Validity and applicability of membrane model systems for studying interactions of peripheral membrane proteins with lipids

BBAMCB-57568; No. of pages: 11; 4C: 2, 3, 4 Biochimica et Biophysica Acta xxx (2014) xxx–xxx

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Review

Validity and applicability of membrane model systems for studying interactions of peripheral membrane proteins with lipids☆ Aleksander Czogalla ⁎, Michał Grzybek, Walis Jones, Ünal Coskun ⁎⁎ Laboratory of Membrane Biochemistry, Paul Langerhans Institute Dresden, Faculty of Medicine Carl Gustav Carus at the TU Dresden, Fetscherstrasse 74, 01307 Dresden, Germany German Center for Diabetes Research (DZD), Germany

a r t i c l e

i n f o

Article history: Received 24 October 2013 Received in revised form 12 December 2013 Accepted 17 December 2013 Available online xxxx Keywords: Phospholipid Membrane Protein–lipid interaction Model membrane system Lipid presentation Lipid identity

a b s t r a c t The cell membrane serves, at the same time, both as a barrier that segregates as well as a functional layer that facilitates selective communication. It is characterized as much by the complexity of its components as by the myriad of signaling process that it supports. And, herein lays the problems in its study and understanding of its behavior — it has a complex and dynamic nature that is further entangled by the fact that many events are both temporal and transient in their nature. Model membrane systems that bypass cellular complexity and compositional diversity have tremendously accelerated our understanding of the mechanisms and biological consequences of lipid–lipid and protein–lipid interactions. Concurrently, in some cases, the validity and applicability of model membrane systems are tarnished by inherent methodical limitations as well as undefined quality criteria. In this review we introduce membrane model systems widely used to study protein–lipid interactions in the context of key parameters of the membrane that govern lipid availability for peripheral membrane proteins. This article is part of a Special Issue entitled Tools to study lipid functions. © 2013 Elsevier B.V. All rights reserved.

1. Introduction Eukaryotic cells invest much of their proteome, energy and time in the production of membrane components and in the maintenance of characteristic membrane properties such as spatiotemporal distribution of membrane lipid species, generation of surface curvature and bilayer asymmetry. Membranes are sites of multiple physiological events, such as signaling, transport, fusion, and fission. They are complex in composition and dynamic in nature and can be either Abbreviations: AFM, atomic force microscopy; BLM, black lipid membrane; Cer, ceramide; CTX-B, B subunit of cholera-toxin; DPH, diphenylhexatriene; DTT, dithiothreitol; ER, endoplasmic reticulum; ESCRT, endosomal sorting complexes required for transport; FCS, fluorescence correlation spectroscopy; GM1, monosialotetrahexosylganglioside; GPMV, giant plasma membrane vesicle; GUV, giant unilamellar vesicle; ITC, isothermal titration calorimetry; ITO, indium tin oxide; Ld, liquid disordered; Lo, liquid ordered; LUV, large unilamellar vesicle; MLV, multilamellar vesicle; NEM, N-ethyl maleimide; PA, phosphatidic acid; PC, phosphatidylcholine; PE, phosphatidylethanolamine; PEG, polyethylene glycol; PFA, paraformaldehyde; PI, phosphatidylinositol; PIP, phosphatidylinositol phosphate; PS, phosphatidylserine; PVDF, polyvinylidene difluoride; SLB, supported lipid bilayer; SM, sphingomyelin; SPR, surface plasmon resonance; SUV, small unilamellar vesicle; TIRF, total internal reflection fluorescence; TLC, thin layer chromatography ☆ This article is part of a Special Issue entitled Tools to study lipid functions. ⁎ Correspondence to: A. Czogalla, Laboratory of Membrane Biochemistry, Paul Langerhans Institute Dresden, Faculty of Medicine Carl Gustav Carus at the TU Dresden, Fetscherstrasse 74, 01307 Dresden, Germany. Tel.: +49 351 796 36604; fax: +49 351 796 6698. ⁎⁎ Correspondence to: Ü. Coskun, Laboratory of Membrane Biochemistry, Paul Langerhans Institute Dresden, Faculty of Medicine Carl Gustav Carus at the TU Dresden, Fetscherstrasse 74, 01307 Dresden, Germany. Tel.: + 49 351 796 5340; fax: + 49 351 796 6698. E-mail addresses: [email protected] (A. Czogalla), [email protected] (Ü. Coskun).

connected with each other or separated spatially and temporally. Studying the role of these multiple components and their mutual relationship is a challenge, particularly when attempting to reconstitute membrane biology in synthetic membrane systems. The biology of the membrane can be described at three levels, first, at the level of the diversity and complexity of the respective membrane compartments, second, their chemical composition and, third, their functional consequences. With respect to membrane components, the endoplasmic reticulum (ER) plays a dominant role in the synthesis of most structural glycerophospholipids and cholesterol [1]. The latter is rapidly transported to other organelles and, as a result, only low concentrations of sterols can be detected in the ER. Complex sphingolipids are mostly synthesized in the Golgi apparatus. The gradient of sterols and sphingolipids along the secretory pathway is reflected in the rather loose packing of ER membranes, facilitating the insertion and transport of newly synthesized lipids and proteins [2,3]. This is in contrast to the 5–10-fold enrichment of these lipids within the plasma membrane itself [4,5] (Fig. 1). The composition of the membrane is remarkable for the diverse lipid chemical identities, which is based upon variability of the lipid head group and fatty acid chains. What is also of note is the complexity in the lateral distribution and the variable head group presentation of lipids in biological membranes. These are key parameters in governing lipid recognition by peripheral membrane proteins. The vital role and significance of lipid recognition are best exemplified by the heterogeneous distribution of the phosphorylated phosphoinositides (PIPs). The levels and turnover of PIPs are carefully controlled via the locally orchestrated action of numerous specific

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Please cite this article as: A. Czogalla, et al., Validity and applicability of membrane model systems for studying interactions of peripheral membrane proteins with lipids, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbalip.2013.12.012

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Fig. 1. Intracellular sorting of lipids resulting in distinct physicochemical properties of biological membranes. Schematic representation of intercellular lipid sorting and its consequences in the generation of membranes with distinct physicochemical properties. (Upper left panel) Membrane order followed by florescence lifetime imaging of human erythroleukemia cells, using di-4-ANEPPS as membrane environment-sensitive dye. Short lifetime corresponds to less ordered intracellular membranes (blue), while long lifetimes denote ordered and more densely packed membranes, such as the plasma membrane (red). FLIM image was kindly provided by Dr. Michal Majkowski, Department of Cytobiochemistry, University of Wroclaw.

