- Email: [email protected]
The role of lipids in defining membrane protein interactions: insights from mass spectrometry
TICB-903; No. of Pages 8
Review
The role of lipids in defining membrane protein interactions: insights from mass spectrometry Nelson P. Barrera1, Min Zhou2, and Carol V. Robinson2 1 2
Department of Physiology, Pontificia Universidad Cato´lica de Chile, Alameda 340, Santiago, 8331150, Chile Department of Chemistry, Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford OX1 3QZ, UK
Cellular membranes comprise hundreds of lipids in which protein complexes, such as ion channels, receptors, and scaffolding complexes, are embedded. These protein assemblies act as signalling and trafficking platforms for processes fundamental to life. Much effort in recent years has focused on identifying the protein components of these complexes after their extraction from the lipid membrane in detergent micelles. Spectacular advances have been made using X-ray crystallography, providing in some cases detailed information about the mechanism of pumping and channel gating. These structural studies are leading to a growing realisation that, to understand their function, it is not only the structures of the protein components that are important but also knowledge of the protein–lipid interactions. This review highlights recent insights gained from this knowledge, surveys methods being developed for probing these interactions, and focuses specifically on the potential of mass spectrometry in this growing area of research. The lipid membrane Considering first the lipid composition of a simple prokaryotic species such as Escherichia coli, the outer and inner membranes limit the cell boundaries and are separated by a network of peptidoglycan. The outer membrane contains large numbers of pore-like proteins through which bulk transport may occur. The inner or cytoplasmic membrane contains numerous specific transport systems such as lactose permease (LacY) and the dicarboxylic acid transport system. Beyond its high protein content, the inner lipid bilayer comprises three main phospholipids: phosphatidylethanolamine (PE; zwitterionic, 74% of the total molar phospholipid content), phosphatidylglycerol (PG; bearing a negative charge, 19%), and cardiolipin (CL; bearing two negative charges, 3%) [1]. Although the composition and structures of many lipids are now established (Box 1), their functions in even a simple organism such as E. coli are not well understood. A further complication in assigning function to lipids arises because intrinsic membrane proteins are in contact Corresponding authors: Barrera, N.P. ([email protected]); Robinson, C.V. ([email protected]). Keywords: mass spectrometry; membrane protein complexes; detergent micelles; lipid binding; V-type ATPase.
with lipids to varying degrees. First, they are solvated by a shell of lipid molecules interacting with the membranepenetrating surface of the protein. These lipid molecules are referred to as annular lipids. Distinct from the annular lipids are the non-annular or ‘structural’ lipids that are found between transmembrane a-helices [2]. The most prevalent interactions with these structural lipids are through salt bridges formed between the lipid phosphate groups and basic residues in the protein, such as arginine and lysine. The acyl chains of these structural lipids are accommodated in hydrophobic pockets in the transmembrane domains (TMDs) [3,4]. Given this diversity in interactions, together with the different functional roles adopted by lipids in conferring structural stability, regulating channel opening and closure, and controlling the oligomeric state of protein complexes, it is unsurprising that cells contain hundreds of different lipids. How specific lipids are selected from the pool of available lipids for explicit tasks and, importantly, how these are then best characterised, is a major question in structural biology. Here we highlight recent literature in which several different roles for lipids have been proposed. We also summarise the various methods that have allowed these observations to take place. Fluorescence methods: observation of oligomeric state Turning our attention to the methods used to define the locations of lipids, solution-based fluorescence methods prove to be particularly exciting. In an elegant study using Fo¨rster resonance energy transfer (FRET), a direct and highly specific interaction of the COPI machinery protein (p24) with a sphingolipid (sphingomyelin) was investigated [5]. A distinct FRET signal was detected from a tryptophan residue in a maltose-binding protein (MBP) fusion of the TMDs of p24 and a fluorescently labelled analogue of sphingomyelin (pentaenoyl-sphingolipid SM18:5). Interestingly, a similar interaction was not detected using FRET between SM18.5 and the TMD of the related p23 protein, which is also involved in COPI vesicle biogenesis. Alanine scanning and a range of sphingomyelins were used to determine that both the hydrophobic acyl chains and the choline phosphate group are needed for the interaction. Binding of SM18:5 not only affects protein transport but was also shown, via crosslinking, to induce p24 dimerisation [5] (Figure 1a). By contrast, p23 does not contain the
0962-8924/$ – see front matter ß 2012 Published by Elsevier Ltd. http://dx.doi.org/10.1016/j.tcb.2012.08.007 Trends in Cell Biology xx (2012) 1–8
1
TICB-903; No. of Pages 8
Review
Trends in Cell Biology xxx xxxx, Vol. xxx, No. x
Box 1. Lipid structures Cells contain a plethora of lipids that fall into eight main classes: fatty acyls, glycerolipids, glycerophospholipids, sphingolipids, sterol lipids, prenol lipids, saccharolipids, and polyketides. All lipids are derived from these basic families and can include variability with respect to chain length and saturations in the hydrophobic carbon tails as well as modifications to the head group. The result of this diversity is that many different lipids constitute the membrane and more than 30 000 different lipid structures have been reported to date (www.lipidmaps.org). The greatest diversity of lipid structures has been observed for the glycerophospholipids. To represent simple lipid structures, the acid/acyl group and the hydrophobic chain are drawn on the right and left sides, respectively [46,47] (Figure I). In a similar manner, more complex lipids, such as glycerolipids, glycerophospholipids, and sphingolipids, are depicted with head groups on the right and hydrocarbon chains on the left. The LIPID MAPS consortium has developed a series of bioinformatics tools to generate lipid structures ‘on demand’ based on stored lipid structure templates [48]. Lipid structures can be identified from total cells or tissues [49] or, more specifically, from isolated protein complexes via mass spectrometry [21]. In the former case, where abundant samples are normally available, lipids are first extracted by chloroform/methanol [50] and then subjected to a combination of different purification and characterisation methods such as liquid chromatography–mass spectrometry (LC–MS). In the case of membrane complexes, an adapted method is applied. Isolated complexes are first digested in the presence of detergents before being applied to reverse-phase liquid chromatography. In this way, peptides, detergents, and lipids are separated and eluted as the gradient approaches 100% organic. The eluent is then analysed by mass spectrometry. The masses of the intact lipids and their fragment are then searched against the LIPID MAPS database to identify polar groups and hydrophobic chains [51]. When quantification of the lipids is necessary, their chromatographic
signature sequence required for binding and dimerisation induced by sphingolipid binding; consequently, no interaction was observed. This study therefore highlights the role of specific binding of a particular lipid structure to a signature sequence and its ability to control the oligomeric state of a protein, and importantly allows proposals to be made about its transport function. A different fluorescence-based method known as fluorescence cross-correlation spectroscopy (FCCS) can also establish oligomerisation, in this case by employing fluorescently labelled protein rather than lipids. Reconstitution of protein into well-defined giant unilamellar vesicles (GUVs) with different lipid components enables oligomerisation to be investigated as a function of the lipid composition. In this way, the oligomeric state of the mitochondrial voltage-dependent anion channel (VDAC) was investigated as a function of lipid structures [6]. VDAC is one of the most abundant channels located at the interface between mitochondria and the cytoplasm. It is implicated in homeostasis because it provides the main pathway for the exchange of metabolites, such as ATP and ADP, between the mitochondria and the cytosol. Using GUVs containing PG or CL, researchers showed that the functional dimeric form is triggered by binding of negatively charged PG lipids (Figure 1bi). By contrast, a CL lipid environment disrupts the oligomerisation of the channel (Figure 1bii). Because CL has four acyl chains compared with the two found in PG, the shape of the hydrophobic tails of the lipids is likely to play an important role in the multimerisation of VDAC [6]. The authors conclude that this difference in oligomerisation is related to the 2
peak areas, based on UV absorbance or total ion count on mass spectrometry, are compared with those of lipids of similar structures of known concentrations.
