temperature was also observed with primaquine, another aminoquinoline antimalarial drug, in DMPC bilayers (Basso et al., 2011). However, a much smaller amount of primaquine (lipid/drug molar ratio 91:1) was required to produce a similar effect than that caused by AQ (lipid/ drug molar ratio 10:1). On the other hand, while the transition enthalpy of DMPC bilayers remained unchanged in the presence of primaquine, AQ induced a large increase in DH of DPPC/DPPS membranes, suggesting different drug-lipid interactions and thus different effects on membranes. Our DSC results show a clear interaction between AQ and the lipid membranes, since AQ changes the shape of the thermograms, decreases the melting temperatures and increases the transition enthalpy of the binary system. These results could indicate a penetration of AQ into the DPPC/DPPS bilayers, since molecules that penetrate lipid bilayers usually lower the gel-to-fluid phase transition (Jain and Wu, 1977). As mentioned before, the narrow DSC peak at T1 (Fig. 1) can be due to the melting of either DPPC-rich or DPPS-rich domains. Using infrared spectroscopy, Schultz et al. showed the coexistence of DPPC-rich and DPPS-rich aggregates in a binary DPPC/DPPS membranes and also showed that DPPC molecules melt at a temperature (45.2 C) lower than that of DPPS (47.8 C) in this lipid system (Schultz et al., 2009). Thus, we propose that the peak at T1 in our DSC thermograms might be due to the melting of DPPC in DPPC-rich domains, whereas the peak at T2 could be related to the melting of DPPS in DPPS-rich domains. Furthermore, lipids do not melt independently of each other but rather simultaneously in clusters of a certain amount of lipids (Heimburg, 2007). Thus, the melting of lipids is a cooperative phenomenon and the cooperativity is proportional to the number of lipids in a cluster or domain. The larger the cooperativity, the narrower and more intense the transition peak. The intensity increase of the DSC peak at T1 caused by AQ binding to the binary system is therefore likely due to an increase of the number of lipids in the domains melting at T1, that is, the size of the DPPC-rich domains increases. Since there is a direct correlation between membrane curvature and both enthalpy changes (Yokoyama et al.

(2013)) and lateral phase separation (Duwe et al., 1990; Sackmann, 2006), the aforementioned effects of AQ on the lipid mixture could be explained by an increase in DPPC/DPPS membrane curvature.

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3.2. ESR results suggest AQ penetration into the bilayer

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Changes in the order and dynamics of the membrane caused by AQ were investigated by means of ESR spin labeling technique at physiological temperature. The hydrophobic region of the bilayer was monitored by using spin labels located at different depths along the lipid acyl chain. Fig. 2a and b shows the experimental and simulated ESR spectra of the head group spin label DOPTC and the acyl chain spin labels n-PCSL (n = 5, 7, 10, 12, 14, and 16) acquired at 36 C in the absence and in the presence of AQ. Generally, the experimental and best-fit theoretical spectra were in very good agreement and considerable changes were observed in all ESR spectra upon incorporation of AQ. Except for the 14- and the 16-PCSL signals (Fig. 2b), all other ESR spectra were simulated with only one component (one spin label population). The twocomponent feature in the 14- and 16-PCSL spectra will be discussed below. Fig. 3 presents the order parameter (S0) profile obtained from the NLLS simulations of the ESR spectra displayed in Fig. 2. Overall, NLLS fits show that AQ increases the ordering and slightly decreases the rotational mobility of the whole hydrophobic region of the lipid bilayers, except for the DOPTC and for 5-PCSL, in which an opposite effect was observed: for example, AQ slightly decreases S0 (from 0.711 to 0.692 – Table S2) and significantly increases R? (from 3.31 to 4.36 107 s1 – Table S2) for 5-PCSL. Interesting features were reported by some of the spin labeled lipids and will be discussed in more details. NLLS simulations of the ESR spectra of the head group spin label DOPTC provided information on the dynamics and on the orientation of the nitroxide radical at the bilayer surface. With a positive value for S0 = 0.518 and a negative value for the nonaxial order parameter S2 = –0.084, the orientation of the trimethyl ammonium (TMA) group of the DOPTC in the binary DPPC/DPPS system is almost similar to that of the dipalmitoylphosphatidyl tempo (2,2,6,6,-tetramethyl-1-oxy) choline (DPPTC) in a pure DPPC dispersion in the gel phase (S0 = 0.50 and S2 = –0.22 at 25 C)