kinases, phosphatases and phospholipases [6,7]. Although PIPs comprise only around 1% of the total cellular lipids, as lipid ligands they play fundamental roles in many biological processes [6,8]. They are involved in membrane recruitment and targeting of peripheral membrane proteins that participate in enzymatic lipid conversion, membrane sculpturing and cell signaling, to name a few [6,9]. Until recently, the majority of protein–lipid interactions were interpreted as simple receptor–ligand binding events, where a given protein interacts specifically with the head group of a lipid ligand. The majority of peripheral membrane proteins containing PIP-binding pleckstrin homology (PH) domains locate to specific cellular membrane compartments but behave in a promiscuous manner in synthetic model systems, showing a rather broad range of specificities and affinities to a set of PIPs. It is supposed that mutual association and cooperative effects with other membrane lipids, additional proteins, or both may amplify their initial low affinity of interaction [10,11]. This model has restricted applicability, however, since various PIPs are not restricted to one membrane compartment only, as exemplified by PI(4)P. It is present in both the Golgi and the plasma membrane, although individual proteins recognize it only in the one or the other compartment [3,12]. Thus, the promiscuous in vitro binding data may not explain the cellular specificity. Some additional interaction mechanism is necessary to account for the specificity of the physical recognition that is encoded as cryptic specificity in the cellular context. A partial explanation of the above may reside in the modulation of the physicochemical properties and, even, conformation of the lipid head groups themselves. This parameter is not solely based on the dependence on the membrane composition, but also on the actual presentation of the molecules in their lipidic environment, giving rise to the concept of lipid head group modulation, referred to as lipid presentation. This concept is difficult to determine experimentally. Apart from the specificity of the membrane composition needed to achieve such an impact, the biological effect is invariably a transient and dynamic one. However, an example of the importance of the actual local lipid environment modulating the behavior of specific lipidic components

is the modification of head group ionization of PA and PI(4,5)P2 , which are modulated by PE and PI, respectively [13,14] (Fig. 2A). Hydrogen bonding between PA and PE results in deprotonation of the phosphomonoester of PA, lowering its pKa and increasing its net negative charge. An exceptional role in modulating lipid presentation in natural membranes is played by sterols, represented by cholesterol in mammalian cells, which are the major modulators of fluidity within a bilayer [4,15]. For instance the plasmalemmal cholesterol was found to play an essential role in controlling accessible pool of PI(4,5)P2 in a number of cell types, although the mechanisms behind these observations are still unknown [16]. A matter of intense debate is the potential targeting of PI(4,5)P2 and other phosphatidylinositols into raft-like membrane domains [7]. The problem here is that the majority of natural PIP species have unsaturated acyl chains, thus the mode of their interactions with cholesterol would differ from the “umbrella model” proposed for SM-cholesterol membrane domains [17]. Another postulated mechanism for generating local enrichment of PI(4,5)P2 is via water-mediated hydrogen-bond networking through polar lipid head groups [18] (Fig. 2B). Such clusters can be disrupted by chaotropic agents, including monovalent salts at physiological concentrations [19]. Conversely, divalent cations, such as Ca2+, can trigger PI(4,5)P2 clustering within a membrane [20] (Fig. 2C). The observed condensing effect can be strong enough not only to change PIPs availability and presentation, but also to drive surface area mismatch between the two leaflets of the lipid bilayer, sufficient to induce membrane curvature [20]. Another example of cholesterol-modulated lipid presentation is observed with the ganglioside GM1. It has been shown that membrane cholesterol has the capacity to regulate glycolipid head group conformation and, thus, receptor function (Fig. 2D). Two prominent examples are the altered recognition of the GM1 head group by cholera-toxin [21] and the sterol-dependent binding of the Alzheimer's beta-amyloid peptide, APP, to GM1 [22]. Interestingly, fatty acid modification of GM1, resulting from the fluorescent labeling of this lipid, also changes its visibility to CTX-B [23], implying a contribution of the fatty acid chemistry to the GM1 head group presentation. Thus, as can be seen, lipids may be presented differently within the membrane, depending on their fatty

Please cite this article as: A. Czogalla, et al., Validity and applicability of membrane model systems for studying interactions of peripheral membrane proteins with lipids, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbalip.2013.12.012

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extreme cases, it can even lead to local formation of more disordered structures, such as hexagonal and cubic phases [26]. The twists and bends of undulating cell membranes are known to act as specific sites of recruitment for numerous curvature-sensing proteins containing such structural elements as amphipathic helices and BAR domains [27,28]. The most widely used in vitro model systems exhibit certain limitations (Fig. 3) which need to be considered not only in terms of understanding the data that they provide but also, more fundamentally, in terms of the validity and applicability of the model to be able to demonstrate the biological function of the membrane. Although model membrane systems are useful in rationalizing the nature of protein–lipid interactions, a “golden standard” of the in vitro membrane system does not exist. The employment of several model systems in an orthogonal manner to study individual phenomenon will help to establish a more holistic picture of biological significance. However, as exemplified above, it is proposed that there is a need to consider a number of factors so far neglected in the general experimental approach in this field. In particular, the dynamic and transient consequences of the membrane lipid composition in determining lipid presentation need careful consideration. In this review we shall provide an overview of the major experimental approaches for studying protein–lipid recognition processes currently employed to bypass the complexity of living cells. A concise summary of these membrane model systems is presented in Table 1. We shall attempt to evaluate them against criteria for the comprehensive understanding of the nature of lipid–protein interactions and to reconstitute a biological model using the minimal number of essential components, under standardized conditions. 2. Protein–lipid overlay assay Fig. 2. Examples of modulating lipid head group presentation. (A) Lipid head group charge dependency of phosphatidic acid (PA) (red) on the local lipid–lipid interactions by intramolecular hydrogen bonds with phosphatidylethanolamine (PE) (blue) [13]. (B) Electrostatic head group expansion effect of PI(4,5)P2 induced by monovalent chaotropic salts, such as NaCl, KCl and LiCl but also urea and temperature [19]. (C) Spatial distribution of phosphoinositides, PI(4,5)P2 under the influence of divalent cations, such as Ca2+ [20]. (D) Membrane cholesterol induced change of the GM1 head group conformation, resulting in formation of so-called “umbrella” configuration over the cholesterol and presentation of the ganglioside with direct consequences in molecular recognition by toxins and antibodies [21].