Glycerophospholipids Key: P C O N Phosphadylethanolamine
Cardiolipin Sphingolipid
Fay acyl
Sphingomyelin
Palmic acid
Sterol lipid
Cholesterol TRENDS in Cell Biology
Figure I. Basic lipid structures are illustrated following the conventions proposed by the LIPID MAPS consortium [48]. Carbon atoms are shown in cyan and heteroatoms phosphorous, oxygen, and nitrogen are colored green, red, and blue respectively.
different structures of the hydrophobic chains of these lipids as well as the nature of the protein–protein interactions responsible for VDAC oligomerisation. Mechanistic insights from X-ray crystallography Defining the oligomeric state of membrane complexes using X-ray crystallography can sometimes be problematic, because the high detergent concentration that is necessary for solubility may disrupt protein interactions during the crystallisation process. This could therefore lead to a dependence on detergent of the oligomeric state observed. This was investigated with an integral membrane protein from the mitochondrial carrier family (MCF) that transports metabolites over the inner mitochondrial membrane [7]. The first high-resolution structure obtained in the presence of the inhibitor carboxyatractyloside revealed a large cavity occupied by the inhibitor, within a monomeric structure [8]. A second X-ray structure demonstrated favourable protein–protein interactions mediated by endogenous CLs, two CLs being sandwiched between two subunits [9] (Figure 1biii). Endogenous lipids that remain bound to the protein during purification appear to be important for function, because previous experiments have shown that covalently linked dimeric assemblies are functional [10]. The observation of the dimeric ADP/ATP carrier in complex with endogenous lipids therefore enables proposals for the mechanism underlying nucleotide exchange across the inner mitochondrial membrane. Lipids also feature in the remarkable X-ray structure of photosystem II, in which densities for 30 subunits, all of
TICB-903; No. of Pages 8
Review
(a)
Trends in Cell Biology xxx xxxx, Vol. xxx, No. x
p23TMD sphingomyelin
p24TMD sphingomyelin
p24TMD dimer
(b)
VDAC dimer PG
(i)
VDAC monomers CL
(ii)
ADP/ATP carrier dimer CL
(iii)
(c)
CL12(PE)6
TRENDS in Cell Biology
Figure 1. Different biophysical approaches can be used to probe membrane lipid interactions, including fluorescence methods, X-ray crystallography, and mass spectrometry. (a) Probing the effects of lipid binding using Fo¨rster resonance
the metal atoms, and the five oxygen atoms of the Mn4CaO5 cluster were resolved [11]. More than 1300 water molecules were located in each photosystem II monomer together with a series of lipids. Interestingly, all of the PGs identified were distributed with their head groups located in the stromal surface of the membrane. This may suggest that the hydrophilic head groups cannot penetrate the membrane, resulting in their preferential distribution on the stromal side. The lipids appear to be dispersed throughout the structure, rather than in discrete binding pockets, mediating interactions between subunits, suggesting that these are annular rather than structural lipids. A particularly dramatic example of structural lipid binding was revealed in the recent X-ray structure of a short-chain derivative of phosphatidylinositol 4,5-bisphosphate (PIP(2)) in complex with Kir2.2 [12]. PIP(2) is a minor component of cell membranes and is known to regulate many ion channels [13]. The X-ray structure revealed that the PIP(2) lipid binds at an interface between the transmembrane and cytoplasmic domains. On lipid binding, a flexible expansion linker in Kir2.2 contracts to a compact helical structure and the cytoplasmic domains ˚ and become tethered to the transmembrane move 6 A domain. This in turn causes the inner helix gate to open. These results illustrate the role of PIP(2) in controlling the resting membrane potential through this specific ion channel–receptor ligand interaction. Furthermore this study highlights the importance of lipid binding in functional control [12]. An intriguing post-translational modification with consequences for lipid binding was reported recently for the bovine c-subunit of the F-ATP synthase [14]. A lysine group at the C terminus of the transmembrane a-helix was found to be completely trimethylated [14]. The resulting quaternary amino group is therefore exposed to the phospholipid bilayer, resulting in a possible steric clash with the head groups of the phospholipids, impeding their binding to the ring. CL is an essential component of the mitochondrial FATP synthase and could bind to trimethyl-lysine residues at the top of the FO-ring such that its acyl side chains would then be in a position to strengthen the membrane ring by filling the gaps between adjacent subunits. Interestingly, Lys-43 is conserved throughout all known c-subunit sequences in Animalia and it is likely that the trimethylation of this residue is conserved as a mean of stabilising FO energy transfer (FRET) measurements between fluorescently labelled sphingomyelin lipid interacting with the COPI machinery protein p24 transmembrane domain (TMD). The related p23 TMD does not bind to sphingomyelin and remains monomeric [5]. Energy-minimised p24 TMD and p23 TMD models were generated by Modeller9v8 [42] using the P-glycoprotein structure (PDB ID 3G61) as a template [43]. (b) Fluorescence correlation spectroscopy was used to probe labelled voltage-dependent anion channel (VDAC) protein bound to phosphatidylglycerol (PG) and cardiolipin (CL). Dimers were formed in the presence of PG (i) but not CL (ii). VDAC structures are from the X-ray structure of human VDAC (PDB ID 2JK4) [44]. X-Ray crystallography was used to examine the dimerisation of ADP/ATP carrier protein in the presence of CL [9] (iii). (c) Mass spectrometry was used to define the presence of six phosphatidylethanolamine (PE) lipids bound to the CL12 membrane rotor of the Thermus thermophilus (Tt) ATPase. Models for the 6-fold symmetric ring in which six L12 dimers (pink) are stabilised by specific lipid binding (green) and the docking of subunit C (blue) were obtained as described previously [34]. The lipid interaction was modelled between the PE head groups and Glu-63 in the L subunit via PatchDock [45] with further energy minimisation [34].
3
TICB-903; No. of Pages 8
Review
Trends in Cell Biology xxx xxxx, Vol. xxx, No. x
rings. CL has also been found in the Enterococcus hirae (Eh) V-type ATPase and PE in the membrane rotor of Thermus thermophilus (Tt) ATPase (Figure 1c) from mass spectrometry experiments described in the next section.
however, have mass spectrometry approaches been applied to intact membrane assemblies [19–21]. Using electrospray ionisation, an unexpected outcome of this research has been the finding that not only are membrane and soluble subunit interactions maintained, but also specific lipid binding is retained in almost all complexes studied to date [21,22]. This has enabled the identification of lipids bound specifically to protein subunits. Mass spectra have been recorded for several intact transporters, ion channels, and molecular machines, each with specific lipids bound tightly within the complexes (Figure 2). The major advantage of mass spectrometry in this area is in defining simultaneously the mass of the lipid and the
Mass spectrometry developments enable observation of specific binding of lipids Mass spectrometry is a relative newcomer to the membrane protein field, having long been the power behind proteomics. For the past two decades, it has been applied to the study of soluble protein complexes providing insight into their assembly [15], dynamics [16], polydispersity [17], and compositional heterogeneity [18]. Only recently, 100 26+
40+
27+
100
100
MacB 38+
23+
%
24+
%
%
MexB
33+
MtrE average charge
m/z 6500
0 6000
0
m/z
m/z
6000
5000
4000
8000
10000
0
50
LmrCD 40
25+
26+
100
100
KirBac3.1 30
28+
27+
24+
22+ 20
%
%
10
0 6000
5000
100
m/z
m/z
0 0
8+
2 100
EmrE
4
6
21+
0 5000
6000
ASA(A2x104) 23+
22+
100
LmrA
CL12
100
BtuCD
17+
%
%
8
%
%
x6 21+
0 3000
4000
m/z
0 6000
8000
m/z
0 6000
7000
m/z
m/z
5000
0 7000
TRENDS in Cell Biology
Figure 2. Mass spectrometry of various intact membrane protein complexes reveals specific binding of lipids. Mass spectra of four ABC transporters (MacB, LmrCD, LmrA, and BtuCD), three multidrug-resistance pumps (MtrE, MexB, and EmrE), an ion channel (KirBac3.1), and a membrane rotor (CL12 from the Tt ATPase). The average charge state recorded for soluble protein complexes is plotted against their accessible surface area (black line) and compared with the same plot for membrane protein complexes (red line) and a plot without the transmembrane regions included in the surface area calculation (pink line). The transmembrane domains in all protein complexes are coloured red. The lipids that were identified in each case are shown in a red/green space-filling representation [phosphatidylethanolamine (PE) and cardiolipin (CL) with two and four acyl chains, respectively].
