Bioactive Structure of Membrane Lipids and Natural Products Elucidated by a Chemistry-Based Approach.

Personal Account DOI: 10.1002/tcr.201402097

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Bioactive Structure of Membrane Lipids and Natural Products Elucidated by a Chemistry-Based Approach Michio Murata,*[a,b] Shigeru Sugiyama,[a,b] Shigeru Matsuoka,[a,b] and Nobuaki Matsumori[b,c] [a] JST ERATO, Lipid Active Structure Project, Machikaneyama, Toyonaka, Osaka 560-0043 (Japan), E-mail: [email protected] [b] Department of Chemistry, Graduate School of Science, Osaka University, Machikaneyama, Toyonaka, Osaka 563-0043 (Japan) [c] Present address: Department of Chemistry, Faculty and Graduate School of Sciences, Kyushu University, 744 Motooka, Nishi-ku, Fukuoka 819-0395 (Japan)

Received: December 9, 2014 Published online: June 30, 2015

ABSTRACT: Determining the bioactive structure of membrane lipids is a new concept, which aims to examine the functions of lipids with respect to their three-dimensional structures. As lipids are dynamic by nature, their “structure” does not refer solely to a static picture but also to the local and global motions of the lipid molecules. We consider that interactions with lipids, which are completely defined by their structures, are controlled by the chemical, functional, and conformational matching between lipids and between lipid and protein. In this review, we describe recent advances in understanding the bioactive structures of membrane lipids bound to proteins and related molecules, including some of our recent results. By examining recent works on lipidraft-related molecules, lipid–protein interactions, and membrane-active natural products, we discuss current perspectives on membrane structural biology. Keywords: fatty acids, lipid rafts, membrane proteins, NMR spectroscopy, sphingolipids

1. Introduction Biomembranes are regarded as the last frontier and a main bottleneck of life science research. In particular, heterogeneous bilayer systems with distinct outer and inner leaflets are extremely complicated molecular assemblies comprising a variety of lipids and diverse associated proteins, which cannot be reproduced by simply reassembling lipids and proteins (Figure 1). To gain deeper insight into cellular functions, a better understanding of biomembranes is crucial, but this has been very difficult to achieve with current technologies, which have been historically developed for primary biopolymers (DNA, RNA, and proteins) without much focus on secondline constituents such as lipids and carbohydrates.

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Until the late 1990s, the fluid mosaic model proposed by Singer and Nicolson prevailed,[1] in which plasma membranes were considered to be two-dimensional homogeneous fluids with free diffusion of lipids and proteins. However, the heterogeneity and complexity of biological membranes have been widely recognized since the lipid raft hypothesis was proposed by Simons and Ikonen,[2] which makes membrane research even more challenging; it was revealed that the lipid bilayers of plasma membranes are assemblies of microdomains consisting of different lipid species (Figure 1). Fortunately, recent research has demonstrated that the basic physicochemical features of lipid rafts can be partly reproduced using artificial

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membranes without proteins,[3] which has greatly accelerated biophysical studies to elucidate the atomistic mechanism of lipid–lipid interactions in membranes.[4] In contrast to soluble proteins, progress in understanding the structural biology of membrane proteins has been slow, largely because of difficulties in functional expression and crystallization. Although they constitute nearly 30% of the human genome, membrane proteins represent only 2.6% of protein structures registered in the Protein Data Bank. Integral membrane proteins usually form complexes with annular lipids, and lipid binding is thought to influence protein function, whereas loosely bound lipids are largely removed from protein surfaces and displaced with detergents during purification. Thus, detailed structural and functional information on the original annular lipids must be accumulated to accurately reproduce the physiological environment of integral membrane proteins. The X-ray crystal structure of the human b2 adrenergic receptor (Figure 2), a member of the GPCR family, includes two tightly bound cholesterol (chol) molecules co-crystallized

with the receptor.[5] The activities of other GPCRs, including the oxytocin and serotonin receptors, are affected by membrane chol, which dramatically increases agonist affinity.[6] However, chol is reported to stabilize the inactive state of rhodopsin, another GPCR member, through indirect effects on plasma membrane curvature.[7] Membrane lipids also influence ion channel proteins. For example, the potassium channel KcsA from Streptomyces lividans binds tightly to anionic phospholipids, which are thought to be necessary for proper channel function.[8] Despite relatively high-resolution X-ray structures for KcsA, the precise lipid-binding sites have not been determined. These examples clearly indicate that membrane lipids/sterols are not merely media incorporating receptor proteins but also important constituents of physiological membrane receptor complexes. Except for lipids strongly binding to proteins, however, conventional strategy alone may not suffice to examine the structures and functions of lipids in biomembranes. A possible approach may be imaging modalities and molecular dynamics (MD) simulations, which have provided a clearer picture of

Michio Murata graduated from Tohoku University, and joined the Suntory Institute for Bioorganic Research (director: Koji Nakanishi). He returned to Tohoku University and obtained his Ph.D. in 1986 (adviser: Prof. Takeshi Yasumoto). In 1993, he moved to the University of Tokyo (Prof. Kazuo Tachibana) as an associate professor. In 1999, he was promoted to a professor at the Graduate School of Science, Osaka University. His research interest is on the structure and function of flexible organic molecules including lipids.

versity in St. Louis as a postdoctoral fellow (adviser: Prof. J. Schaefer), he joined the Graduate School of Pharmaceutical Sciences, the University of Tokyo, as an assistant professor. He is currently an associate professor at the Graduate School of Science, Osaka University, and a group leader in the JST ERATO Murata Lipid Active Structure project.