acid composition and/or their immediate membrane lipid surroundings. This directly implies that membranes can contain discreet pools of lipids with varying presentations that may be differently recognized by different proteins. A clear message emerging from the recent studies on protein–lipid interactions is the need to adopt appropriate models to determine the parameters of the membrane composition influencing the availability of bioactive lipid species that are required for the regulation and function of peripheral membrane proteins. Other membrane parameters can also influence protein binding. For example, dystrophin, a cytoskeleton protein vital for muscle function, directly associates with the membrane and is considered to mechanically regulate membrane stress during contraction and relaxation of muscle fibers. It has been demonstrated that lipidpacking and lateral membrane pressure have significant effects on the binding and organization of dystrophin. At low lipid surface pressures, the lipid binding of dystrophin is dependent on the accessibility of hydrophobic zones, whereas at high surface pressure, lipid binding is driven by electrostatic forces [24]. A particularly important natural feature of biological membranes is their asymmetry. Both the composition and biophysical properties of the two leaflets are different [25], which is usually difficult to challenge in synthetic model membrane systems. Membrane asymmetry may also contribute to curvature stress within the bilayer. In

One of the most widely used approaches to assess lipid-binding specificity of proteins is based on resolving the level of protein interaction with pure lipids immobilized at defined positions on nitrocellulose or PVDF [29,30], or with biotinylated lipids adhered to streptavidin-coated solid supports [31]. The major advantages of such systems are their convenience and relative high throughput, which facilitate the creation of miniaturized arrays capable of handling the lipid-binding profiles for a large number of proteins in a row [30,32]. However, the usage of lipid strips has major disadvantages. Most importantly, lipids are not presented to their recognition partners in the physiological context of the lipid membrane. Additionally, a homogenous distribution of lipids is often unattainable at the various individual spots because of the nature of the dispensing method itself. Moreover, this effect is exacerbated by the different water-solubilities of the lipid head groups, particularly of the phosphoinositides, that will result in a variable deposition efficiency and susceptibility to removal during the wash cycles [33]. Thus, it is suggested to qualitatively confirm the presence of bound lipids after completing the assay, such as by staining with molybdenum blue [34]. In general, protein–lipid overlay assays can be considered as a first-pass assessment only, with further confirmation using membrane-mimetic systems, as discussed below. 3. Lipid monolayers Lipid monolayers are films of single-molecule thickness spontaneously formed at the air/water interface after spreading of the lipid solution in an appropriate solvent mixture [35]. In the case of phospholipids, their hydrophobic tails are exposed to the air, while their hydrophilic head groups are embedded within the water phase. Structurally, such films resemble the single-leaflet of the lipid bilayer of natural membranes and are thermodynamically comparable to bilayers at the ‘so-called’ monolayer–bilayer equivalent pressure [36,37]. The

Please cite this article as: A. Czogalla, et al., Validity and applicability of membrane model systems for studying interactions of peripheral membrane proteins with lipids, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbalip.2013.12.012

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Fig. 3. Representative quality controls and properties of selected model membrane systems. (A) An example of TLC (silica 60, 5–6 μm) analysis of PC vesicles containing PI(4,5)P2 (asterisk) before (left) and after (right) treatment with high temperature/sonication resulting in partial decomposition of PI(4,5)P2. (B) A mica-deposited SLB labeled with Ld-phase preferring fluorescent lipid analog FastDiO; the used lipid composition of dioleoyl-PC/dipalmitoyl-PC/cholesterol (1/1/1 molar ratio) results in round shaped Lo-domains in free-standing membranes but are often difficult to reproduce on supported membranes because of non-specific interactions with the support. (C) GUVs composed of palmitoyl-oleoyl-PC/cholesterol/PI(4,5)P2 (60/35/5 molar ratio) labeled with membrane marker DiD (red channel, left); heterogeneity in lipid composition at the individual specimen level leads to heterogeneous binding of syndapin-GFP (green channel, right) to PI(4,5)P2 containing vesicles. (D) GPMVs derived from A431 cells using NEM blebbing procedure [146]; upon controlled cooling down to 4 °C, the majority of GPMV specimen exhibits phase separation (segregation visualized by Ld-phase marker FastDiI).

major advantages of this system are: homogeneity, stability, planar geometry and controllable lateral packing density of the lipids. Moreover, a useful feature of this method is that monolayers can also be formed with non-bilayer forming lipids in an aqueous environment. Typically, the monolayer approach is used to follow the penetration of proteins into a defined lipid surface [38]. When a membranepenetrating protein is injected into the subphase, the change in surface pressure (Δπ) is measured as a function of time, related to the protein concentration and lipid composition. As a control, the non-specific absorption of the protein at a bare air/water interface should also be taken into consideration in such studies [35]. Another very useful parameter to characterize lipid specificities is the maximum insertion pressure [39], also described as the exclusion pressure (πex) [38]. This parameter defines the upper limit of the initial surface pressure of the lipid monolayer (π0) into which a protein can penetrate, and can be estimated by an extrapolation of the plot depicting inverse proportionality of Δπ versus π0 . Given that the lateral pressure of cell membranes has been estimated in the range of 30–35 mN m− 1 [40], similar or higher πex values should characterize those peripheral proteins whose biological activity relies on membrane penetration. This method is faced with three important issues, namely that lipids are not presented in the context of a lipid bilayer, leaflet-coupling effects cannot be taken into consideration, and the method requires a relatively large amount of protein. An additional consideration is that this method is restricted to the study of planar lipid monolayers and, as such, is not amenable for the study of curvature-related effects. However, these systems may provide detailed thermodynamic analysis [35,41] and they are amenable to orthogonal measurements using a range of complementary techniques, such as Brewster angle microscopy and fluorescence microscopy. The conformation and orientation of monolayer lipids and proteins can also be extracted from polarized infrared spectra of such preparations [38]. Recent technical improvements have made the lipid monolayer approach accessible to more advanced techniques such as confocal imaging and fluorescence correlation spectroscopy [42,43], allowing the above data to be supplemented with spatial information about the binding of fluorescently-labeled proteins to discrete domains within monolayers. The stable Langmuir film can be transferred to a solid support, such as a mica plate or an electron-microscopy grid, by slow vertical lift of the slide from the subphase to air at constant surface pressure, according to the Blodgett deposition approach. This then allows further study of the same surface with scanning probes such as atomic force microscopy (AFM), or electron microscopy techniques,