4
TICB-903; No. of Pages 8
Review stoichiometry of its binding from measurement of the intact mass of membrane-containing complexes generated in solution or in the gas phase. From the mass of the lipid and its gas phase fragmentation pathway, it is often possible to define the head group, branching, and unsaturation sites. In this review, we focus on recent applications of mass spectrometry to the identification of lipids and the stoichiometry of their binding to membrane complexes, with special attention on their effect on subunit interactions and links to function. The major technological breakthrough that made this possible was the introduction into the gas phase of membrane complexes in protective detergent micelles. The precise mechanism by which these so-called ‘gas-phase micelles’ protect membrane complexes continues to inspire both theoretical simulations [23] and experimental investigation [24]. It is clear that during the phase transition, under the same instrumental set-up, soluble complexes are reduced to monomeric subunits [21]. However, how do the gas phase micelles protect the membrane complexes and what is the mechanism for their dissociation in the gas phase? Recent studies have shown that it is not the solution phase critical micelle value that governs the size of these gas phase micelles, because a plot of the average aggregation number against the alkyl chain length shows that the series from C6 trimethylammonium bromide (TAB) to C16TAB leads to a decrease in the aggregation number in the gas phase. Interestingly, therefore, this observation is in contrast to the established relationship in which increasing chain length leads to larger aggregation numbers in solution [24]. That the longer alkyl chain length is less stable in the gas phase provides a rationale for the choice of DDM over shorter chain detergents for a wide range of protein micelle complexes [21]. Recent proposals for the electrospray mechanism are relevant when considering the dissociation of gas phase micelles. A new view that appears to be well supported by experiment is that protein charging does not take place in the droplet, but rather at the point at which the protein enters the gas phase, after emission of charge carriers during droplet evaporation [25]. This has implications for the folded state of protein complexes, particularly those that have been expelled from micelles. If the micelle is still surrounding the transmembrane regions at the time of charging, these areas, which typically have fewer charges anyway, will not be charged. We investigated the relationship between charge and membrane surface area systematically by plotting the solvent-accessible surface area of several membrane complexes against their average charge state determined experimentally (Figure 2). We compared this relationship with a plot of the charge state of a representative selection of soluble complexes. It is clear that the average charge for membrane complexes is lower than for soluble ones of similar surface area. Interestingly, however, if the surface area attributed to the membrane regions is subtracted from the total surface area, the two lines largely coalesce. This implies that the membrane regions are not charged by virtue of their sequence because of their protection in the detergent micelle. Whatever the origin of this low charge, the absence of detergent molecules from the membrane complex points strongly to their
Trends in Cell Biology xxx xxxx, Vol. xxx, No. x
Box 2. IM of membrane protein complexes IM is an analytical technique that separates ions based on their gasphase mobilities. Application of this technique to membrane protein complexes, after their emergence from detergent micelles in the gas phase, presents a unique opportunity to probe their structure and dynamics, especially in regions that are otherwise wrapped by detergents in solution. In a typical experiment, membrane complexes are electrosprayed directly from micellar solutions and transferred intact into the gas phase protected within detergent micelles. The encapsulated complexes are subsequently released and pulsed into a drift tube filled with inert buffer gas, then migrate under a low electric field. While in the drift tube, ions undergo numerous collisions with the buffer gas molecules. The greater the CCS of an ion, the more gas molecules will collide with it, impeding their progress and thereby increasing the time taken for the ions to migrate through the drift tube. A detector at the end of the drift tube records the arrival times of the ions. These arrival times are then converted to experimental CCS values, which are a measurement of the orientationally averaged projections of a particular ion and is characteristic of its size and shape. When this is coupled with mass spectrometry (IM–MS), information on the size and shape of the ions can be obtained simultaneously. Ions of the same mass but different topology have distinct arrival times on IM. For example, ions with a compact conformation experience fewer collisions in the drift tube and hence travel faster and arrive earlier than those of similar size but with an extended conformation. Moreover, an increase in conformational dynamics and heterogeneity is evident from an expansion of the IM arrival time distribution (ATD) of the ions. This is informative for assessing structural dynamics and conformational heterogeneity in both the soluble and the transmembrane regions of membrane complexes.
role in forming a ‘protective shield’. This protective shield is lost during activation in the mass spectrometer, to reveal well-resolved mass spectra, enabling lipid binding to be defined. The protection afforded by the micelle, however, may not be sufficient to prevent unfolding of the membrane complex in the gas phase. We considered this possibility in a series of ion mobility (IM) experiments (Box 2) in which two membrane complexes were investigated: the transporter BtuC2D2 and the channel KirBac3.1 [26]. Because it is established that activation can lead to unfolding of protein complexes in the gas phase [27], we were keen to determine whether the activation conditions applied to remove detergent were sufficient to unfold the membrane protein complexes. To investigate this, we selected two membrane protein complexes of similar mass and surface area but with different topological arrangements. In the case of KirBac3.1, all four subunits make contact with the membrane, whereas for BtuC2D2 only the two BtuC subunits are protected by the micelle. Clear differences were observed, with the transporter being much less protected than the ion channel by virtue of the surrounding micelle [26]. However, under very low activation conditions, it was possible to obtain relatively compact structures for both complexes, validating the use of IM mass spectrometry for membrane complexes released from detergent micelles. In addition to annular and non-annular or structural lipids, there is a third class that is used as substrates. This is well documented for ABC transporters that interact with membrane lipids and in some cases use phospholipids as substrates [28–30]. In addition, these transporters can require specific lipids to perform substrate transport. The ABC transporter Aus1, reconstituted in proteoliposomes, 5
TICB-903; No. of Pages 8
Review
Trends in Cell Biology xxx xxxx, Vol. xxx, No. x
increases its ATPase activity by direct interaction between the protein and phosphatidylserine [31]. These results suggest strong interactions between lipids and the transmembrane domains of this ABC transporter. Similarly, mass spectra of the ABC transporter MacB revealed the presence of two PE molecules [21]. Mass spectra of two other ABC transporters showed binding of one and two CL molecules to LmrA [32] and LmrCD [21], respectively. Under the conditions used in these experiments, it is likely that annular lipids are absent [33]. Moreover, the absence of lipids associated with monomeric subunits implies that lipid binding occurs primarily within subunit interfaces. Whether these lipids are transported or are required for optimal protein stability is difficult to define in the absence of a functional assay. However, their persistence in various different detergents implies a high specificity of interaction. Different modes of lipid binding revealed by mass spectrometry Turning to the largest of the membrane complexes recently studied by electrospray mass spectrometry, two rotary ATPases/synthases from Tt and Eh, comprising almost 30 subunits and many lipids, were preserved intact in the gas phase [34]. The fact that the subunit–lipid interactions were maintained in these assemblies is particularly interesting, because two very different lipid-binding patterns were uncovered. Using a combination of the mass spectra of the intact species together with liquid chromatography of the individual subunits and gas phase fragmentation of the endogenous lipids, the masses and structures of the lipids could be ascertained. Quantitative proteomics and lipidomics were then used to confirm their relative abundances. For the membrane rotor from Eh ATPase, which contains 10 K subunits each with four transmembrane helices, a series of CLs were identified with a stoichiometry of 1:1 for rotor subunit:CL. This represents a change in the interpretation of the lipids bound to the K ring. These lipids were previously observed in an X-ray crystallographic structure of the isolated ring and assigned to 20 PG molecules based on the high cellular abundance of this lipid [35]. The similar volume occupied by four acyl chains in 10 CLs as opposed to two acyl chains (a)