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1.0 DPPC/DPPS - comp 1 DPPC/DPPS - comp 2

0.8 DOPTC

Order parameter

223

5

DPPC/DPPS/AQ - comp 1 DPPC/DPPS/AQ - comp 2

0.6

0.4

0.2

0.0

HG

4

6

8

10

12

14

16

n-PCSL Fig. 3. Order parameter profile obtained from non-linear least-squares simulations of DOPTC and n-PCSL (n = 5, 7, 10, 12, 14, and 16) 36- C ESR spectra in the absence (full circle) and in the presence (open triangle) of AQ. HG stands for head group.


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(Ge and Freed, 1998) or to that of 4-O-(1,2-dipalmitoyl-sn-glycero3-phospho) 4-hydroxy-2,2,6,6-tetramethyl-piperidine-1-oxy (DPP-Tempo) in dioleoylphosphatidylcholine (DOPC) dispersions at –11 C (S0 = 0.516 and S2 = –0.238) in the presence of excess water (Ge and Freed, 2003). This means that the principal diffusion axis (zR), which coincides with xm (parallel to the N—O bond – see Section 2.2.3), is preferentially aligned along the local director of the bilayer surface. The main difference in the orientation of the TMA group of DOPTC in DPPC/DPPS membranes compared to that of the other spin labels (DPPTC in DPPC or DPP-Tempo in DOPC) is the lower preference of either xR or yR axes to be oriented perpendicular to director, due to the low value of S2 (–0.084). To make this clear, we decomposed the spherical tensor components S0 and S2 in their Cartesian counterparts (Ge and Freed, 2003): p p Szz = S0 = 0.518, Sxx = 1/2( 3/2 S2 – S0) = –0.310, and Syy = –1/2( 3/ 2 S2 + S0) = –0.207. As discussed before, the high positive value of Szz means a strong preference for zR to be aligned along the normal to the bilayer, whereas negative and close values of Sxx and Syy indicate an almost equal preference for xR and yR diffusion axes to be oriented tangential to the bilayer. The addition of AQ to DPPC/ DPPS samples changes the ordering and dynamics of DOPTC, which means that AQ binds to and interacts with the lipid head group of the bilayers. Interestingly, AQ significantly decreases both the perpendicular rotational mobility R? (from 3.02 to 2.45 107 s1) and the order parameter S0 (from 0.518 to 0.452) of the spin label. The latter result implies that lipid-AQ interaction increases the average angular amplitude of the wobbling motion of the nitroxide moiety (the larger S0, the smaller the amplitude). In terms of the Cartesian components, Sxx = –0.291, Syy = –0.160, and Szz = 0.452, we can also infer that AQ binding to membranes induces zR to be much less oriented along the local director, whereas xR and yR axes retain their orientation tangential to the bilayer normal, but yR becomes less oriented. Another interesting feature observed in Fig. 2a is that the spectrum of 12-PCSL from the AQ-free sample presented a much broader line than would be normally expected for this spin label. This clearly indicates strong spin–spin interactions that likely arise from cluster formation of 12-PCSL molecules in the DPPC/DPPS membranes. Partial segregation of spin labels was also noted, for instance, by Fajer et al. for n-PCSL in the DPPC gel phase in a saturation transfer ESR study (Fajer et al.,1992) and by Earle et al. in a rigid-limit 250 GHz ESR study of n-PCSL incorporated in DPPC and in palmitoyl-oleoyl-PC (POPC) lipid vesicles (Earle et al., 1994). In fact, the best least-square fits to 12-PCSL ESR spectra were achieved by allowing for a Heisenberg exchange coupling in the simulations, which yielded a vHE = 4.9 107 s1. Interestingly, addition of AQ into the preformed 12-PCSL-clustered DPPC/DPPS lipid vesicles abolished the spectral broadening, thus suggesting disruption of those clusters. Direct lipid-AQ interactions might promote a better miscibility of the 12-PCSL molecules in the binary system. A distinct behavior was observed in the spectra of 14- and 16PCSL (Fig. 2b), which exhibited additional spectral features (see the low-field and high-field lines) characteristic of a second spin population, indicating the coexistence of two types of domains in the 1:1 DPPC/DPPS lipid bilayers, which is in agreement with our DSC results. A two-component system was also observed, for instance, by Basso et al. in the rippled phase of dimyristoyl-PC (DMPC) lipid bilayers (Basso et al., 2011) and by Chiang et al. in a ternary DPPC/dilauroyl-PC/cholesterol (DPPC/DLPC/Chol) lipid mixture (Chiang et al., 2004). Our result shows that ESR is very sensitive to detect the heterogeneity of different micro-environments in a binary system. Thus, an extra ESR signal with different ordering and rotational dynamics was added to the fitting process. From the NLLS fits, we found that the most populated component of the two spin labels presented the higher order parameter: S0 of 0.367 for 14-PCSL (87% population) and of 0.343 for 16-PCSL (93%