which can reveal not only lipid domain structures but also peripheral protein organization at the monolayer as a function of surface pressure [24,44]. 4. Hybrid lipid bilayers The formation of hybrid lipid bilayers on solid supports usually involves the deposition of phospholipids on an electrode or chip surface previously coated with self-assembled monolayers of organic molecules, such as an alkanethiol (e.g. octadecanethiol) spontaneously absorbed to the support [45]. This yields a hybrid bilayer membrane (sometimes referred to as a hemi-lipid layer), where the upper, or distal, leaflet is composed of phospholipids and the lower leaflet, proximal to the support, consists of hydrophobic chains directly coupled to the sensor surface. This approach allows labelfree protein binding analysis using various techniques, such as surface plasmon resonance (SPR) [46], electrochemical impedance spectroscopy [47] and surface-enhanced Raman scattering [48]. A common phospholipid deposition method is based on the spontaneous fusion of small unilamellar vesicles to a hydrophobic surface. Employing cell membrane vesicles offers an opportunity to retain at least some of the membrane-embedded protein receptors [49]. Alternatively, a phospholipid monolayer from a Langmuir trough can be transferred directly onto the sensor surface from the air–water interface [50]. The biophysical properties of hybrid bilayers can be altered through modifying the functional surface using alkanethiols of various chain lengths and the additional introduction of ethylene oxide spacer units attached to these compounds. Hybrid phospholipid bilayers are usually more robust than fully supported lipid bilayers, due to the strong interactions they have with their support. Conversely, the same argument applies to some of the limitations associated with this membrane model, as the phospholipid monolayer is adsorbed onto another lipid monolayer that is fixed to a solid surface, resulting in a system of restricted fluidity and relatively high packing density. However, such systems have been reported to have utility in studies of membrane-disturbing proteins, such as antimicrobial peptides [51]. 5. Supported lipid bilayers More commonly used are supported lipid bilayers (SLBs) [52], where phospholipids are deposited as bilayers on hydrophilic substrates such as fused silica, borosilicate glass, mica or oxidized silicon [53]. One of the simplest means to form supported lipid bilayers is

Please cite this article as: A. Czogalla, et al., Validity and applicability of membrane model systems for studying interactions of peripheral membrane proteins with lipids, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbalip.2013.12.012

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Table 1 Overview of the most commonly used membrane model systems. Experimental approach

Controllable parameters

Major advantages

Major disadvantages

Protein–lipid overlay [33]

- lipid composition

- fast and easy to perform - relative high throughput

-

Monolayers [35]

- lipid composition - tunable lateral pressure

- defined geometry of lipid assembly (flat membrane) - homogeneity of the system - accessibility to fluorescence (confocal) microscopy, FCS, Brewster angle microscopy, AFM (after transfer to support)

- single leaflet presentation - requirement for relatively large amount of protein. - restricted to planar lipid monolayers

Supported lipid bilayers [53,54]

- lipid composition - incorporation of integral proteins/ compounds - membrane curvature/patterning

- flat geometry - asymmetric lipid distribution is amenable - accessibility to broad range of biophysical methods (e.g. AFM, TIRF, SPR) - accessibility of both leaflets (polymercushioned systems)

- interactions with support may result in restricted fluidity of lipids and segregation between the leaflets (partially overcame in polymer-cushioned systems) - possible defects within a bilayer

Pore-suspending membranes [86]

- lipid composition - incorporation of integral proteins/ compounds

- flat geometry - asymmetric lipid distribution is amenable - suitable for broad range of biophysical methods (particularly for conductance measurements, fluorescence microscopy) - accessibility of both leaflets

- reduced stability comparing to SLB - presence of residual organic solvents (primarily in BLMs)

Liposomes [89,93]

- lipid composition - membrane curvature/lipid packing (vesicle size-dependent) - incorporation of integral proteins/ compounds

- simple preparation procedure - free-standing membrane - compatibility with multiple methodical approaches

- possible polydispersity in terms of size and multilamellarity - symmetric lipid distribution - size below optical resolution - only one leaflet accessible

GUVs [116,142]

- lipid composition - membrane tension - membrane deformation can be induced (deflation, tubulation) - incorporation of integral proteins/ compounds

-

micrometer-scale structure free-standing membranes microscopically accessible compatibility with multiple methodical approaches

- heterogeneity in lipid composition at the individual specimen level - increased fragility compared to SUVs and LUV

-

relatively high population homogeneity micrometer-scale structure free-standing membranes complex lipid and protein composition microscopically accessible

- undefined lipid and protein composition - chemically induced crosslinking of lipids and proteins - possible depletion of specific lipid classes

GPMVs [146]

- protein content by overexpression

the absorption and fusion of small or large unilamellar vesicles (SUVs or LUVs, respectively) [53,54]. Vesicle adsorption is usually accelerated by addition of divalent cations such as calcium and magnesium. Subsequent fusion and rupture can also be enhanced either by heating, osmotic gradient or exposure to versatile fusogenic agents, such as polyethylene glycol.

lipid presentation in non-physiological context difficult to quantitate variable deposition efficiency susceptibility to removal during the incubation and wash cycles

The major drawbacks of liposomal deposition are excessive vesicles that need to be rinsed off and susceptibility to membrane defects [54]. Moreover, both the process of membrane deposition and the physical properties of the membrane strongly depend on ionic interactions between lipids and the support, such as the calcium-mediated interactions between phosphatidylserine (PS) and the mica support [55].