(b)
Dri me (ms)
22
100
16.91
19.4
100 10.62
17 0
5
20 35
26+
24+
12
7
12-fold symmetry
20 Dri me (ms)
6-fold symmetry
in 20 PG molecules is, however, consistent with two possible assignments to this electron density. The mass of the intact membrane complex together with the lipidomics enables us to delineate these possibilities and to confirm that 10 CLs are located in the orifice of the ring [34]. It should be noted, however, that without the X-ray structure, the location of the 10 CLs would not have been possible from mass spectrometry experiments alone. For Tt ATPase, 12 L subunits each with two transmembrane helices occupy the membrane rotor. Six PE molecules per intact Tt ATPase were identified using quantitative proteomics and lipidomics as well as knowledge of the mass of the intact ATPase and its subcomplexes. Results from the intact mass and the lipidomics were consistent with a stoichiometry of 2:1 for L subunit:lipid interactions. This substoichiometric binding suggests that the 12 subunits undergo rotation to form six dimeric L subunit pairs, each stabilised by specific lipid binding, consequently with four transmembrane helices per subunit (Figure 3 left). This changes the proton:ATP ratio in favour of proton pumping rather than ATP synthesis. Interestingly, this configuration is in accord with the 6fold electron density of the L ring observed previously [36]. This 6-fold symmetric state is also of interest mechanistically because, previously it was established that Tt ATPase can operate both as a proton pump and to synthesise ATP [37]. This led to the proposal that this specific lipid binding switches this rotary ATPase from an ATP synthase to an ion-pumping V-type ATPase [34]. In this mode, pairs of subunits interact and are stabilised by specific lipid binding to produce subunits with four transmembrane helices in line with all other membrane rotors of the V-type ATPases examined to date [35]. Interestingly, the central subunit within all membrane rotors of the rotary ATPases makes remarkably similar contact with subunits in the central stalk to propagate rotation from the membrane rotor to the soluble head. The size of this central subunit in the centre of the rotor is largely conserved, suggesting that the orifices of the membrane rotors should be similar. However, this is clearly not the case. The number of subunits in the ring appears to be fixed for a given species, but this stoichiometry varies
25+
15 0
5
11.49
21+ 22+ 20 35 23+
10
5
21+
m/z
5000 6000 7000 8000 9000
5000 6000 7000 8000 9000 TRENDS in Cell Biology
Figure 3. Ion mobility (IM) mass spectrometry of the Thermus thermophilus (Tt) ATPase membrane-containing subcomplexes. IM arrival time distribution (ATD) (blue) reveals a broadening of the distribution for ICL12 (a) compared with CL12 (b). This broadening is also observed in the extracted ATD peaks (green and blue base, respectively). Models of the 6-fold symmetry (left) stabilised by the binding of six phosphatidylethanolamine (PE) lipids (red) and the membrane rotor with 12-fold symmetry (right). Homology models for the 12-fold and 6-fold symmetry L12 rings were built using the L subunit model described previously [34]. Hybrid atomic and coarsegrained (dark green spheres) models for the I subunit were built based on both previous I subunit modelling and the recent electron microscopy density map [34,39].
6
TICB-903; No. of Pages 8
Review between species [38]. How, then, are the different membrane rings adapted for rotation of the central shaft? Consider the two examples examined here, where in situ lipid binding was identified and molecular modelling of the 12-fold L12 ring based on homology to two transmembrane helices from the Eh K ring structure [35] gave rise to a ˚ . Rotation to form six L dimers (L2)6, central orifice of 43 A with each dimer stabilised by one lipid, reduces the inner ˚ . This compares with the orifice diameter of (L2)6 to 39 A calculated for the K10 membrane rotor of Eh ATPase, in which 10 bulky CLs bind to the internal cavity, reducing it ˚ in the absence of lipids to 39 A ˚ in the presence of from 54 A these lipid brushes. This implies that lipid binding to both membrane rotors reduces the internal cavities to similar values for Eh and Tt ATPases, although the effects are much more marked in the case of the Eh ATPase. To investigate conformational heterogeneity in the rotary ATPases, we used IM mass spectrometry, as outlined above, to probe the collision cross-section (CCS) of various subcomplexes. The broadening of the arrival time distribution peaks for the membrane-embedded Tt ICL12, compared with the CL12 peaks, is consistent with enhanced conformational dynamics of subunit I (Figure 3a,b). Subunit I contains a transmembrane portion which is at 908 to the soluble arm. IM CCS data, together with molecular modelling, suggest that this soluble arm can undergo conformational dynamics from 908 to 1208. This observation, coupled with a proposed nucleotide-binding site in subunit I at the hinge region of the 908 configuration, which binds preferentially to ADP, led us to propose that ADP binding triggers a conformational change. This conformational change in turn causes subunit I to move away from the proton conductance channel formed between the membrane I and the L subunits in the ring, and disrupt the channel. Lipids from the membrane would then seal the gap left by the subunit movement when ATP levels are depleted, and in so doing preserve the proton gradient created at the expense of ATP [34]. Subsequently, a high-resolution electron cryomicroscopy (cryo-EM) structure of the intact Tt ATPase at ˚ was reported. This technological breakthrough shows 9.7 A 12-fold symmetry for the L12 ring [39]; presumably in this case, by contrast to the one above, the Tt ATPase was caught in an ATP synthesis mode (Figure 3 right). Although lipid binding could not be discerned in this cryoEM structure and the detergent micelle contributes to the overall dimensions, when compared with a sixfold model produced from our experimental data, the two rotor rings appear remarkably similar. This would imply that only minor rearrangements in the ring would be enough to trigger the switch between pumping and ATP synthesis. Isolating this rotary Tt ATPase in its two different modes of action and investigating lipid-binding patterns in the two possible forms is now a major goal. In summary, these studies of the two rotary ATPases described here highlight the importance of combining electron microscopy and X-ray crystallography with mass spectrometry. The synergy of these techniques has not only enabled the location of lipids to be defined, but also their identity and stoichiometry to be deduced. In the future, coupling this lipid-binding information with the
Trends in Cell Biology xxx xxxx, Vol. xxx, No. x
location of post-translational modifications will enable their effects on subunit interfaces and lipid binding to be understood. Ultimately, this will add to our understanding of the regulation and control of these fascinating rotary motors. Concluding remarks The diversity of lipid structures that are being uncovered and the beginnings of an understanding of their roles in organising and maintaining membrane protein interactions is sparking considerable interest from structural biologists and lipid chemists alike. Interestingly, lipids that are found in subunit interfaces are often of minimal abundance in the membrane in which the complex is embedded, as was found for the V-type ATPases [40] and for the ion channels that select (PIP)2, a minor component of the cell membrane [41]. This leads to the proposal that complexes select lipids from the available pool for specific tasks rather than using the most abundant lipids. The four transmembrane K subunits of Eh ATPase select lipids with four acyl chains, whereas the two transmembrane L subunits use PE lipids with two acyl chains. This specific tailoring of lipids is also observed for the VDAC dimer, p24, and the ADP/ATP carrier protein, which binds preferentially to PG, sphingomyelin, and CL, respectively. At this stage, it is unclear how these recognition events occur. It is clear, however, that more information is needed before definitive conclusions can be made based not only on structural recognition but also on functional requirements. Increasingly evident is the fact that lipids can no longer be ignored in the structures of membrane complexes; their ability to fine-tune interactions and to stabilise different interfaces is leading to numerous mechanistic insights. With the new biophysical approaches that are coming to the fore to uncover lipid binding, it seems likely that mechanistic understanding of the role of lipids will advance rapidly. In the near future, it is hoped that this will lead to fascinating insights into the role of the diversity of lipid structures and into their correlated binding events. Acknowledgements Funding from Fondo Nacional de Desarrollo Cientı´fico y Tecnolo´gico (FONDECYT) regular grant #1120169 and the Millennium Scientific Initiative (Ministerio de Economı´a, Fomento y Turismo) #P10-035-F (N.P.B.), an ERC advanced grant and the Wellcome Trust (M.Z.), and the Royal Society (C.V.R.) is acknowledged.