population), S0 values of the less ordered domain were 0.196 for 14PCSL (13% population) and 0.117 for 16-PCSL (7% population). As shown in Fig. S2 of the Supporting information, the theoretical 16PCSL spectrum of the ordered domain (component 1) in the binary system is very similar to the experimental spectrum of this same spin label in pure DPPS vesicles at the same temperature. Similarly, the theoretical spectrum of the less ordered component resembles, despite the difference in intensity, that of the 16-PCSL incorporated in pure DPPC lipid bilayers at the same temperature (Fig. S2). Thus, the coexistence of the ordered and disordered regions in the binary DPPC/DPPS lipid system is most likely due to the coexistence of DPPS-rich (ordered) and DPPC-rich (disordered) domains. The slight differences in the spin population values of the ordered and disordered domains for 14- and 16-PCSL suggest that these spin-labeled lipids have different partition coefficients KP between those domains. Since the mole fraction of 14-PCSL in the less ordered state is higher than the population of 16-PCSL in this same domain (0.13 and 0.07, respectively), 14-PCSL must partition better in the disordered domains than 16-PCSL. In fact, a slighter preference of 14-PCSL for the disordered phase compared to 16PCSL in a two-phase coexisting region of a ternary system has been reported (Swamy et al., 2006). Upon AQ addition to the lipid samples, changes in the lineshape of the 14- and 16-PCSL are observed (Fig. 2b). The nonlinear leastsquares fits show that binding of AQ to the DPPC/DPPS membranes increases the ordering (S0: from 0.367 to 0.413 for 14-PCSL and from 0.343 to 0.379 for 16-PCSL) and decreases the perpendicular rotational diffusion rate (R?: from 6.76 to 6.31 107 s1 for 14-PCSL and from 2.09 to 1.78 108 s1 for 16-PCSL) of the DPPS-rich domains (Table S2). The overall result is an increased packing of the bilayer center of the DPPS-rich domain, whereas almost no change was observed on ordering and dynamics of the center of the hydrophobic core of the DPPC-rich domains (Table S2). However, both 14- and 16-PCSL spectra of the AQ-containing samples show an increase of the spin-labeled lipid population of the less ordered DPPC-rich domain (from 0.13 to 0.28, for 14-PCSL, and from 0.07 to 0.17, for 16-PCSL). The redistribution of the spin label molecules between the two domains upon AQ addition might be a result of two main effects: (1) changes in the equilibrium constant between the DPPC-rich and DPPS-rich domains and/or (2) changes in the partition coefficient of the spin labels between the two regions. We explored these two possibilities as follows. Firstly, we defined the partition coefficient KP as the ratio of spin-label concentration in the ordered (Lo) and disordered (Ld) states as in Swamy et al. (2006): KP ¼

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PðLo Þ ½mol%ofLd x PðLd Þ ½mol%ofLo

where P(Ld) and P(Lo) are the populations of the spin labels incorporated in the disordered (Ld) and ordered (Lo) domains, respectively, obtained from the NLLS fits of the ESR spectra, and the second term on the right hand side of the equation represents the percentage of lipids in the DPPC-rich (Ld) and DPPS-rich (Lo) domains. KP would be unity if spin labels partition equally into each domain. If we further suppose that the relative populations (percentage of lipids) of the ordered and disordered domains are characteristic of the binary system and thus do not change with the addition of 0.5 mol% of the spin label (this is reasonable since the acyl chain of spin-labeled lipids are almost identical to that of DPPC and DPPS), the ratio K between KP values for 14- and 16-PCSL can be directly calculated as the ratio of the spin label populations obtained from the NLLS fits: K¼