Please cite this article as: A. Czogalla, et al., Validity and applicability of membrane model systems for studying interactions of peripheral membrane proteins with lipids, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbalip.2013.12.012

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Alternatively, deposition of a lower leaflet by the Langmuir–Blodgett technique and the subsequent transfer of an upper leaflet by horizontal dipping the support to another monolayer in a Langmuir trough are possible [52]. The vesicle deposition approach can also be applied to proteoliposomes, and enables relatively easy incorporation of transmembrane proteins into SLBs, but at the expense that their orientation within the bilayer is difficult to control. A combination of the two approaches, where liposomes are fused with a pre-deposited phospholipid monolayer, yields the gentlest method to reconstitute integral membrane proteins into SLBs in an orientation-controllable way with respect to the bilayer [56]. This also enables the formation of asymmetric bilayers. A substantial improvement in the formation of highly uniform and defect-free supported membranes has been achieved by the spin-coating of lipids [57]. Planar SLBs can be also made from native cell membrane [58]. Such “membrane sheets” or “cell cortices” provide membranes of natural composition and may be valuable for reconstitution of dynamic membrane processes. Of particular note is that this technique enables the deposition of a cell-derived asymmetric plasma membrane where the cytosolic leaflet is orientated distally to the support, and thus accessible for further experimentation [59]. What distinguishes SLBs from hybrid lipid bilayers is a gap of around 1–2 nm in thickness between the SLB membrane and the supporting substratum that is occupied by a water phase. This phase is sufficient to lubricate the lower leaflet of the bilayer and increase the membrane fluidity, although some residual frictional coupling with the support makes the lipids within the leaflet proximal to the support diffuse slower [60]. Electrostatic or frictional coupling may also result in undesirable asymmetries in structural and compositional properties of SLBs. For instance, lipid domains of SLBs exhibiting Ld/Lo liquid phase separation differ in morphology from those observed in free-standing membranes (Fig. 3B) [61]. The hydration layer between the membrane and its support may also be insufficient to accommodate large extramembranous parts of integral proteins reconstituted into SLBs or retained within native membrane sheets. This may lead to the displacement or deformation of the proteins, potentially leading to their adsorption onto the support or even their denaturation. This problem can be alleviated in one of two ways. In the first instance, the introduction of spacer components between the substratum and the lipid bilayer, such as through tethering SLBs to a support. Such an example is the use of ligand–receptor pairs, where membranes bearing biotinylated lipids are captured to a low-density streptavidin sublayer [62]. In the second approach, the deposition of a polymeric cushion directly onto the support can be used as the basis for creating an environment that maintains membrane fluidity. Among several types of polymers explored as cushion supports, two major polymer classes appear to be of particular interest: polyelectrolytes, such as polyethylenimine, and lipopolymers [53,56,63–65]. Lipids with head groups modified with a thiol group linked via a hydrophilic polymer are often employed to attach lipid bilayers to gold sensors used in SPR [66]. The most common examples are polyethylene glycol (PEG)-based polymers that bridge a phospholipid on one end and a support-tethering silane group on the other end of the molecule [67]. Free diffusion of lipid components within the bilayer is preserved when the concentrations of the lipopolymer in the proximal leaflet do not exceed a critical mol% of the total membrane lipids [56,68]. The polymer-supported approach is also suited to generate membranes with asymmetric lipid composition built via layer-by-layer deposition, allowing the study of membrane domain-registration and leaflet-coupling phenomena [69]. Supported lipid bilayers provide an elegant approach to control selected properties of membranes coupled to structured surfaces exhibiting chemical and/or topological patterns [70]. A wide array of techniques is available to fine-tune the architecture and biophysical parameters of SLBs for optimal reconstitution and presentation of membrane proteins [65].

An exciting new field relates to studies on protein–membrane interactions involving curvature-sensing mechanisms using SLBs. A robust way to generate membranes with highly uniform curvatures is provided by small microparticles or beads (3–30 μm in diameter) of various compositions as a support for lipid bilayers. The unique properties of such systems allow membrane-coated beads to be readily mixed with water-soluble proteins and for the beads to be then captured in laser traps, or for their colloidal properties to be exploited to probe proteinbinding events at membrane surfaces [71]. The accessibility of SLBs is advantageous for applying a broad array of biophysical methods, including AFM [72], total internal reflection fluorescence microscopy [73], attenuated total reflectance [74] and surface plasmon resonance [75], to name but a few. These applications support the study of not only kinetics of protein–lipid interactions [46], but also changes in lipid-chain order [76], lipid-driven structural transitions of proteins [77], lateral diffusion of proteins [78] and lipids [79], oligomerization and self-assembly of proteins on membrane surfaces [80,81], membrane fusion [81] and pore formation within the lipid bilayer [82,83]. Nevertheless, it must also be stressed that the underlying support will always have some influence on the structural and functional properties of the lipid bilayer and lipid presentation. Although these undesirable effects can be minimized to various extents, it is not possible to fully exclude them using the approaches described above. Another disadvantage of SLBs is the relatively poor membrane integrity due to the frequent presence of defects in the bilayer. 6. Pore-suspending membranes Attempts to eliminate the effect of support while, at the same time, preserving the experimental advantages of SLBs, such as stability, optical symmetry and ease of buffer exchange, have led to the development of systems with membranes suspended over open apertures. The use of a microstructured silicon or thiol-functionalized gold support for studying pore-suspending membranes has been reported, enabling parallel real-time kinetic studies of individual membrane transporters or the study of oligopeptide permeation through phospholipid bilayers [84]. On the other hand, apertures larger than 1 μm should provide access to free-standing regions of a membrane for optical approaches. This can be achieved either by Langmuir–Blodgett transfer of lipid monolayers to a grid [85] or by electrostatically enhanced bursting of giant unilamellar vesicles on functionalized silicon nitride grids [86]. Such systems share the general concept with black lipid membranes (BLMs), where a free-standing lipid bilayer is formed by painting of the lipid solution in an organic solution over a large aperture (up to 1 mm) in a hydrophobic material [87]. As the membrane usually separates two aqueous compartments that can be equipped with electrodes, BLMs are particularly used to investigate ion-channel activity. However, this system is usually characterized by poor stability and the presence of residual organic solvents. In particular, the shorter alkanes that are sometimes used in these methods may partially reside in the BLM, which results in potential solvent-dependence artifacts interfering with the diffusion coefficients of lipids in membrane [88]. 7. Liposomes Liposomes are aqueous-filled structures of lipid bilayers spontaneously formed during dispersion of phospholipids in water and have been extensively used over the years for membrane-binding assays. Some lipid mixtures do not spontaneously form vesicles consisting of bilayers: in particular those with high cholesterol content (N 50%), high amount of charged lipids, or lipids with a packing parameter (P = v / al; where v—molecular volume, a—cross-sectional area of the head group; l—length of the molecule) far smaller or greater than 1.0 (thus having conical rather than cylindrical molecular shape), which tend to form micelles, hexagonal, or cubic phases [26].