References 1 Dowhan, W. (1997) Molecular basis for membrane phospholipid diversity: why are there so many lipids? Annu. Rev. Biochem. 66, 199–232 2 Lee, A.G. (2011) Lipid-protein interactions. Biochem. Soc. Trans. 39, 761–766 3 Adamian, L. et al. (2011) Lipid-binding surfaces of membrane proteins: evidence from evolutionary and structural analysis. Biochim. Biophys. Acta 1808, 1092–1102 4 Lensink, M.F. et al. (2010) Identification of specific lipid-binding sites in integral membrane proteins. J. Biol. Chem. 285, 10519–10526 5 Contreras, F.X. et al. (2012) Molecular recognition of a single sphingolipid species by a protein’s transmembrane domain. Nature 481, 525–529 6 Betaneli, V. et al. (2012) The role of lipids in VDAC oligomerization. Biophys. J. 102, 523–531 7
TICB-903; No. of Pages 8
Review 7 Palmieri, F. (2004) The mitochondrial transporter family (SLC25): physiological and pathological implications. Pflugers Arch. 447, 689– 709 8 Pebay-Peyroula, E. et al. (2003) Structure of mitochondrial ADP/ATP carrier in complex with carboxyatractyloside. Nature 426, 39–44 9 Nury, H. et al. (2005) Structural basis for lipid-mediated interactions between mitochondrial ADP/ATP carrier monomers. FEBS Lett. 579, 6031–6036 10 Trezeguet, V. et al. (2000) A covalent tandem dimer of the mitochondrial ADP/ATP carrier is functional in vivo. Biochim. Biophys. Acta 1457, 81–93 11 Umena, Y. et al. (2011) Crystal structure of oxygen-evolving photosystem II at a resolution of 1.9 A. Nature 473, 55–60 12 Hansen, S.B. et al. (2011) Structural basis of PIP2 activation of the classical inward rectifier K+ channel Kir2.2. Nature 477, 495–498 13 Suh, B.C. and Hille, B. (2008) PIP2 is a necessary cofactor for ion channel function: how and why? Annu. Rev. Biophys. 37, 175–195 14 Watt, I.N. et al. (2010) Bioenergetic cost of making an adenosine triphosphate molecule in animal mitochondria. Proc. Natl. Acad. Sci. U. S. A. 107, 16823–16827 15 Sakata, E. et al. (2011) The catalytic activity of Ubp6 enhances maturation of the proteasomal regulatory particle. Mol. Cell 42, 637–649 16 Uetrecht, C. et al. (2008) High-resolution mass spectrometry of viral assemblies: molecular composition and stability of dimorphic hepatitis B virus capsids. Proc. Natl. Acad. Sci. U. S. A. 105, 9216–9220 17 Stengel, F. et al. (2010) Quaternary dynamics and plasticity underlie small heat shock protein chaperone function. Proc. Natl. Acad. Sci. U. S. A. 107, 2007–2012 18 Ebong, I.O. et al. (2011) Heterogeneity and dynamics in the assembly of the Heat Shock Protein 90 chaperone complexes. Proc. Natl. Acad. Sci. U. S. A. 108, 17939–17944 19 Morgner, N. et al. (2007) A novel approach to analyze membrane proteins by laser mass spectrometry: from protein subunits to the integral complex. J. Am. Soc. Mass Spectrom. 18, 1429–1438 20 Barrera, N.P. et al. (2008) Micelles protect membrane complexes from solution to vacuum. Science 321, 243–246 21 Barrera, N.P. et al. (2009) Mass spectrometry of membrane transporters reveals subunit stoichiometry and interactions. Nat. Methods 6, 585–587 22 Barrera, N.P. and Robinson, C.V. (2011) Advances in the mass spectrometry of membrane proteins: from individual proteins to intact complexes. Annu. Rev. Biochem. 80, 247–271 23 Friemann, R. et al. (2009) Molecular dynamics simulations of a membrane protein-micelle complex in vacuo. J. Am. Chem. Soc. 131, 16606–16607 24 Borysik, A.J. and Robinson, C.V. (2012) Formation and dissociation processes of gas-phase detergent micelles. Langmuir 28, 7160–7167 25 Hogan, C.J., Jr et al. (2008) Charge carrier field emission determines the number of charges on native state proteins in electrospray ionization. J. Am. Chem. Soc. 130, 6926–6927 26 Wang, S.C. et al. (2010) Ion mobility mass spectrometry of two tetrameric membrane protein complexes reveals compact structures and differences in stability and packing. J. Am. Chem. Soc. 132, 15468–15470 27 Ruotolo, B.T. et al. (2007) Ion mobility-mass spectrometry reveals longlived, unfolded intermediates in the dissociation of protein complexes. Angew. Chem. Int. Ed. Engl. 46, 8001–8004
8
Trends in Cell Biology xxx xxxx, Vol. xxx, No. x
28 Doshi, R. et al. (2010) Dissection of the conformational cycle of the multidrug/lipidA ABC exporter MsbA. Proteins 78, 2867–2872 29 Nagao, K. et al. (2010) Lipid outward translocation by ABC proteins. FEBS Lett. 584, 2717–2723 30 van Meer, G. et al. (2006) ABC lipid transporters: extruders, flippases, or flopless activators? FEBS Lett. 580, 1171–1177 31 Marek, M. et al. (2011) The yeast plasma membrane ATP binding cassette (ABC) transporter Aus1: purification, characterization, and the effect of lipids on its activity. J. Biol. Chem. 286, 21835–21843 32 Velamakanni, S. et al. (2009) A multidrug ABC transporter with a taste for salt. PLoS ONE 4, e6137 33 Hite, R.K. et al. (2010) Principles of membrane protein interactions with annular lipids deduced from aquaporin-0 2D crystals. EMBO J. 29, 1652–1658 34 Zhou, M. et al. (2011) Mass spectrometry of intact V-type ATPases reveals bound lipids and the effects of nucleotide binding. Science 334, 380–385 35 Murata, T. et al. (2005) Structure of the rotor of the V-Type Na+ATPase from Enterococcus hirae. Science 308, 654–659 36 Bernal, R.A. and Stock, D. (2004) Three-dimensional structure of the intact Thermus thermophilus H+-ATPase/synthase by electron microscopy. Structure 12, 1789–1798 37 Yokoyama, K. et al. (2000) V-Type H+-ATPase/synthase from a thermophilic eubacterium, Thermus thermophilus. Subunit structure and operon. J. Biol. Chem. 275, 13955–13961 38 Pogoryelov, D. et al. (2012) Engineering rotor ring stoichiometries in the ATP synthase. Proc. Natl. Acad. Sci. U. S. A. 109, E1599–E1608 39 Lau, W.C. and Rubinstein, J.L. (2012) Subnanometre-resolution structure of the intact Thermus thermophilus H+-driven ATP synthase. Nature 481, 214–218 40 Ray, P.H. et al. (1971) Effect of growth temperature on the lipid composition of Thermus aquaticus. J. Bacteriol. 108, 227–235 41 Jiang, Q.X. and Gonen, T. (2012) The influence of lipids on voltagegated ion channels. Curr. Opin. Struct. Biol. 22, 529–536 42 Sali, A. and Blundell, T.L. (1993) Comparative protein modelling by satisfaction of spatial restraints. J. Mol. Biol. 234, 779–815 43 Aller, S.G. et al. (2009) Structure of P-glycoprotein reveals a molecular basis for poly-specific drug binding. Science 323, 1718–1722 44 Bayrhuber, M. et al. (2008) Structure of the human voltage-dependent anion channel. Proc. Natl. Acad. Sci. U. S. A. 105, 15370–15375 45 Schneidman-Duhovny, D. et al. (2005) PatchDock and SymmDock: servers for rigid and symmetric docking. Nucleic Acids Res. 33, W363–W367 46 Fahy, E. et al. (2005) A comprehensive classification system for lipids. J. Lipid Res. 46, 839–861 47 Sud, M. et al. (2007) LMSD: LIPID MAPS structure database. Nucleic Acids Res. 35, D527–D532 48 Subramaniam, S. et al. (2011) Bioinformatics and systems biology of the lipidome. Chem. Rev. 111, 6452–6490 49 Ivanova, P.T. et al. (2009) Lipidomics: a mass spectrometry based systems level analysis of cellular lipids. Curr. Opin. Chem. Biol. 13, 526–531 50 Bligh, E.G. and Dyer, W.J. (1959) A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol. 37, 911–917 51 Harkewicz, R. and Dennis, E.A. (2011) Applications of mass spectrometry to lipids and membranes. Annu. Rev. Biochem. 80, 301–325
Review
The role of lipids in defining membrane protein interactions: insights from mass spectrometry Nelson P. Barrera1, Min Zhou2, and Carol V. Robinson2 1 2
Department of Physiology, Pontificia Universidad Cato´lica de Chile, Alameda 340, Santiago, 8331150, Chile Department of Chemistry, Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford OX1 3QZ, UK
Cellular membranes comprise hundreds of lipids in which protein complexes, such as ion channels, receptors, and scaffolding complexes, are embedded. These protein assemblies act as signalling and trafficking platforms for processes fundamental to life. Much effort in recent years has focused on identifying the protein components of these complexes after their extraction from the lipid membrane in detergent micelles. Spectacular advances have been made using X-ray crystallography, providing in some cases detailed information about the mechanism of pumping and channel gating. These structural studies are leading to a growing realisation that, to understand their function, it is not only the structures of the protein components that are important but also knowledge of the protein–lipid interactions. This review highlights recent insights gained from this knowledge, surveys methods being developed for probing these interactions, and focuses specifically on the potential of mass spectrometry in this growing area of research. The lipid membrane Considering first the lipid composition of a simple prokaryotic species such as Escherichia coli, the outer and inner membranes limit the cell boundaries and are separated by a network of peptidoglycan. The outer membrane contains large numbers of pore-like proteins through which bulk transport may occur. The inner or cytoplasmic membrane contains numerous specific transport systems such as lactose permease (LacY) and the dicarboxylic acid transport system. Beyond its high protein content, the inner lipid bilayer comprises three main phospholipids: phosphatidylethanolamine (PE; zwitterionic, 74% of the total molar phospholipid content), phosphatidylglycerol (PG; bearing a negative charge, 19%), and cardiolipin (CL; bearing two negative charges, 3%) [1]. Although the composition and structures of many lipids are now established (Box 1), their functions in even a simple organism such as E. coli are not well understood. A further complication in assigning function to lipids arises because intrinsic membrane proteins are in contact Corresponding authors: Barrera, N.P. ([email protected]); Robinson, C.V. ([email protected]). Keywords: mass spectrometry; membrane protein complexes; detergent micelles; lipid binding; V-type ATPase.