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K 14PC P K 16PC P

¼

412 411 413 414 415 416 417 418 419 420 421 422 423 424 425

½PðLo Þ=PðLd Þ14PC ½PðLo Þ=PðLd Þ16PC


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Thus, K gives a good estimate of the relative partition coefficient of both spin labels into the Ld and Lo phases. We found that K = 0.504 for the AQ-free sample and K = 0.527 for the AQcontaining sample, that is, the difference is within the uncertainty of the spin label populations obtained from NLLS simulations, which is 3%. This result suggests that the partitioning of the spin labels in the ordered and disordered domains does not depend on the AQ binding to membranes at 36 C (at least on the specific conditions of our experiments). Consequently, changes on spin label population upon AQ binding reflect changes on the equilibrium constant between DPPC-rich and DPPS-rich domains in the binary system. As mentioned before, our DSC experiments suggest an increase of the cooperativity of the DPPC-rich domains upon AQ incorporation (see Section 3.1). Our ESR results also support this hypothesis. Binding of AQ to DPPC/DPPS membranes could shift the equilibrium constant between DPPC-rich and DPPS-rich domains towards the DPPC-rich one. As a result, there may be an increase in the number of lipids in the DPPC-rich clusters, which can contribute to the increased cooperativity of the phase transition of this domain. Taken together, the ESR/NLLS results suggest that AQ inserts into the DPPC/DPPS membranes and changes the molecular force field, i.e., the orienting potential, in the headgroup and in the acyl chain regions. We suggest that both the phenol ring and the protonated tertiary amine of AQ (at pH 7.4, AQ is mostly monoprotonated) are most likely located in the headgroup region of the lipid bilayer. This can enhance the hydrogen bond network of that region (due to additional hydrogen bonds between AQ and the lipid head groups) and also decrease electrostatic repulsion among

7

PS in DPPS-rich domains. Direct AQ-head group interaction via the aforementioned groups also changes the orientation of the TMA group of the DOPTC spin label, which impacts the membrane orienting potential. Additionally, except for the local environment around carbon 5, the increased S0 values of almost the entire acyl chain region indicate an increased lateral packing density of the lipids induced by AQ, which can enhance van der Waals interactions between lipids. This increased bilayer ordering is consistent with a higher transition enthalpy, since more heat is required to break molecular interactions and melt the lipids. The slight reduction of S0 with a remarkably increase of rotational mobility around carbon 5 – as shown by NLLS simulations of 5-PCSL spectra – can be explained by the insertion of the quinoline ring of AQ into the lipid matrix. With the quinoline ring located into the hydrophobic core of the lipid bilayer, but close to the polar–apolar interface, and the phenol ring OH and the tertiary amine directed towards the head group, a free volume could be created around the very first carbon atoms of the lipid acyl chains, which can explain the markedly increase of the rotational degree of freedom of 5-PCSL. Insertion of the quinoline structure into lipid matrix may also enhance van der Waals attractions between lipids, which can contribute to increase the lateral packing. In fact, insertion of both unprotonated and protonated forms of the quinoline ring of another 4-aminoquinoline antimalarial drug, chloroquine, into the apolar region of an anionic micelle has been also proposed (Santos et al., 2014; Usman and Siddiq, 2013). To gain further insights on the hypothesis of AQ penetration into the membrane, as strongly suggested by our results, we also compared the effect of AQ on acyl chain order and mobility with

Fig. 4. Experimental (black) and best-fit (gray) of ESR spectra of the head group spin label DOPTC and the acyl chain n-PCSL (n = 5, 7, 10, 12, 14, and 16) in either DPPC/DPPS (left) or DPPC/DPPS/Chol (right) membranes. Spectra of 14- and 16-PC spin labels were simulated with two components. All the spectra were acquired at 36 C.