Shigeru Sugiyama graduated from the Graduate School of Engineering, Himeji Institute of Technology, in 1992 (adviser: Prof. N. Yasuoka). He received his Ph.D. from Osaka University in 1997 (adviser: Prof. Y. Kai). He joined the Kyowa Hakko Kogyo Co., Ltd. Presently, he is an associate professor at the Graduate School of Science, Osaka University, and a group leader in the JST ERATO Murata Lipid Active Structure project.

Nobuaki Matsumori received his B.Sc. (1992) and his Ph.D. (1997) from the University of Tokyo under the supervision of Prof. K. Tachibana. His Ph.D. research was on the development of a Jbased configuration analysis method. After two years of postdoctoral research in antibiotic biosynthesis with Prof. S. Horinouchi at the University of Tokyo, he was appointed as an assistant professor (1999) and then an associate professor (2010) at Osaka University, where he worked with Prof. M. Murata. In 2000, he spent a year in the solid-state NMR laboratory of Prof. R. G. Griffin at MIT. In 2014, he was promoted to a professor at Kyushu University. His research centers on the integrated analysis of membrane systems, including membrane proteins and lipid rafts.

Shigeru Matsuoka graduated from the University of Tokyo in 1997 (adviser: Prof. K. Tachibana). He received his Ph.D. from Osaka University in 2003, followed by two years of postdoctoral training (adviser: Prof. M. Murata). After spending two years at Washington Uni-

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Fig. 1. Cellular membrane with lipid rafts and fatty acid binding protein (blue) in cytosol.

intermolecular interactions of lipids and other membrane components in biomembranes. Another potential approach is subdivision of the membrane assemblies into molecular components that permit application of structural biological and/or biophysical methodologies. This approach focuses on examining the lipid–lipid interactions in the membrane and the annular lipids associated with integral membrane proteins. This lipid-based strategy for investigating biomembranes sounds

promising (Figure 3), but there are some difficult issues: lipid– lipid and lipid–protein assemblies are usually very dynamic without clear periodic structures and are therefore rarely subjected to conventional structural biology methods such as Xray and solution NMR spectroscopy. More importantly, the experimental protocols to deal with lipids and other highly hydrophobic ligands are not completely established in structural biology; dissolving both lipids and proteins in aqueous

Fig. 2. X-ray structure of human b2 adrenergic receptor.[5] The picture was obtained from the 2.8 A˚ resolution crystal structure of the thermally stabilized receptor bound to cholesterol and the partial inverse agonist timolol. The figure was reproduced from reference [5] with permission.

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Fig. 3. Three areas of active lipid structure investigation required for a better understanding of biomembranes. Research on these topics must establish methods for handling highly hydrophobic ligands such as water-insoluble lipids for structural biology experiments, including X-ray crystallography, NMR, and evaluation of lipid–protein interactions.

media is still one of the most serious problems in X-ray and NMR experiments. In this review, we describe recent findings on the bioactive structures of membrane lipids and related molecules mainly based on our recent results. Through our discussion of lipidraft-related molecules, lipid–protein interactions, and membrane-active natural products, we endeavor to provide a clear perspective on the future directions of lipid structural biology.

2. Lipid-Raft-Related Molecules, Conformation, and Dynamics 2.1. Lipid Rafts and Sphingomyelin Sphingomyelin (SM), the most abundant sphingolipid in biomembranes, forms ordered microdomains called lipid rafts with chol in plasma membranes (Figure 4).[2] The lipid rafts are segregated from outer disordered bilayers composed of unsaturated phosphatidylcholine (PC) and are characterized by tight packing of lipid molecules despite their relatively high lateral mobility. Lipid rafts are also believed to play a key role in various cellular processes such as signal transduction, protein sorting, and chol shuttling.[2,9] Furthermore, lipid rafts are recognized as potential sites for toxin interactions, pathogen entry,[10] and fusion of influenza viruses and human immunodeficiency virus[11] and have therefore attracted much attention from multidisciplinary researchers. Despite such biological and

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Fig. 4. (a) Schematic representation of lipid rafts. (b) Structures of sphingomyelin (SM) and cholesterol.