with lipids to varying degrees. First, they are solvated by a shell of lipid molecules interacting with the membranepenetrating surface of the protein. These lipid molecules are referred to as annular lipids. Distinct from the annular lipids are the non-annular or ‘structural’ lipids that are found between transmembrane a-helices [2]. The most prevalent interactions with these structural lipids are through salt bridges formed between the lipid phosphate groups and basic residues in the protein, such as arginine and lysine. The acyl chains of these structural lipids are accommodated in hydrophobic pockets in the transmembrane domains (TMDs) [3,4]. Given this diversity in interactions, together with the different functional roles adopted by lipids in conferring structural stability, regulating channel opening and closure, and controlling the oligomeric state of protein complexes, it is unsurprising that cells contain hundreds of different lipids. How specific lipids are selected from the pool of available lipids for explicit tasks and, importantly, how these are then best characterised, is a major question in structural biology. Here we highlight recent literature in which several different roles for lipids have been proposed. We also summarise the various methods that have allowed these observations to take place. Fluorescence methods: observation of oligomeric state Turning our attention to the methods used to define the locations of lipids, solution-based fluorescence methods prove to be particularly exciting. In an elegant study using Fo¨rster resonance energy transfer (FRET), a direct and highly specific interaction of the COPI machinery protein (p24) with a sphingolipid (sphingomyelin) was investigated [5]. A distinct FRET signal was detected from a tryptophan residue in a maltose-binding protein (MBP) fusion of the TMDs of p24 and a fluorescently labelled analogue of sphingomyelin (pentaenoyl-sphingolipid SM18:5). Interestingly, a similar interaction was not detected using FRET between SM18.5 and the TMD of the related p23 protein, which is also involved in COPI vesicle biogenesis. Alanine scanning and a range of sphingomyelins were used to determine that both the hydrophobic acyl chains and the choline phosphate group are needed for the interaction. Binding of SM18:5 not only affects protein transport but was also shown, via crosslinking, to induce p24 dimerisation [5] (Figure 1a). By contrast, p23 does not contain the
0962-8924/$ – see front matter ß 2012 Published by Elsevier Ltd. http://dx.doi.org/10.1016/j.tcb.2012.08.007 Trends in Cell Biology xx (2012) 1–8
1
TICB-903; No. of Pages 8
Review
Trends in Cell Biology xxx xxxx, Vol. xxx, No. x
Box 1. Lipid structures Cells contain a plethora of lipids that fall into eight main classes: fatty acyls, glycerolipids, glycerophospholipids, sphingolipids, sterol lipids, prenol lipids, saccharolipids, and polyketides. All lipids are derived from these basic families and can include variability with respect to chain length and saturations in the hydrophobic carbon tails as well as modifications to the head group. The result of this diversity is that many different lipids constitute the membrane and more than 30 000 different lipid structures have been reported to date (www.lipidmaps.org). The greatest diversity of lipid structures has been observed for the glycerophospholipids. To represent simple lipid structures, the acid/acyl group and the hydrophobic chain are drawn on the right and left sides, respectively [46,47] (Figure I). In a similar manner, more complex lipids, such as glycerolipids, glycerophospholipids, and sphingolipids, are depicted with head groups on the right and hydrocarbon chains on the left. The LIPID MAPS consortium has developed a series of bioinformatics tools to generate lipid structures ‘on demand’ based on stored lipid structure templates [48]. Lipid structures can be identified from total cells or tissues [49] or, more specifically, from isolated protein complexes via mass spectrometry [21]. In the former case, where abundant samples are normally available, lipids are first extracted by chloroform/methanol [50] and then subjected to a combination of different purification and characterisation methods such as liquid chromatography–mass spectrometry (LC–MS). In the case of membrane complexes, an adapted method is applied. Isolated complexes are first digested in the presence of detergents before being applied to reverse-phase liquid chromatography. In this way, peptides, detergents, and lipids are separated and eluted as the gradient approaches 100% organic. The eluent is then analysed by mass spectrometry. The masses of the intact lipids and their fragment are then searched against the LIPID MAPS database to identify polar groups and hydrophobic chains [51]. When quantification of the lipids is necessary, their chromatographic
signature sequence required for binding and dimerisation induced by sphingolipid binding; consequently, no interaction was observed. This study therefore highlights the role of specific binding of a particular lipid structure to a signature sequence and its ability to control the oligomeric state of a protein, and importantly allows proposals to be made about its transport function. A different fluorescence-based method known as fluorescence cross-correlation spectroscopy (FCCS) can also establish oligomerisation, in this case by employing fluorescently labelled protein rather than lipids. Reconstitution of protein into well-defined giant unilamellar vesicles (GUVs) with different lipid components enables oligomerisation to be investigated as a function of the lipid composition. In this way, the oligomeric state of the mitochondrial voltage-dependent anion channel (VDAC) was investigated as a function of lipid structures [6]. VDAC is one of the most abundant channels located at the interface between mitochondria and the cytoplasm. It is implicated in homeostasis because it provides the main pathway for the exchange of metabolites, such as ATP and ADP, between the mitochondria and the cytosol. Using GUVs containing PG or CL, researchers showed that the functional dimeric form is triggered by binding of negatively charged PG lipids (Figure 1bi). By contrast, a CL lipid environment disrupts the oligomerisation of the channel (Figure 1bii). Because CL has four acyl chains compared with the two found in PG, the shape of the hydrophobic tails of the lipids is likely to play an important role in the multimerisation of VDAC [6]. The authors conclude that this difference in oligomerisation is related to the 2
peak areas, based on UV absorbance or total ion count on mass spectrometry, are compared with those of lipids of similar structures of known concentrations.