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CPL 4341 1–12 8 485 486 487 488 489 490 491 492 493 494 495 496 497 498 499 500 501 502 503 504 505 506 507 508 509 510 511 512 513 514 515 516 517 518 519 520 521 522


those provoked by cholesterol. It is well known that cholesterol is located within the lipid bilayers (Huang, 1977; Ahmad and Mellors, 1978; Subczynski et al., 1992) and, as will be discussed further, cholesterol is also soluble in PS bilayers. Fig. 4 shows the ESR spectra of DOPTC and n-PCSL (n = 5, 7, 10, 12, 14, and 16) in DPPC/DPPS (1:1) and DPPC/DPPS/Cholesterol (1:1:0.1) model membranes and Table S2 summarizes the dynamics and order parameters obtained from the NLLS simulations. As before, the spectra of 14- and 16-PCSL were simulated with two components, with the greater contribution (most populated) coming from the ordered, DPPS-rich domains, for both spin labels (Table S2). Incorporation of 5 mol% of cholesterol into the binary DPPC/ DPPS system causes a rigidifying effect on the whole acyl chain of the lipids (Table S2). Cholesterol also provokes a depth-dependent effect on the perpendicular rotational diffusion rate of the lipids: a slight reduction of R? from carbon 5 to carbon 7 and then a prominent increase of R? from carbon 12 to the end of the lipid acyl chain (Table S2). This depth-dependent effect of cholesterol on the structural dynamics of lipid acyl chains in lipid bilayers is well known (Costa-Filho et al., 2003; Mainali et al., 2013). Besides that, cholesterol causes no effect on the ordering of the head groups (S0 = 0.52; S2 = –0.11), but slightly reduces the rotational mobility (from 3.16 to 3.02 107 s1), as monitored by DOPTC spin label. Despite the small difference in R? (4.4%), this variation is higher than the uncertainty calculated for this parameter (3% – see Section 2.2.3). Also, the observed trend of reduced head group mobility in cholesterol-containing samples is obtained in different lipid preparations: there is a consistent decrease of the ratio between the intensity of the low-field and the center-field lines of the DOPTC ESR spectrum in the presence of cholesterol. Changes in lipid order parameters can be used to infer how deep foreign molecules go inside the lipid bilayers (Biaggi et al., 1997). To gain further insights into the localization of AQ inside DPPC/DPPS bilayers, we plotted the variation of S0 induced by AQ or cholesterol (Chol) relative to the pure lipid bilayer as a function of the labeled carbon atoms of the lipid acyl chain (Fig. 5). AQ and Chol affect the order parameter along the whole acyl chain with Chol effects

decreasing from n = 7 down the chain, whereas AQ-induced alterations oscillate in the same region. Furthermore, the increase in S0 at carbons 14 and 16 caused by AQ in the more ordered domain is larger than that caused by cholesterol (component 1 – Fig. 5). Unlike Chol, AQ also causes a significant perturbation of the headgroup organization, which indicates a distinct mode of organization of the drug in the bilayer. The previous experiments were carried out with AQ addition to preformed liposomes (see Section 2), which means that only the external part (the outer leaflet) of the bilayers was available to interact with AQ. However, cholesterol was not incorporated into the bilayers by the same procedure, but rather by preparing the liposome as a lipid film constituted of DPPC, DPPS, and cholesterol. Thus, as both the inner and outer monolayers were available to cholesterol, the effect of AQ on the lipid ordering and mobility might be even stronger than that caused by cholesterol. To test this hypothesis, we qualitatively studied the effect of AQ on the acyl chain ordering and dynamics when the drug was also allowed to interact with the inner monolayers of the liposomes. In this condition, we compared the effect of AQ and Chol on the lineshape of 14-PCSL spectra at temperatures above and below the phase transition temperatures (above T2 and below T1) of the binary system (Fig 6a). Above the phase transition, there is no change in the lineshape of the spectra due to addition of AQ. However, cholesterol slightly decreases the ratio between the high-field and the center-field lines, as indicated by the arrow in the figure, which means that 5 mol% of Chol slightly decreases the mobility and/or increases the ordering of the region monitored by 14-PCSL. On the other hand, below the phase transition, both AQ and cholesterol broaden the spectrum relative to the pure lipid mixture. This means that both molecules increase the bilayer ordering, but the effect caused by AQ is stronger than that of cholesterol, which corroborates our initial hypothesis. The lineshape alterations in the 14-PCSL spectrum caused by incorporation of AQ into the inner and outer monolayers (Fig. 6a) are even more pronounced than those caused by incorporation of the drug only into the outer monolayer (see Fig. 2b). The effect of cholesterol on the heat capacity of a DPPC/DPPS/Chol lipid dispersion was also similar to the effect

Fig. 5. Variation of the order parameter, S0, relative to the pure lipid bilayer as a function of nitroxide position, n, of n-PCSL in DPPC/DPPS membranes in the presence of AQ or cholesterol. Temperature: 36 C.