pathological significance, the manner in which raft-forming lipids self-assemble to form ordered domains is still unclear, largely due to the dynamic properties of the lipid molecules in rafts. To address this question, we have explored the structures, dynamics, and interaction of SM and chol in lipid rafts by combining various NMR techniques with organic synthesis of labeled SMs, which led to some findings on the conformation, orientation, and dynamics of SM in membranes. 2.2. Conformation of Sphingomyelin in Isotropic Bicelles To date, the conformation of SM in membrane environments has not been elucidated, but its conformation in organic solvents was extensively analyzed based on coupling constants from NMR experiments.[12] Bicelles provide a closer mimic of the membrane environment than micelles since bicelles possess true lipid bilayer portions.[13] Bicelles are generally comprised of a long-chain phospholipid such as dimyristoyl phosphatidylcholine (DMPC) and a detergent such as dihexanoyl phosphatidylcholine (DHPC, Figure 5a,b) and adopt a disk-shaped morphology in which the planar bilayer region consisting of the long-chain lipid is surrounded by a rim composed of the detergent. The size and shape of bicelles can be controlled by the q value, the ratio of long-chain phosphatidylcholine to detergent. At higher q values, bicelles are oriented in an external magnetic field, whereas bicelles with q values less than one undergo fast tumbling and can be regarded as isotropic in aqueous solutions, thus allowing high-resolution 1H NMR measurements.[14] We noticed some similarity between lipid rafts and bicelles: lipid rafts were originally defined as detergentresistant membranes, and the bilayer membranes in bicelles are also insoluble to the surrounding detergent. Hence, under the



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Fig. 5. (a) Schematic structures of bicelles composed of dimyristoyl phosphatidylcholine (DMPC) and dihexanoyl phosphatidylcholine (DHPC). (b) Oriented (left) and isotropic (right) bicelles with respect to the external magnetic field, B0. (c) 2H NMR spectrum of stearoyl-SM/100 -d2-stearoyl-SM (SSM)/DHPC (3:1:1) bicelles at 37 C, suggesting the formation of magnetically oriented bicelles. Total lipid concentration: 500 mM. (d) Conformation of the central region of SM in small isotropic bicelles (SSM/DHPC 5 1:2) deduced from 1H NMR spin–spin coupling constants (blue arrows) and NOEs (red arrows).[15,17]

assumption that bicelles can be used as a lipid raft model, we applied the bicelle technique to the structure analysis of SM in a membrane environment.[15] Because there was no preceding study on the formation of SM/DHPC bicelles, we first showed that the SM/DHPC system forms both oriented and isotropic bicelles as for the well-established DMPC/DHPC bicelles, by synthesizing deuterated stearoyl-SM (100 -d2-SSM, Figure 5c) and measuring the 2H NMR spectrum of 100 -d2-SSM/DHPC bicelles (q 5 4), which gave a well-resolved doublet without any center peak (Figure 5c).[15] Lipid molecules residing in the bilayer portion of oriented bicelles provide sharp doublet signals in 2H NMR spectra, while those in the rim portion of oriented bicelles show peaks near the spectral center because of the isotropic nature of the rim portion (Figure 5).[16] We also utilized small isotropic bicelles composed of SSM and DHPC (q 5 0.5) to determine the conformation of SM in isotropic bicelles on the basis of nuclear Overhauser effects and vicinal 1H–1H coupling constants.[17] The NOESY spectra provided conformationally relevant NOEs such as H-3/H-5, H-2/H-4, and NH/H-3 (Figure 5d). We further synthesized 3d-stearoyl-SM to deplete the overlapping H-3 signal and confirmed the presence of the NH/H-3 NOE correlation. The large values of 3JH-2/H-3 (10.7 Hz), 3JNH/H-2 (9.1 Hz), and 3JH-

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3/H-4 (7.7 Hz) in bicelles indicated anti conformations for H2/H-3, H-2/NH, and H-3/H-4 (Figure 5d).[15] From these NMR data and energy minimization, we determined the conformation of the SM partial structure (the C1–C6 and amide portions) in small bicelles (Figure 5d). The observed coupling constants typical for the anti orientation suggest that the C2– C6 and amide portions of SM adopt relatively rigid conformations in membrane environments.[15]

2.3. Orders of Alkyl Chains of SM To gain insight into the role played by chol in raft formation,[18] we next examined the effect of chol on the alkyl chains of SM using solid-state (SS) NMR. Solid-state 2H NMR spectroscopy allows noninvasive investigation of the order and mobility of acyl chains in lipid bilayers and has frequently revealed the chain order of phospholipids in membranes.[19] These 2H NMR measurements provide quadrupole splittings (Dm) of deuterium atoms located in lipid chains, which allow evaluation of the segmental dynamics (Figure 6). However, previous 2H NMR spectra were measured with perdeuterated acyl chains, which sometimes caused severe overlapping of quadrupole splitting signals,[20] thus hampering the specific



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Fig. 6. (a) Schematic of 2H NMR quadrupole splitting pattern of lipid alkyl chains. The quadrupole coupling directly indicates local mobility of each methylene group in the alkyl chains. (b) Quadrupole splitting magnitude obtained from the site-specifically deuterated stearoyl-SM derivatives on the sphingosine (left) and stearoyl (right) chains. Data were collected in the absence (blue circles) and presence (red circles) of 50 mol % cholesterol at 45 C.