Please cite this article as: A. Czogalla, et al., Validity and applicability of membrane model systems for studying interactions of peripheral membrane proteins with lipids, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbalip.2013.12.012

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Liposomes are usually classified according to their size and lamellarity, that is the number of lipid bilayers that make up an individual vesicle [89]. Multilamellar vesicles (MLVs) are rarely used for quantitative binding assays, since most of the formed bilayers are not accessible to membrane-binding molecules and their geometry is undefined. On the other hand, small (SUVs; d b 100 nm) and large (LUVs; d N 100 nm) unilamellar vesicles are relatively well-defined and easy to prepare. Among various preparation methods [89], the simplest and most extensively used is hydration of a dry thin lipid film, yielding a heterogeneous population of liposomes in terms of size and lamellarity. Further sonication of the suspension results in the formation of SUVs, with their average size-distribution highly dependent on molecular shape and charge of the lipids used. For natural glycerophospholipids, the minimum diameter of thermodynamically favored vesicles is approximately 20 nm [90]. More precise size calibration is usually achieved by applying a number of cycles of freezing and thawing and subsequent extrusion of liposomes through a polycarbonate membrane of a defined pore-size. Besides lipid composition, size is one of the most critical parameters influencing membrane–protein interactions. Decreasing the diameter of liposomes imposes increased but opposed spacing constraints at both the head group level and the tails of the individual leaflet constituents of the liposomes. In membranes containing at least two lipids with opposing molecular shapes, these constrains might lead to spontaneous asymmetric distribution of lipids across the membrane according to their packing parameters. A number of techniques have been adapted to measure the size of liposomes, in particular, light scattering and electron microscopy [91]. Both of these techniques are also commonly used to assay protein-driven tubulation and fragmentation of liposomes [92]. A possible problem of the lipid film hydration method is that within the population of heterogeneous liposomes a noticeable fraction consists of stable small vesicles. The latter are unaffected by extrusion through filters with pore diameters N100 nm, resulting in a polydispersed population [93]. Thus, measurements using heterogeneous vesicle populations may influence experimental results, particularly if the smaller vesicle population has a significant and specific effect of its own. The problem of differential responses observed across heterogeneous vesicle populations has been recently raised for curvature-selective binding of amphipathic proteins to individual liposomes, where both vesicle size and protein recruitment were measured via quantitative fluorescence microscopy [94]. The simplicity of the liposomal system triggered its widespread use in studying protein–membrane interactions. A broad array of methods is based on the sensitivity of tryptophan fluorescence to its microenvironment [95] and fluorescence resonance energy transfer (FRET) [96]. The latter approach often includes the chemical labeling of proteins and lipids for a variety of methods. The standard chemical labeling protocols for proteins, however, make use of probes that are linked either to lysines or free cysteines. Positively charged lysine residues are often present in the binding interface of proteins and their modification may, therefore, change their binding specificity and affinity. Additionally, an avidity effect is observed where initially low binding-affinities are amplified through oligomerization of proteins at the membrane [97,98]. Binding of fluorescently modified proteins to liposomes can be also investigated by fluorescence-correlation spectroscopy, where the membrane partitioning coefficient is extracted from the multi-component autocorrelation function [99]. Direct, label-free quantification of membrane–ligand binding can be achieved by detecting slight changes in the refractive index of a solution via backscattering interferometry [100]. In some cases common principles underlying separation are used for measuring the liposome-bound protein fraction, which is usually directly performed through dialysis [101]. The major disadvantage of this approach is the long time of equilibration (up to 24 h). More convenient are the methods based on centrifugation, such as sedimentation of sucrose- or dextran-loaded vesicles after incubation

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with a protein [102,103]. Although extensively employed, the possibility of perturbation of the equilibrium between bound and free protein fractions following extended periods of centrifugation needs to be considered. Another issue is a possibility of protein precipitation during centrifugation. A solution for the latter problem may involve the flotation of vesicles in a sugar gradient, rather than their sedimentation [104]. Liposomes can also be immobilized on a solid support of a microarray, or a sensor. Arrangements of surface-immobilized vesicles prepared through such links as oligonucleotides covalently coupled to cholesterol [105] or specific streptavidin–biotin binding [106], may be used for fastthroughput screening of membrane-interacting compounds. Moreover, SPR may be directly applied to vesicles immobilized on a sensor for measuring protein–membrane interactions. This can be achieved either by direct attachment of unmodified liposomes to hydrophobic anchors deposited on a chip, or via ligand–receptor pairing [46]. The integrity of immobilized liposomes is usually preserved, as confirmed in fluorescence-leakage experiments [107]. 8. Giant unilamellar vesicles The attractiveness of giant unilamellar vesicles (GUVs) as membrane model systems derives mostly from their geometry, as they are cellsized vesicles approximately 10–100 μm in diameter. Consequently, they are easily accessible to either standard contrast enhanced microscopy or fluorescence microscopy. GUVs provide well-characterized membranes that are completely free of any kind of supporting surface. This is reflected by unconstrained diffusion within the plane of the bilayer [108]. The simplest and, historically, earliest method of GUV formation is based on gentle hydration of a dry lipid film [109]. Although the addition of charged phospholipids is necessary to provide electrostatic repulsion between bilayers, guaranteeing sufficient yield and quality of vesicles, this approach allows the preparation of GUVs at physiological ionic strength [110]. Nowadays, the most widely used method to produce GUVs is the electroformation of vesicles. This affords the opportunity to produce GUVs more rapidly with higher reproducibility and yield [111,112]. The alternating current (AC) exerts an ordering effect on the movement of lipid molecules deposited on electrodes, such as ITOcovered microscopic slides or platinum wires. This helps the lipids to adopt bilayer-packing and to arrange into unilamellar structures. Depending on electric field conditions, the vesicles can remain attached to the electrodes, allowing long-term observations and mechanical manipulations, or can be fully detached into solution. The simplest protocols delimit formation conditions to buffers of low ionic strength and a relatively low content of charged lipids in membranes, usually not exceeding approximately 20% of the total lipid component [113]. However, tight control of the electric-field parameters, such as voltage and/or frequency, facilitates electroformation at physiologically relevant electrolyte concentrations using synthetic mixtures containing negatively charged lipids, natural lipid extracts or preformed proteoliposomes containing integral membrane proteins [114,115]. These methods are used routinely, although many factors that strongly influence the quality of the vesicles, such as temperature and electrodecatalyzed oxidation and decomposition of lipids, need to be taken into consideration [116]. One of the major issues concerning the use of GUVs is the limited control of the membrane lipid composition of individual vesicles, also reflected in selective binding of proteins to vesicles within one population (Fig. 3C). Thus, in many cases, interesting phenomena are observed on a fraction of total vesicle populations selected from a group that, on average, may not show distinctive effects. Most recently, efforts have focused on the development of new techniques to produce GUVs, such as the utilization of water/oil emulsion-transfer [117] or microfluidic jetting [118], which allow the precise control of lipid composition, trans-bilayer asymmetry,