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Fig. 6. (a) ESR spectra of 14-PCSL in DPPC/DPPS membranes in the absence and in the presence of AQ or Chol, in the gel phase (left) and fluid phase (right). (b) DSC traces of 10 mM DPPC/DPPS in Hepes buffer with (gray) and without (black) cholesterol. 561 562 563 564 565 566 567 568 569 570 571 572

caused by AQ, though there was no change in the transition enthalpy (Fig. 6b). Despite the low miscibility of cholesterol in phosphadityl serines (PS) bilayers reported by several authors (Bach and Wachtel, 2003), the effect of limiting amounts of cholesterol on the thermotropic behavior of binary phospholipid mixture indicated that anionic phospholipids exhibit greater affinity for cholesterol than do the zwitterionic phospholipids PC and PE (Van Dijck, 1979). McMullen et al. (2000) also showed that cholesterol exhibits greater miscibility in anionic PG than in uncharged glycolipid bilayers. A personal communication published in the McMullen et al. paper (McMullen et al., 2000) reports an up-to 67 mol% cholesterol solubility in 1-palmitoyl-oleoyl-PS

bilayers. Cholesterol incorporation, up to 15 mol%, in the positively charged DODAB bilayers was also reported (Benatti et al., 2007). We used a cholesterol concentration of 5 mol% of the total lipid and, considering its reported higher affinity to PS compared to PC, we suggest that cholesterol partitions better in PS-rich domains than in PC-rich ones. The incorporation of cholesterol in the bilayer is related to hydrophobic effects between the cholesterol and the lipid bilayer. Since AQ is a hydrophobic drug, and it is worth noting that its hydrophobicity has been linked to its antimalarial activity (Elslager et al., 1964), we expect that hydrophobic effects between AQ and the bilayers could also take place. However, besides the


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hydrophobic effect, electrostatic forces between AQ and the negatively charged DPPC/DPPS bilayers may occur, since at the Q4 pH studied, about 98% of AQ was in the monoprotonated form ( Naisbitt et al., 1997). To verify the role of the bilayer charge in the interaction with AQ, we measured the effect of the same amount of AQ on the ESR spectrum of 12-PCSL in DPPC/DPPS bilayers with increasing amounts of DPPS (Fig. 7). Although variations can be observed even in the absence of DPPS, which means that AQ also interacts with pure DPPC bilayers, it is only with 50% that we observed significant changes in the ratio h+1/h0, a parameter that increases to unity with increasing disordering. This indicates that the bilayer surface charge has a crucial role in the AQ-bilayer interaction. 3.3. Possible role of drug-membrane interactions to the antimalarial mechanism of action The amphiphilic or hydrophobic character of many pharmacologically active compounds confers them different types of association, such as self-association and specific and/or nonspecific binding to proteins and lipid membranes. Usually, the plasma membrane is one of their sites of action and, even if the target is intracellular, the interaction with this first barrier plays an important role (Schreier et al., 2000). Since several antimalarials are amphiphilic, such as AQ, chloroquine, primaquine, quinacrine, mefloquine, and quinine, drug-membrane interactions may contribute to their whole mechanism of action. Indeed, interactions between halofantrine and lipid bilayers of PC and PE were observed by DSC measurements (Lim and Go, 1999). Halofantrine caused a broadening of the phase transition of PC bilayers and a decrease in both enthalpy and transition temperature. A similar

Fig. 7. Ratio between the amplitudes of the low- and central-field lines (h+1/h0) of 12-PCSL in 10 mM of DPPC and different amounts of DPPS in the absence (closed square) and in the presence (open square) of AQ. Temperature: 36 C.