assignment of deuterium signals. Another important consideration when using perdeuterated lipids is that the accumulated deuterium isotope effects alter the physicochemical properties of their lipid bilayers; e.g., the chain melting temperatures of perdeuterated acyl lipids decrease by several degrees.[21] To avoid these problems and, more importantly, to gain accurate local information on the lipid molecule dynamics, we synthesized a series of site-specifically deuterated SMs (Figure 6), which allowed us to accurately evaluate the segmental motions of the whole SM molecule and to examine the effect of chol on the SM membrane in detail.[22] The quadrupole splitting profiles of SSM in the absence and presence of chol (Figure 6) clearly indicated that chol enhances the order of the middle portions of the sphingosine and acyl chains.[22] Because the rigid alicyclic skeleton of chol decreases the fluctuation of the acyl chains of SSM,[23] thereby effectively augmenting the quadrupole coupling magnitude,[24] the 2H NMR derived ordering profiles in Figure 6 imply that the alicyclic skeleton of chol is preferentially located near the middle portions of the SSM hydrocarbon chains. This is consistent with the model that direct hydrogen-bonding interactions between the SM

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amide and the chol hydroxy are weak, if present at all, because direct interaction should enhance the order of the upper positions of the alkyl chains more prominently by anchoring the rigid alicycles of chol close to the upper chains. The results described above led to the following discoveries: (1) the conformation and orientation around the amide group of SM are relatively rigid and seem suitable for forming an intermolecular hydrogen bond with a neighboring SM molecule, (2) chol significantly enhances the order of the central hydrocarbon chains of SM, and (3) the SM/chol system is tolerant to temperature change, even at low chol concentrations. Collectively, these findings support a mechanism of raft formation wherein the rigid alicyclic chol enhances the order of the central SM alkyl chains by restricting the chain fluctuation, consequently shortening the intermolecular distances between SM molecules and facilitating the formation of intermolecular hydrogen bonds among SM molecules. The hydrogen bonds between SMs promote the formation of a hydrogen bond network of SM molecules, thereby forming a stable ordered phase robust to temperature fluctuations. The resultant higher thermal stability of SM/chol membranes at various chol



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Fig. 7. Fatty acid binding proteins (FABPs) are widely distributed in various tissues and play important cytoplasmic roles. (a) Fatty acids (FAs) are used in several organelles. In addition to their roles in energy production in mitochondria and peroxisomes by b-oxidation, FAs are signaling molecules for nuclear receptors such as peroxisome proliferator-activated receptor (PPAR), components of membrane lipids, and energy storage sources in the form of triglycerides. Reproduced with permission from reference [26]. (b) X-ray structure of FABP3 with stearic acid (Protein Data Bank code: 3WVM).

concentrations is indispensable for maintaining physiological homeostasis in lipid rafts. Therefore, SM is preferable to diacyl phosphatidylcholines as a component of lipid rafts.[25]

3. Lipid–Protein Interactions 3.1. Fatty Acid Binding Proteins (FABPs) As a first example of lipid–protein interactions (LPIs), we focus on fatty acid (FA) recognition by heart-type fatty acid binding protein (FABP3). Before undertaking the more challenging study of interactions between integral membrane proteins and their annular lipids, we chose to study a small protein (FABP3) that binds a simple lipid (FA) as an appropriate target. There

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are at least ten subtypes of human FABPs (Figure 7), among which FABP3 is the most abundant, particularly in the heart and skeletal muscle, and provides mitochondria with FAs as fuel to produce the ATP necessary for heart muscle contraction through b-oxidation (Figure 7). Thus, FABP3 functions intracellularly to transport FAs to and from membranes. In addition, FABPs play diverse cellular roles. The main difficulty in evaluating the binding affinity of FAs to FABP3 is a common problem for in vitro analysis of LPIs: hydrophobic lipids aggregate in aqueous media and/or adsorb on the instrument surface and thereby prevent accurate determination of effective ligand concentrations. To overcome this problem, we developed a method to comprehensively analyze FAs with long acyl chains. We used liposomes to mimic



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Fig. 8. Heat response obtained from ITC experiments. (a) Titration of FABP3 with FA-incorporating liposomes. (b) Oleic acid (OA, C18:1 n-9c) micelles, DMPC liposomes, and OA/DMPC (1:11 mol/mol) mixed liposomes were titrated into FABP3 solutions. The heat of binding was observed only when OA and DMPC were added as mixed liposomes. (c) Affinity index and thermal parameters obtained from ITC.[27]

the physiological environment and make the long-chain FAs (LCFAs) available to FABP3 (Figure 8a).[27,28] Our isothermal titration calorimetry (ITC) experiments clearly exhibited heat release upon binding of oleic acid to FABP3 (Figure 8b). Based on the heat generated, we determined the dissociation constant and the entropy and enthalpy of binding (Figure 8c) for a series of saturated FAs (SFAs) with varying chain lengths. FABP3 shows a clear preference for binding SFAs with C10–C18 chain lengths. The affinities for C10 and C18 were more than tenfold higher than for C8 and C20,[27] respectively, and we were intrigued by this broad but specific substrate selectivity for C10–C18 chain length SFAs.