Please cite this article as: A. Czogalla, et al., Validity and applicability of membrane model systems for studying interactions of peripheral membrane proteins with lipids, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbalip.2013.12.012

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as well as internal contents of vesicles. However, the presence of residual organic solvents in the GUV bilayer is one of the major disadvantages of these methods. The microscopic accessibility of GUVs triggered studies on the formation and properties of membrane lipid-phases (liquid-disordered (Ld), liquid-ordered (Lo) or gel) and the phase partitioning properties of molecules [119–122]. This combination of tools has also enabled investigating the role of individual lipid species in membrane recruitments of proteins [113] as well as tracing the preferential association of proteins to coexisting Ld/Lo lipid-phases and the interplay between lipid organization and protein activity [123,124]. Small vesicles usually will not provide an insight into reactions occurring within or on the surface of an individual vesicle, since only ensemble-averaged observations are possible with such structures. On the other hand, since GUVs act as freestanding membranes, in contrast to SLBs, the processes of membrane-induced oligomerization of proteins as well as protein-driven lipid clustering are easier to observe and recognized as more physiologically relevant [113,125]. Potential destabilization of GUVs as a result of clustering events and/or protein oligomerization can be easily observed with fluorescence microscopy in a real-time regime. The different responses can be classified as membrane fusion, budding, or tubule formation, which give information on the modes of action of proteins or protein assemblies on lipid bilayer [126,127]. Incubating GUVs with cytosolic fractions and selectively fluorescenttagged proteins makes it possible to reconstitute and trace even highly complex multicomponent processes, such as COPII membrane scaffolding and remodeling [128], clathrin-AP-1-coated membrane carrier biogenesis [129] or formation of multivesicular bodies mediated by endosomal sorting complexes required for transport (ESCRT) [130]. As recently demonstrated, sequential recruitment of protein complexes to GUVs, along with membrane-assisted activation and vesicle rupture, can be assayed with flow cytometry [131]. Membrane-leaking experiments on GUVs allow for simultaneous microscopic observations of protein binding, membrane permeabilization and vesicle integrity. This provides additional information about heterogeneities within membrane structure and some insights into the mechanical aspects of membrane pore formation [132,133]. Another important aspect of GUVs as a model system is the possibility to control membrane tension, which may influence protein binding and/or protein organization on the surface of membranes [134]. This parameter can be adjusted relatively easy by changing the osmotic gradient across the membranes or can be more precisely controlled with micropipette aspiration [135]. GUVs are also suitable for generating curved membranes in a controllable way by motor proteins attached to the vesicular membrane reservoir via polystyrene beads and biotin–streptavidin recognition pair [136,137]. Nanotubes pulled from vesicles closely represent various stages of tubular carrier formation in cells. More precise control of nanotube formation can be obtained by pulling nanotubes from micropipette-aspirated GUVs with a bead trapped by an optical tweezers [138,139] or by capillary aspiration [140,141]. Such systems are powerful in studying spatial distribution of membraneattached proteins or protein-assisted lipid sorting between flat and curved regions. Another application is monitoring concentrationdependent functional switching between membrane curvaturesensing and curvature-generating processes by peripheral membrane proteins. Moreover, this set of tools enables valuable insights into details of the action of mechano-enzymes, such as the membrane fission-triggering molecule, dynamin [142]. 9. Giant plasma membrane vesicles As GUVs are limited in their capacity to reconstitute or represent the compositional diversity of cellular membranes, one consideration is to induce cell swelling by nutrient depletion in order to enforce the

generation of single-cell plasma membrane spheres [143]. Another, more frequently used method, is the use of chemically-induced production of giant plasma membrane-derived vesicles (GPMVs) [144]. The most common GMPV preparation method involves the treatment of cells with a combination of chemical cross-linkers and reducing agents such as paraformaldehyde (PFA) and dithiothreitol (DTT) [145]. Alternative protocols that avoid chemical crosslinkers involve N-ethyl maleimide (NEM) to induce cell blebbing [146]. Similarly, formation of vesicles from the plasma membrane of living cells can be triggered by their exposure to hypotonic conditions [147]. The cellular events that induce membrane vesiculation are complex and not well understood. Moreover, the resulting membrane composition and asymmetry of the bilayer are usually compromised to some extent when compared to cells they derive from [145,148]. It should also be noted that treatment with PFA/DTT greatly affects the lipid composition of the resultant vesicles, depleting them almost completely of some lipidic species, such as PE and phosphoinositides [146,148]. In this respect the usage of NEM seems much more gentle since, at least, the levels of major membrane phospholipids are unaffected, although, most probably, these vesicles will also be depleted of phosphoinositides [146]. Apart from these limitations, GPMVs appear to be a promising tool that may serve to bridge the data obtained using fully artificial systems and the observations from cells. Studies on plasma membrane vesicles have provided meaningful data for lipid-phase coexistence in natural membranes (Fig. 3D) and selective partitioning of lipid-modified proteins in more ordered phases [23,149]. Particularly informative in the former case is the labeling of the GPMVs with membrane-penetrating dyes that are sensitive to water content, from which the relative packing of the bilayer can be extrapolated [150]. It appeared that in plasma-membrane derived vesicles, membrane-domains exhibit properties different to liquid disordered/liquid ordered (Ld/Lo) phases in synthetic systems (e.g. GUVs), where enhanced order difference is observed [23]. Here, data on selective binding of peripheral proteins to pre-existing membrane domains of GPMVs and their influence on the formation and coalescence of such domains are of particular interest, such as the clustering of lipid ligands [23,143]. 10. Conclusions The complexity of cellular membranes and their behavior as collectives are encoded within the biochemical diversity of membrane lipid composition itself. It is this biochemical diversity, which imposes the biophysical properties of the membrane. While a number of important methods exist for determining lipid behavior in model systems, it is becoming increasingly clear that reconstitution of specific membrane/lipid–protein interactions using synthetic membrane model systems needs to account for the functional influence of the biochemical complexity on lipid presentation. The molecular modulation of lipid presentation at the biochemical level ultimately manifests itself as specific membrane-level effects in terms of physicochemical properties that constitute the various membrane compartments, such as the plasma membrane and the various internal membrane components, such as the ER and Golgi. The over-simplification imposed by the limitations of our current range of model systems may not allow the expression of the full spectrum of possible regulatory mechanisms needed to demonstrate function. Indeed, it is becoming increasingly evident that they may not even be able to support the biological behavior of the processes that they are being used to study in some instances. Consideration must be given to both validating lipid chemical identity by quality controls (e.g. TLC, MS, see Fig. 3A), and achieving appropriate conditions for lipid presentation (e.g. buffer conditions, temperature, membrane morphology, integrity etc.). These two criteria are poorly accommodated in our current methodologies. It is recommended that the community approaches this area with standardization