effect of halofantrine was observed with PE. Our results also showed a decrease in the transition temperature caused by AQ. In another study, the effects caused by halofantrine on the phase transition of DPPC bilayers were compared with those caused by quinine and mefloquine (Lim and Go, 1995). Similar to halofantrine, the latter drugs also interacted with DPPC bilayers, changing the phase transition and decreasing the transition temperature. Pajeva et al. showed that quinacrine was also able to interact with membranes, since the drug caused changes in the thermotropic behavior of DPPS bilayers by decreasing the transition enthalpy (Pajeva et al., 1996). Primaquine-membrane interactions were shown by a combination of pressure perturbation calorimetry, DSC and ESR (Basso et al., 2011). It was shown that primaquine stabilizes the fluid phase of different lipid model membranes, causing a local disorder on the acyl chains of the lipids. This fluidizing effect seems to be due to the interaction of primaquine with the lipid head groups, causing a less dense gel phase. Interactions of mefloquine, quinacrine, chloroquine, and quinine with DPPC bilayers were investigated by 2H and 31P NMR (Zidovetzki et al., 1989). Although chloroquine did not seem to affect the bilayer structure of DPPC, a weak chloroquine–DPPC interaction could not be disregarded by the authors. Their results also pointed that quinacrine interacts only with the surface of the DPPC bilayers, whereas both quinine and mefloquine penetrate into the membrane. A further investigation of DPPC–chloroquine interactions using a fluorescent probe inserted into the bilayer showed that chloroquine rigidified DPPC membranes (Ghosh et al., 1995). Our results also show a rigidifying effect on the hydrophobic core of the bilayers caused by AQ. It is believed that the heme is the target of both AQ and chloroquine, since these drugs prevent heme detoxification. The parasite protects itself against the toxicity of heme by polymerizing it to insoluble hemozoin. As weak bases, AQ and chloroquine accumulate in the acidic environment of the parasite's food vacuole and inhibit heme polymerization. After exiting the food vacuole, the heme that is not polymerized is degraded by glutathione. Ginsburg et al. found that AQ and chloroquine competitively inhibit the degradation of heme by glutathione, allowing it to accumulate into the membranes (Ginsburg et al., 1998). Heme, which is released as ferriprotoporphyrin IX (FPIX) in the presence of oxygen (Jacob and Winterhalter, 1970), disrupts the membrane barrier properties and disturbs the ion homeostasis in the cells, which kills the parasites. Famin et at. showed that the inhibition constant of glutathione-mediated destruction of heme in ghost membranes by AQ is 25% greater than by chloroquine (Famin et al., 1999). It was supposed that this difference might be related to different locations of AQ, chloroquine, and FPIX within the membranes. It was proposed that chloroquine is located mainly in the head group region of the lipid bilayer, but not intercalated into the phospholipids (Ginsburg and Demel, 1983; Kursch et al., 1983 Zidovetzki et al., 1989). However, AQ is much more hydrophobic than chloroquine (the partition coefficient of AQ in octanol is 359% greater than that of chloroquine, according to Bray et al., 1996) and thus it may intercalate in the bilayer, as we showed in the present paper. In this way, the non-specific interaction between lipids and AQ, as other antimalarial drugs, may have an important role in their overall and complex mechanism of action.

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4. Conclusions

672

We showed that AQ is able to interact non-specifically with DPPC/DPPS lipid bilayers affecting the gel phase more than the lipid fluid phase. AQ seems to influence more organized lipid domains, since the fluid phase of the model membrane investigated was not affected. This AQ-bilayer interaction is driven

673


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by a combination of electrostatic forces and hydrophobic effects. Firstly, the approximation between the drug and the bilayer surface is driven by electrostatic forces between the positively charged amino group of AQ and the negatively charged PS headgroups. Once close the bilayer, hydrophobic effects also take place. Our results indicate that AQ binding to the binary DPPC/ DPPS system shifts the equilibrium constant between the disordered DPPC-rich and the ordered DPPS-rich domains towards the DPPC-rich one. We suggest that AQ is located completely inside the bilayer with the positively charged amino group and the OH of the phenol group pointing outwards to the polar headgroups of the bilayer.

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Conflict of interest

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The authors declare that there are no conflicts of interest.

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Transparency document

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The Transparency document associated with this article can be found in the online version.

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Acknowledgments

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This research was supported by USP, FAPESP (Grants No. 2010/ 17662-8, 2011/1755-1 and 2012/20367-3) and CNPQ. The authors thank Professor P. Ciacanglini and Professor M. T. Lamy for allowing access to the DSC equipments.

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Appendix A. Supplementary data

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Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j. chemphyslip.2014.12.003.