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Co-crystals of FABP3 and C10–C18 SFAs were subjected to X-ray diffraction at 100 K, which yielded crystal structures with a very high resolution of 0.8–0.9 A˚ (Figure 9). Interestingly, the carbon atoms from C1 to C10 completely overlapped, while those beyond C10 were significantly different. The mechanism behind the promiscuity for C10–C18 chains is indicated by these crystal structures, in which the space created by the binding of a shorter FA is filled with water molecules. This finding is surprising because it suggests that an alkyl chain can be replaced by water without a large energy difference, but hydrocarbons and water molecules are usually considered to have opposite natures in terms of hydrophobicity and hydrophilicity.



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Fig. 9. Co-crystal structures of saturated fatty acids (C10:0–C18:0) bound to FABP3. (a) Electron density 2jFoj2jFcj maps of five C10–C18 saturated fatty acids (SFAs) bound to FABP3 at cryogenic temperatures. The resolution of each map was very high, ranging from 0.86 A˚ to 0.92 A˚. (b) Superimposed main-chain structures of five FABP3-bound SFAs with the key intermolecular interactions in FABP3–FA recognition indicated. (c) Contacts of C18:0 FA with water molecules (red balls) from the bottom and top of the co-crystal in the orientation in Figure 7b. (d) Binding pockets with C10:0 and C18:0 with the two clusters of water molecules. Red balls in the right panel denote the water molecules that are replaced by the C11–C18 part of C18 FA.

To illuminate the effect of water molecules on the “Ushaped” SFA-binding mechanism,[29] the hydration state of the C10:0–C18:0 FA/FABP3 complexes and the apo form were analyzed by MD using WaterMap,[30] which predicts hydration sites and allows estimation of the free energy associated with their formation. The MD simulations predicted that, upon FA binding, these water molecules segregated into two distinct water clusters (clusters 1 and 2) with significantly different energies and positioned in close agreement with the crystallographic data (Figure 10b).[27] The water molecules in cluster 1, which were highly ordered, formed a stable hydrogen-bonding network[29] (Figures 10a–c). In contrast, cluster 2 was confined to an unstable (high-energy) hydrophobic area surrounded by nonpolar amino acids and the FA hydrocarbon, and a significant desolvation energy release was

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expected by the displacement of water molecules with SFAs. The crystallographically observed U-shaped conformation was consistent with the energy conditions of binding-site hydration. Notably, the C18:0 chain almost exactly aligned with high-energy hydration sites (Figure 10c). ITC experiments performed under cell-mimicking conditions demonstrated that FABP3 recognizes FAs by the presence of free carboxyl and alkyl chains longer than C8, with particular preference for C10–C18, and by the flexibility of FAs to adopt the U-shaped conformation. The crystal structures with sub-angstrom resolution and the WaterMap analysis (Figure 10) confirm that the chain-length adaptability is supported by the disposition of the high-energy water molecules in cluster 2; the displacement with C10–C18 alkyl chains leads to enthalpy gain from desolvation, which is compensated by the entropy



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Fig. 10. Hydration sites identified by WaterMap in the binding pocket of FABP3 co-crystallized with SFAs.[27] Color gradation of hydration sites is based on the free energy relative to bulk water. Stable sites are shown in green, and unstable sites are shown in red. (a) Hydration state of the apo binding site. (b) Hydration state of FABP3 bound to C10:0. Note that water molecules are segregated into two clusters with distinct stabilities. (c) The extended chain of C18:0 displaces unstable water molecules in cluster 2.

lost in the same process (Figures 8 and 10). The lower affinity of C20–C24 FAs could be accounted for by disruption of highly ordered water molecules, which contact the C18 FA at the top part of cluster 1 (Figure 10c). Further elongation of an FA chain should disrupt the stable water network, thus destabilizing binding. Formation of this network is affected by the FA carboxylate, suggesting that the U shape of SFA in FABP3 is the ultimate design for measuring FA chain length.[27] From these observations, we conclude that FABP3 performs a fuzzy search for fuel FAs. Indeed, the promiscuity of FABP3 for binding FAs is broad enough to include major FAs and their derivatives in energy metabolism. FAs are thought to stabilize the proper structure of FABP3 since the protein in an apo form is easily degraded. We assumed that this instability should be related to conformational change for the apo protein to load an FA molecule, where a rigid b-clam structure is needed to become flexible in the ligand-absent form, and carried out an MD simulation.[31] The results clearly revealed that a relatively large gap between b strands 4 and 5 (Figure 7b) could be further enlarged in an apo form while, in the FA-bound form, this gap is shielded by a hydrogen-bond network to stabilize the FA-incorporated conformation.[31] Furthermore, we examined the co-crystal of FABP3 with a non-FA ligand since we expected that the specific binding mode for FAs should become clear by comparing ligands other than FAs. The cocrystal with 1-anilinonaphthalene-8-sulfonic acid (ANS) revealed that the orientation of ANS binding to FABP3 is completely opposite to that of ANS binding to FABP4 and, unlike FA, the anionic sulfonate group of ANS is not directly hydrogen bonded to Y128 and R126, suggesting that the binding site of acidic ligands in FABP3 is rather flexible, as is the case with FA chain length.[32]