Please cite this article as: A. Czogalla, et al., Validity and applicability of membrane model systems for studying interactions of peripheral membrane proteins with lipids, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbalip.2013.12.012

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in mind, before considering whether they are able to demonstrate natural physiological behavior. The long-term aim should be to improve these methods so as to develop superior protocols that can take account of the subtle modulation of the membrane at the level of controlling lipid presentation. Here, we offer a minimum set of standardized quality criteria, which should be included as part of any experimental rationale designed to study dynamic mechanisms involving the interaction of peripheral membrane proteins with the membrane itself. The initial step of introducing an appropriate level of quality control of lipid composition is essential, so that the true lipid composition within the model system itself is validated and quantified. It is then recommended that the influence of the experimental conditions themselves on lipid presentation is evaluated in terms of vesicle type, size and membrane morphology. Finally, it is necessary to determine whether the method itself satisfies the following requirements with respect to: i) Lipid composition — Is it possible to validate that the composition of the lipid in the synthetic model truly reflects the minimum requirements to demonstrate a physiological effect in vitro? ii) Lipid presentation — Is it possible to modulate the presentation of the lipid to membrane proteins, specifically the hydrophilic head group together with the fatty acid chain, within its local environment in the model system? iii) Independent assay validation — The validation of the model via an orthogonal measurement, whenever possible. Acknowledgements This work was supported by the following agencies: Deutsche Forschungsgemeinschaft (DFG) “Transregio 83” Grant TRR83 TP18 and by the German Federal Ministry of Education and Research (BMBF) grant to the German Center for Diabetes Research (DZD e.V.). References [1] R.M. Bell, L.M. Ballas, R.A. Coleman, Lipid topogenesis, J. Lipid Res. 22 (1981) 391–403. [2] U. Coskun, K. Simons, Membrane rafting: from apical sorting to phase segregation, FEBS Lett. 584 (2010) 1685–1693. [3] R. Behnia, S. Munro, Organelle identity and the signposts for membrane traffic, Nature 438 (2005) 597–604. [4] G. van Meer, D.R. Voelker, G.W. Feigenson, Membrane lipids: where they are and how they behave, Nat. Rev. Mol. Cell Biol. 9 (2008) 112–124. [5] G. van Meer, A.I. de Kroon, Lipid map of the mammalian cell, J. Cell Sci. 124 (2011) 5–8. [6] G. Di Paolo, P. De Camilli, Phosphoinositides in cell regulation and membrane dynamics, Nature 443 (2006) 651–657. [7] K. Kwiatkowska, One lipid, multiple functions: how various pools of PI(4,5)P(2) are created in the plasma membrane, Cell. Mol. Life Sci. 67 (2010) 3927–3946. [8] J. Saarikangas, H. Zhao, P. Lappalainen, Regulation of the actin cytoskeleton–plasma membrane interplay by phosphoinositides, Physiol. Rev. 90 (2010) 259–289. [9] P. Mayinger, Phosphoinositides and vesicular membrane traffic, Biochim. Biophys. Acta 1821 (2012) 1104–1113. [10] M.A. Lemmon, Membrane recognition by phospholipid-binding domains, Nat. Rev. Mol. Cell Biol. 9 (2008) 99–111. [11] T.G. Kutateladze, Translation of the phosphoinositide code by PI effectors, Nat. Chem. Biol. 6 (2010) 507–513. [12] A. Balla, G. Tuymetova, A. Tsiomenko, P. Varnai, T. Balla, A plasma membrane pool of phosphatidylinositol 4-phosphate is generated by phosphatidylinositol 4-kinase type-III alpha: studies with the PH domains of the oxysterol binding protein and FAPP1, Mol. Biol. Cell 16 (2005) 1282–1295. [13] E.E. Kooijman, K.M. Carter, E.G. van Laar, V. Chupin, K.N. Burger, B. de Kruijff, What makes the bioactive lipids phosphatidic acid and lysophosphatidic acid so special? Biochemistry 44 (2005) 17007–17015. [14] Z.T. Graber, Z. Jiang, A. Gericke, E.E. Kooijman, Phosphatidylinositol-4,5-bisphosphate ionization and domain formation in the presence of lipids with hydrogen bond donor capabilities, Chem. Phys. Lipids 165 (2012) 696–704. [15] F.R. Maxfield, G. van Meer, Cholesterol, the central lipid of mammalian cells, Curr. Opin. Cell Biol. 22 (2010) 422–429. [16] D.M. Taglieri, D.A. Delfin, M.M. Monasky, Cholesterol regulation of PIP(2): why cell type is so important, Front. Physiol. 3 (2012) 492. [17] D. Lingwood, K. Simons, Lipid rafts as a membrane-organizing principle, Science 327 (2010) 46–50.

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Please cite this article as: A. Czogalla, et al., Validity and applicability of membrane model systems for studying interactions of peripheral membrane proteins with lipids, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbalip.2013.12.012