Glycerophospholipids Key: P C O N Phosphadylethanolamine
Cardiolipin Sphingolipid
Fay acyl
Sphingomyelin
Palmic acid
Sterol lipid
Cholesterol TRENDS in Cell Biology
Figure I. Basic lipid structures are illustrated following the conventions proposed by the LIPID MAPS consortium [48]. Carbon atoms are shown in cyan and heteroatoms phosphorous, oxygen, and nitrogen are colored green, red, and blue respectively.
different structures of the hydrophobic chains of these lipids as well as the nature of the protein–protein interactions responsible for VDAC oligomerisation. Mechanistic insights from X-ray crystallography Defining the oligomeric state of membrane complexes using X-ray crystallography can sometimes be problematic, because the high detergent concentration that is necessary for solubility may disrupt protein interactions during the crystallisation process. This could therefore lead to a dependence on detergent of the oligomeric state observed. This was investigated with an integral membrane protein from the mitochondrial carrier family (MCF) that transports metabolites over the inner mitochondrial membrane [7]. The first high-resolution structure obtained in the presence of the inhibitor carboxyatractyloside revealed a large cavity occupied by the inhibitor, within a monomeric structure [8]. A second X-ray structure demonstrated favourable protein–protein interactions mediated by endogenous CLs, two CLs being sandwiched between two subunits [9] (Figure 1biii). Endogenous lipids that remain bound to the protein during purification appear to be important for function, because previous experiments have shown that covalently linked dimeric assemblies are functional [10]. The observation of the dimeric ADP/ATP carrier in complex with endogenous lipids therefore enables proposals for the mechanism underlying nucleotide exchange across the inner mitochondrial membrane. Lipids also feature in the remarkable X-ray structure of photosystem II, in which densities for 30 subunits, all of
TICB-903; No. of Pages 8
Review
(a)
Trends in Cell Biology xxx xxxx, Vol. xxx, No. x
p23TMD sphingomyelin
p24TMD sphingomyelin
p24TMD dimer
(b)
VDAC dimer PG
(i)
VDAC monomers CL
(ii)
ADP/ATP carrier dimer CL
(iii)
(c)
CL12(PE)6
TRENDS in Cell Biology
Figure 1. Different biophysical approaches can be used to probe membrane lipid interactions, including fluorescence methods, X-ray crystallography, and mass spectrometry. (a) Probing the effects of lipid binding using Fo¨rster resonance
the metal atoms, and the five oxygen atoms of the Mn4CaO5 cluster were resolved [11]. More than 1300 water molecules were located in each photosystem II monomer together with a series of lipids. Interestingly, all of the PGs identified were distributed with their head groups located in the stromal surface of the membrane. This may suggest that the hydrophilic head groups cannot penetrate the membrane, resulting in their preferential distribution on the stromal side. The lipids appear to be dispersed throughout the structure, rather than in discrete binding pockets, mediating interactions between subunits, suggesting that these are annular rather than structural lipids. A particularly dramatic example of structural lipid binding was revealed in the recent X-ray structure of a short-chain derivative of phosphatidylinositol 4,5-bisphosphate (PIP(2)) in complex with Kir2.2 [12]. PIP(2) is a minor component of cell membranes and is known to regulate many ion channels [13]. The X-ray structure revealed that the PIP(2) lipid binds at an interface between the transmembrane and cytoplasmic domains. On lipid binding, a flexible expansion linker in Kir2.2 contracts to a compact helical structure and the cytoplasmic domains ˚ and become tethered to the transmembrane move 6 A domain. This in turn causes the inner helix gate to open. These results illustrate the role of PIP(2) in controlling the resting membrane potential through this specific ion channel–receptor ligand interaction. Furthermore this study highlights the importance of lipid binding in functional control [12]. An intriguing post-translational modification with consequences for lipid binding was reported recently for the bovine c-subunit of the F-ATP synthase [14]. A lysine group at the C terminus of the transmembrane a-helix was found to be completely trimethylated [14]. The resulting quaternary amino group is therefore exposed to the phospholipid bilayer, resulting in a possible steric clash with the head groups of the phospholipids, impeding their binding to the ring. CL is an essential component of the mitochondrial FATP synthase and could bind to trimethyl-lysine residues at the top of the FO-ring such that its acyl side chains would then be in a position to strengthen the membrane ring by filling the gaps between adjacent subunits. Interestingly, Lys-43 is conserved throughout all known c-subunit sequences in Animalia and it is likely that the trimethylation of this residue is conserved as a mean of stabilising FO energy transfer (FRET) measurements between fluorescently labelled sphingomyelin lipid interacting with the COPI machinery protein p24 transmembrane domain (TMD). The related p23 TMD does not bind to sphingomyelin and remains monomeric [5]. Energy-minimised p24 TMD and p23 TMD models were generated by Modeller9v8 [42] using the P-glycoprotein structure (PDB ID 3G61) as a template [43]. (b) Fluorescence correlation spectroscopy was used to probe labelled voltage-dependent anion channel (VDAC) protein bound to phosphatidylglycerol (PG) and cardiolipin (CL). Dimers were formed in the presence of PG (i) but not CL (ii). VDAC structures are from the X-ray structure of human VDAC (PDB ID 2JK4) [44]. X-Ray crystallography was used to examine the dimerisation of ADP/ATP carrier protein in the presence of CL [9] (iii). (c) Mass spectrometry was used to define the presence of six phosphatidylethanolamine (PE) lipids bound to the CL12 membrane rotor of the Thermus thermophilus (Tt) ATPase. Models for the 6-fold symmetric ring in which six L12 dimers (pink) are stabilised by specific lipid binding (green) and the docking of subunit C (blue) were obtained as described previously [34]. The lipid interaction was modelled between the PE head groups and Glu-63 in the L subunit via PatchDock [45] with further energy minimisation [34].
3
TICB-903; No. of Pages 8
Review
Trends in Cell Biology xxx xxxx, Vol. xxx, No. x
rings. CL has also been found in the Enterococcus hirae (Eh) V-type ATPase and PE in the membrane rotor of Thermus thermophilus (Tt) ATPase (Figure 1c) from mass spectrometry experiments described in the next section.
however, have mass spectrometry approaches been applied to intact membrane assemblies [19–21]. Using electrospray ionisation, an unexpected outcome of this research has been the finding that not only are membrane and soluble subunit interactions maintained, but also specific lipid binding is retained in almost all complexes studied to date [21,22]. This has enabled the identification of lipids bound specifically to protein subunits. Mass spectra have been recorded for several intact transporters, ion channels, and molecular machines, each with specific lipids bound tightly within the complexes (Figure 2). The major advantage of mass spectrometry in this area is in defining simultaneously the mass of the lipid and the
Mass spectrometry developments enable observation of specific binding of lipids Mass spectrometry is a relative newcomer to the membrane protein field, having long been the power behind proteomics. For the past two decades, it has been applied to the study of soluble protein complexes providing insight into their assembly [15], dynamics [16], polydispersity [17], and compositional heterogeneity [18]. Only recently, 100 26+
40+
27+
100
100
MacB 38+
23+
%
24+
%
%
MexB
33+
MtrE average charge
m/z 6500
0 6000
0
m/z
m/z
6000
5000
4000
8000
10000
0
50
LmrCD 40
25+
26+
100
100
KirBac3.1 30
28+
27+
24+
22+ 20
%
%
10
0 6000
5000
100
m/z
m/z
0 0
8+
2 100
EmrE
4
6
21+
0 5000
6000
ASA(A2x104) 23+
22+
100
LmrA
CL12
100
BtuCD
17+
%
%
8
%
%
x6 21+
0 3000
4000
m/z
0 6000
8000
m/z
0 6000
7000
m/z
m/z
5000
0 7000
TRENDS in Cell Biology
Figure 2. Mass spectrometry of various intact membrane protein complexes reveals specific binding of lipids. Mass spectra of four ABC transporters (MacB, LmrCD, LmrA, and BtuCD), three multidrug-resistance pumps (MtrE, MexB, and EmrE), an ion channel (KirBac3.1), and a membrane rotor (CL12 from the Tt ATPase). The average charge state recorded for soluble protein complexes is plotted against their accessible surface area (black line) and compared with the same plot for membrane protein complexes (red line) and a plot without the transmembrane regions included in the surface area calculation (pink line). The transmembrane domains in all protein complexes are coloured red. The lipids that were identified in each case are shown in a red/green space-filling representation [phosphatidylethanolamine (PE) and cardiolipin (CL) with two and four acyl chains, respectively].