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3.2. Bacteriorhodopsin We focused on bacteriorhodopsin (bR) as a model membrane protein since this protein has been subjected to extensive structural studies by SS NMR and X-ray crystallography.[33] Moreover, numerous reports have described the biochemical properties of bR in reconstituted membranes and detailed studies on the structure–function relationships have been conducted using DMPC and DPPC as model lipids.[34] Therefore, we could confirm the integrity of the protein by evaluating the retention of the original trimeric structure and proton-pumping function. The reconstituted lipid membranes of bR were prepared with diverse mammalian phospholipids and subjected to quantitative kinetic analysis of the decay rate constant of the M intermediate in the bR photocycle based on the data from laser flash photolysis (Figure 11). We found that addition of the acidic phospholipids phosphatidic acid (PA), phosphatidylglycerol (PG), and phosphatidylserine (PS) resulted in activity that was similar to or slightly lower than that of the purple membrane (PM).[35] In contrast, phosphatidylcholine (PC) caused markedly reduced activity. These changes due to differences in the structure of the lipid headgroup are related to acidification of the membrane surface, which seems to result from enriched proton levels. In addition, unsaturated lipids tended to preserve protein function, whereas a fully saturated lipid did not. Our findings showed that PGcontaining bilayers efficiently restore bR activity in reconstituted lipid membranes.[35] Thus, these lipids may substitute for PM lipids during functional reconstitution of bR and may be useful as a model system for studies of the molecular basis of LPIs in higher animals. In particular, both bulk lipids with low affinity and annular lipids with high affinity for the integral protein are necessary when the interaction is selectively



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Fig. 11. Phospholipid-binding structure of bacteriorhodopsin and its lipid assemblies. Left: A schematic bR trimer is shown, with one monomer showing the surface charges and another with the residue numbers of charged amino acids. Acidic amino acids gather at the inner part of the trimer, which, together with acidic lipids, promotes the H1pumping activity of bR.[37] As shown by the green wires, a few archaeal lipids are bound to specific positions of bR (PDB code: 2ZZL), while most of the annular lipids are not shown due to their disorder in crystals. Right: As the lipid/protein ratio is increased, bR-containing lipid assemblies take the shape of bilayer membranes while the trimers dissociate to monomers.

observed in membranes, which contain an excess amount of lipids over incorporated proteins to form stable proteoliposomes. To monitor the affinity of LPIs, our group developed an SS NMR methodology called centerband-only analysis of rotor-unsynchronized spin echo (COARSE),[36] which evaluates LPIs by measuring the chemical shift anisotropy (CSA) of 31P signals. We examined structural factors for the affinity between the PM lipid and bR by utilizing 31P COARSE, where inhibition of PM lipid–bR interaction by added phospholipid was evaluated as a decrease in the size of 31P CSA of the PM lipid. The results reinforced that both negative charge on the headgroup and the methyl-branched bulky alkyl chain are essential for lipids to have strong affinity to bR.

4. Membrane-Active Natural Products and Their Interactions with Lipids 4.1. Natural Products that Increase Membrane Permeability As described for bacteriorhodopsin in Section 3.2, LPIs are sometimes too weak to detect by SS NMR or X-ray crystallography. During our research on amphotericin B (AmB), the well-known and clinically important antifungal drug, we learned that natural products often show very high affinities for phospholipids and sterols in bilayer membranes when

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exerting their biological activities.[38] Thus, SS NMR studies on the interaction between membrane lipids and natural products should provide valuable information regarding possible NMR experiments for investigating LPIs. Furthermore, conventional solution NMR techniques can elucidate the structures of membrane-bound small molecules; e.g., standard twodimensional 1H–1H experiments are feasible with micelle suspensions or even with hydrated membrane dispersions. In this section, we review our recent findings on the membrane-active marine natural products theonellamide A (TNM-A) and amphidinol 3 (AM3). 4.2. Theonellamide A Theonellamides (TNMs, Figure 12), which were isolated from the marine sponge Theonella sp., belong to a family of unique bicyclic dodecapeptides.[39] TNM homologues possess a common bis-macrocyclic structure encompassing some unusual amino acids and a histidinoalanine bridge. Yeast chemical genomics revealed that TNMs were less effective on mutated cells lacking the ergosterol biosynthesis pathway,[40] and in vitro binding assays using a fluorescent derivative demonstrated that the peptide specifically bound to 3bhydroxysterols such as chol and ergosterol.[41] Judging from the phenotypic changes of yeasts, the membrane-related activity of TNMs is distinct from that of polyene antifungals such as AmB. However, direct interaction between TNMs and sterols in lipid bilayers had not been demonstrated. To address this