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Wiley-VHC, Weinheim. Hsiao, L.L., Howard, R.J., Aikawa, M., Taraschi, T.F., 1991. Modification of host cell membrane lipid composition by the intra-erythrocytic human malaria parasite Plasmodium falciparum. Biochem. J. 274, 121–132. Huang, C.H., 1977. A structural model for the cholesterol–phosphatidylcholine complexes in bilayer membranes. Lipids 12, 348–356. Jacob, H., Winterhalter, K., 1970. Unstable hemoglobins: the role of heme loss in Heinz body formation. Proc. Natl. Acad. Sci. U. S. A. 65, 697–701. Jain, M.K., Wu, N.M., 1977. Effect of small molecules on the dipalmitoyl lecithin liposomal bilayer: III. Phase transition in lipid bilayer. J. Membr. Biol. 34, 157–201. Könczöl, F., Farkas, N., Dergez, T., Belágyi, J., LÅrinczy, D., 2005. Effect of tetracaine on model and erythrocyte membranes by DSC and EPR. J. Therm. Anal. Calorim. 82, 201–206. Kursch, B., Lüllmann, H., Mohr, K., 1983. Influence of various cationic amphiphilic drugs on the phase-transition temperature of phosphatidylcholine liposomes. Biochem. Pharmacol. 32, 2589–2594. Lee, A.G., 2004. How lipids affect the activities of integral membrane proteins. Biochim. Biophys. Acta 1666, 62–87. Lim, L.Y., Go, M.L., 1995. A differential scanning calorimetry study of the interaction of the antimalarial agent halofanthrine with dipalmitoyl phosphatidyl choline bilayers. Chem. Pharm. Bull. 43, 2226–2231. Lim, L.Y., Go, M.L., 1999. The antimalarial agent halofantrine perturbs phosphatidylcholine and phosphatidylethanolamine bilayers: a differential scanning calorimetric study. Chem. Pharm. Bull. 47, 732–737. Luna, E.J., McConnell, H.M., 1977. Lateral phase separations in binary mixtures of phospholipids having different charges and different crystalline structures. Biochim. Biophys. Acta 470, 303–316.


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G Model

CPL 4341 1–12 12 822 823 824 825 826 827 828 829 830 831 832 833 834 835 836 837 838 839 840 841 842 843 844 845 846 847 848 849 850 851 852 853 854 855 856 857 858 859 860 861 862


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α-Synuclein interactions with phospholipid model membranes: Key roles for electrostatic interactions and lipid-bilayer structure.

Opioid peptide interactions with lipid bilayer membranes.

Protein-lipid interactions in bilayer membranes: a lattice model.

Lipid-polypeptide interactions in bilayer lipid membranes.

Lipid-protein interactions in membranes.

Interactions of the anticancer drug tamoxifen with lipid membranes.

Lipid and peptide specificities in signal peptide--lipid interactions in model membranes.

Interactions of an antimicrobial peptide, tachyplesin I, with lipid membranes.

Amphiphilic drug interactions with model cellular membranes are influenced by lipid chain-melting temperature.

Interactions between lipid vesicles and cell membranes.

Nanoparticles meet cell membranes: probing nonspecific interactions using model membranes.

Repulsive interactions and mechanical stability of polymer-grafted lipid membranes.

Theoretical study of protein--lipid interactions in bilayer membranes.

Buffers affect the bending rigidity of model lipid membranes.

Influence of amodiaquine on the antimalarial activity of ellagic acid: crystallographic and biological studies.

Lipid-Loving ANTs: Molecular Simulations of Cardiolipin Interactions and the Organization of the Adenine Nucleotide Translocase in Model Mitochondrial Membranes.

Continuous flow atomic force microscopy imaging reveals fluidity and time-dependent interactions of antimicrobial dendrimer with model lipid membranes.

Lipid Interactions and Organization in Complex Bilayer Membranes.

Electrostatic interactions among hydrophobic ions in lipid bilayer membranes.

Lipid peroxidation modifies the assembly of biological membranes "The Lipid Whisker Model".

Introducing New Antimalarial Analogues of Chloroquine and Amodiaquine: A Narrative Review.

Interactions of a bacterial trehalose lipid with phosphatidylglycerol membranes at low ionic strength.

Magnetic resonance studies on the interaction of antidepressants with lipid model membranes.

A general model for the interaction of foreign molecules with lipid membranes: drugs and anaesthetics.