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Fig. 12. Structures of theonellamides (top) and solid-state 2H NMR spectra of sterols in the presence and absence of theonellamide A (bottom).[42] The POPC membrane dispersions were used for wide-line spectra in the absence (A, C, and E) and presence (B, D, and F) of TNM-A. 3-d-Cholesterol (A and B), 3-d-ergosterol (C and D), and 3-d-epicholesterol (E and F) were used. Center peaks are due to micelles and/or deuterium in water.

need, we carried out SS 2H NMR measurements using TNMA in POPC membranes containing chol, ergosterol, or 3-epicholesterol (Figure 12).[42] The 2H NMR spectra showed that TNM-A significantly inhibits the fast rotational motion of chol and ergosterol but not 3-epi-cholesterol, thus confirming the direct intermolecular interaction between TNM-A and 3b-hydroxysterols in lipid bilayers. The membrane action of TNMs is apparently different from that of AmB. We previously reported that sterols, particularly ergosterol, prominently promote not only the initial surface-binding step of AmB but also the subsequent reorientation process presumably corresponding to pore formation.[43] In particular, the second step of AmB action hardly proceeds without sterols present, suggesting that sterols, particularly ergosterol, are involved in the pore complex as shown below:

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AmB in H2O ! AmB on membrane ! AmB–sterol assembly in membrane TNM-A in H2O ! TNM-A on membrane ! TNM-A aggregate on membrane

Surface plasmon resonance (SPR) experiments clearly indicated that the second step of TNM-A binding proceeds without sterols and is not significantly accelerated by sterols.[41] These data indicate that TNM-A mostly recognizes the sterol hydroxy group in the initial binding process while AmB recognizes not only the 3-hydroxy group but also the steroid rings and side chains. This difference is consistent with the observation that TNM-A is unlikely to form distinctive pores as AmB does. Rather, it is more plausible that the dense accumulation of TNM-A in the sterol-containing membrane disturbs the bilayer morphology and integrity, which would correspond to the second step in TNM-A binding.



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Fig. 13. Lowest-energy conformation for the C20–C54 portion of amphidinol 3 (AM3) in isotropic bicelles (q 5 0.5) and hypothetical channel assembly of AM3. (a) Structure of AM3. (b) One of the lowest-energy conformations based on a systematic Monte Carlo method constrained by NOE and 3JHH data. Hydrogen atoms are omitted for clarity. (c) Flexible barrel-stave model for AM3-induced membrane channel.

4.3. Amphidinols Amphidinols (AMs) were isolated as potent antifungal agents from the epiphytic dinoflagellate Amphidinium klebsii. Several congeners, including some with closely related structures but different names, have been reported.[44] Of these, AM3 (Figure 13a) exhibits the most potent antifungal and hemolytic activities. AM3 enhances the permeability of bilayer membranes in a strictly sterol-dependent manner by forming pores or lesions in lipid bilayers,[45] which is thought to be responsible for the potent antifungal activity of AMs. Therefore, the 3D structure of AM3 in membranes probably accounts for its strong permeabilizing activity and strict sterol requirement. In contrast to AmB, which has a rigid conformation, AM3 is a highly flexible molecule, and elucidation of its conformation in membrane systems is therefore challenging. To gain insight into the active structure of AM3 in membranes, we adopted small bicelles and elucidated the conformation and topological orientation of AM3.[46,47] First, the conformation of AM3 was examined on the basis of the NOEs and 3JHH values in isotropic bicelles composed of DMPC-d54 and DHPC-d22 at a 1:2 molar ratio (see Figure 5). Although relevant NOEs could not be observed for C1–C20 or the poly-

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ene portion, probably due to molecular flexibility, the conformation for C20–C50 was established (Figure 13b). Measurements of the 1H NMR T1M values of AM3 in bicelles containing Mn21, which revealed how far apart the 1H position was from the membrane surface, showed deeper insertion of the olefinic terminus into the membrane interior, suggesting the predominance of an extended conformation for the hydrophobic polyene portion. We have reported that AM3 adopts a hairpin-like conformation with a relatively rigid turn structure near the center including the tetrahydropyran rings.[46–48] Based on these findings obtained from NMR results, a structural model of AM3 in the membrane was proposed (Figure 13c).[48] As described above, the turn region, which is conserved among other amphidinols, recognizes the 3b-OH group of sterols via hydrogen bonds. Therefore, a new type of barrelstave model with high flexibility may possibly explain the membrane permeabilization activity of AM3 interacting with sterol molecules.

5. Summary and Outlook In this review, we have first demonstrated that the combination of SS NMR techniques and organic synthesis is a powerful and



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critical tool to scrutinize atomic-level lipid interactions and dynamics. In particular, NMR-based studies of lipid–lipid and lipid–small molecule interactions require site-specifically labeled lipids. Closer collaborations between biophysicists and synthetic chemists will accelerate fundamental research in the structural biology of membrane lipids. In the next step, to elucidate the atomistic mechanism of LPIs, we utilized the ultrahigh-resolution X-ray structure of FABP3 obtained at a cryogenic temperature at which the highly flexible acyl chains of lipids and water molecules are mostly converged into a single conformer. Realistic lipid behaviors can be deduced from these high-resolution structures in combination with MD simulations and affinity evaluations at room temperature. Our results on FABP3 disclosed that the water clusters play crucial roles in the promiscuous recognition of C10–C18 fuel FAs. Lipids have great structural diversity, particularly with respect to chain length and degree of unsaturation in the acyl moieties. Thus, molecular recognition in biomembranes by these structurally similar but chemically diverse lipid mixtures is difficult to investigate with current methods in structural biology. Moreover, their inherent promiscuity in LPIs is another important feature of biomembranes. In our study on FABP3, we comprehensively examined the binding affinity of ten SFAs (C6–C24) and subjected them to X-ray crystallography to obtain accurate structures of their complexes.[27] Our results suggested an atomic-level mechanism underlying the promiscuous recognition of fuel FAs by FABP3. This strategy, based on combining affinity data and X-ray structure analysis for a series of structurally similar lipids, could potentially be useful for investigating LPIs, including those between membrane lipids and integral proteins. However, determining the accurate structure of their complexes is extremely difficult even with the use of X-ray crystallography or SS NMR, mainly because of the rapid exchange and high flexibility of annular lipids. We believe that these methods are still most reliable to elucidate the atomistic mechanism of LPIs, as briefly described in our study on bR, while combination with the other techniques such as imaging modalities, X-ray/neutron scattering and MD simulations would be necessary to efficiently promote LPI research. In this review, we have described our recent findings on natural products that bind to lipids in bilayer membranes. Their interactions including those of sterols are sometimes very strong, as we have revealed molecular contact between amphotericin B and ergosterol using SS NMR.[49] Theonellamide, bearing a cyclic peptide skeleton, also shows high affinity to chol. The detailed mechanism for this affinity, albeit currently unknown, may lead to a better understanding of sterol recognition by peptides and proteins. Thus, some of the natural products bearing potent membrane activities may serve as versatile tools for investigating the atomistic interaction mech-

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anisms implicated in LPIs and for developing methodologies necessary for their detection. Our current understanding of LPIs does not substantially elucidate the atomistic properties of biological membranes. To accelerate LPI research, experimental techniques for handling highly hydrophobic and water-insoluble ligands must be developed. Our recent results have indicated the usefulness of some experimental methods that may partly address this solubility problem; e.g., liposomes were an excellent vehicle in ITC and SPR experiments to obtain affinity indices for LPIs,[27,28] which were otherwise extremely difficult to obtain. In addition, X-ray crystallography of protein-bound lipids is very important for observing accurate 3D structures. Recent developments in crystallization of integral membrane proteins by lipid cubic phases and bicelles may further increase opportunities to obtain more accurate and biologically relevant structures of bound and annular lipids. Lipid research is also becoming more important in drug development. Besides oxidized metabolites of FAs such as prostaglandins, leukotriene and oxidized polyunsaturated fatty acids (PUFAs), many lipid derivatives play crucial roles in signal transduction, e.g., sphingosine-1-phosphate, diacylglycerol, lysophospholipids, and even FAs. Thus, LPIs will continue to provide basic knowledge about drug–receptor interactions. Moreover, membrane proteins targeted by current drugs continually interact with lipids. Thus, the drugs must have higher affinities for their targets than annular lipids, which further emphasizes the importance of LPIs. As we described in the introduction, biomembranes remain the bottleneck of life science research. Long-term basic investigations are still necessary to tackle this extremely challenging research topic.

Acknowledgements The results presented in this review were mainly achieved by the members of the JST ERATO “Lipid Active Structure” project, including Drs. Toshiaki Hara, Satoshi Kawatake, Masanao Kinoshita, Yuichi Umekawa, Sebastian Lethu, Daisuke Masutoka, and Fuminori Satou. We are grateful to Ph.D. students Tomokazu Yasuda, Rafael Espiritu, Kimberly Cornelio, Nagy Morsy, and Respati Suwasono in our laboratory, and Drs. Ichihara and Kimura in Schr€odinger Inc. for collaboration in WaterMap calculations. This research was partly supported by Grant-in-Aid No. 25242073 from MEXT Japan.

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Bioactive oligosaccharide natural products.

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Efficient total synthesis of bioactive natural products: a personal record.

Structure and Function of Macroalgal Natural Products.

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Structure Determination of Natural Products by Mass Spectrometry.

Natural bioactive compounds from winery by-products as health promoters: a review.

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Bioactive natural products with anti-herpes simplex virus properties.

Chemical and bioactive natural products from Microthyriaceae sp., an endophytic fungus from a tropical grass.

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Bioactive natural products from fungicolous Hawaiian isolates: secondary metabolites from a Phialemoniopsis sp.

apoptotic molecular pathways.

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Cardiovascular protection by natural products.

Regulation of membrane enzymes by lipids.

Asymmetric extraction of membrane lipids by CHAPS.

Mode of coupling between hormone receptors and adenylate cyclase elucidated by modulation of membrane fluidity.