4
TICB-903; No. of Pages 8
Review stoichiometry of its binding from measurement of the intact mass of membrane-containing complexes generated in solution or in the gas phase. From the mass of the lipid and its gas phase fragmentation pathway, it is often possible to define the head group, branching, and unsaturation sites. In this review, we focus on recent applications of mass spectrometry to the identification of lipids and the stoichiometry of their binding to membrane complexes, with special attention on their effect on subunit interactions and links to function. The major technological breakthrough that made this possible was the introduction into the gas phase of membrane complexes in protective detergent micelles. The precise mechanism by which these so-called ‘gas-phase micelles’ protect membrane complexes continues to inspire both theoretical simulations [23] and experimental investigation [24]. It is clear that during the phase transition, under the same instrumental set-up, soluble complexes are reduced to monomeric subunits [21]. However, how do the gas phase micelles protect the membrane complexes and what is the mechanism for their dissociation in the gas phase? Recent studies have shown that it is not the solution phase critical micelle value that governs the size of these gas phase micelles, because a plot of the average aggregation number against the alkyl chain length shows that the series from C6 trimethylammonium bromide (TAB) to C16TAB leads to a decrease in the aggregation number in the gas phase. Interestingly, therefore, this observation is in contrast to the established relationship in which increasing chain length leads to larger aggregation numbers in solution [24]. That the longer alkyl chain length is less stable in the gas phase provides a rationale for the choice of DDM over shorter chain detergents for a wide range of protein micelle complexes [21]. Recent proposals for the electrospray mechanism are relevant when considering the dissociation of gas phase micelles. A new view that appears to be well supported by experiment is that protein charging does not take place in the droplet, but rather at the point at which the protein enters the gas phase, after emission of charge carriers during droplet evaporation [25]. This has implications for the folded state of protein complexes, particularly those that have been expelled from micelles. If the micelle is still surrounding the transmembrane regions at the time of charging, these areas, which typically have fewer charges anyway, will not be charged. We investigated the relationship between charge and membrane surface area systematically by plotting the solvent-accessible surface area of several membrane complexes against their average charge state determined experimentally (Figure 2). We compared this relationship with a plot of the charge state of a representative selection of soluble complexes. It is clear that the average charge for membrane complexes is lower than for soluble ones of similar surface area. Interestingly, however, if the surface area attributed to the membrane regions is subtracted from the total surface area, the two lines largely coalesce. This implies that the membrane regions are not charged by virtue of their sequence because of their protection in the detergent micelle. Whatever the origin of this low charge, the absence of detergent molecules from the membrane complex points strongly to their
Trends in Cell Biology xxx xxxx, Vol. xxx, No. x
Box 2. IM of membrane protein complexes IM is an analytical technique that separates ions based on their gasphase mobilities. Application of this technique to membrane protein complexes, after their emergence from detergent micelles in the gas phase, presents a unique opportunity to probe their structure and dynamics, especially in regions that are otherwise wrapped by detergents in solution. In a typical experiment, membrane complexes are electrosprayed directly from micellar solutions and transferred intact into the gas phase protected within detergent micelles. The encapsulated complexes are subsequently released and pulsed into a drift tube filled with inert buffer gas, then migrate under a low electric field. While in the drift tube, ions undergo numerous collisions with the buffer gas molecules. The greater the CCS of an ion, the more gas molecules will collide with it, impeding their progress and thereby increasing the time taken for the ions to migrate through the drift tube. A detector at the end of the drift tube records the arrival times of the ions. These arrival times are then converted to experimental CCS values, which are a measurement of the orientationally averaged projections of a particular ion and is characteristic of its size and shape. When this is coupled with mass spectrometry (IM–MS), information on the size and shape of the ions can be obtained simultaneously. Ions of the same mass but different topology have distinct arrival times on IM. For example, ions with a compact conformation experience fewer collisions in the drift tube and hence travel faster and arrive earlier than those of similar size but with an extended conformation. Moreover, an increase in conformational dynamics and heterogeneity is evident from an expansion of the IM arrival time distribution (ATD) of the ions. This is informative for assessing structural dynamics and conformational heterogeneity in both the soluble and the transmembrane regions of membrane complexes.
role in forming a ‘protective shield’. This protective shield is lost during activation in the mass spectrometer, to reveal well-resolved mass spectra, enabling lipid binding to be defined. The protection afforded by the micelle, however, may not be sufficient to prevent unfolding of the membrane complex in the gas phase. We considered this possibility in a series of ion mobility (IM) experiments (Box 2) in which two membrane complexes were investigated: the transporter BtuC2D2 and the channel KirBac3.1 [26]. Because it is established that activation can lead to unfolding of protein complexes in the gas phase [27], we were keen to determine whether the activation conditions applied to remove detergent were sufficient to unfold the membrane protein complexes. To investigate this, we selected two membrane protein complexes of similar mass and surface area but with different topological arrangements. In the case of KirBac3.1, all four subunits make contact with the membrane, whereas for BtuC2D2 only the two BtuC subunits are protected by the micelle. Clear differences were observed, with the transporter being much less protected than the ion channel by virtue of the surrounding micelle [26]. However, under very low activation conditions, it was possible to obtain relatively compact structures for both complexes, validating the use of IM mass spectrometry for membrane complexes released from detergent micelles. In addition to annular and non-annular or structural lipids, there is a third class that is used as substrates. This is well documented for ABC transporters that interact with membrane lipids and in some cases use phospholipids as substrates [28–30]. In addition, these transporters can require specific lipids to perform substrate transport. The ABC transporter Aus1, reconstituted in proteoliposomes, 5
TICB-903; No. of Pages 8
Review
Trends in Cell Biology xxx xxxx, Vol. xxx, No. x
increases its ATPase activity by direct interaction between the protein and phosphatidylserine [31]. These results suggest strong interactions between lipids and the transmembrane domains of this ABC transporter. Similarly, mass spectra of the ABC transporter MacB revealed the presence of two PE molecules [21]. Mass spectra of two other ABC transporters showed binding of one and two CL molecules to LmrA [32] and LmrCD [21], respectively. Under the conditions used in these experiments, it is likely that annular lipids are absent [33]. Moreover, the absence of lipids associated with monomeric subunits implies that lipid binding occurs primarily within subunit interfaces. Whether these lipids are transported or are required for optimal protein stability is difficult to define in the absence of a functional assay. However, their persistence in various different detergents implies a high specificity of interaction. Different modes of lipid binding revealed by mass spectrometry Turning to the largest of the membrane complexes recently studied by electrospray mass spectrometry, two rotary ATPases/synthases from Tt and Eh, comprising almost 30 subunits and many lipids, were preserved intact in the gas phase [34]. The fact that the subunit–lipid interactions were maintained in these assemblies is particularly interesting, because two very different lipid-binding patterns were uncovered. Using a combination of the mass spectra of the intact species together with liquid chromatography of the individual subunits and gas phase fragmentation of the endogenous lipids, the masses and structures of the lipids could be ascertained. Quantitative proteomics and lipidomics were then used to confirm their relative abundances. For the membrane rotor from Eh ATPase, which contains 10 K subunits each with four transmembrane helices, a series of CLs were identified with a stoichiometry of 1:1 for rotor subunit:CL. This represents a change in the interpretation of the lipids bound to the K ring. These lipids were previously observed in an X-ray crystallographic structure of the isolated ring and assigned to 20 PG molecules based on the high cellular abundance of this lipid [35]. The similar volume occupied by four acyl chains in 10 CLs as opposed to two acyl chains (a)
(b)
Dri me (ms)
22
100
16.91
19.4
100 10.62
17 0
5
20 35
26+
24+
12
7
12-fold symmetry
20 Dri me (ms)
6-fold symmetry
in 20 PG molecules is, however, consistent with two possible assignments to this electron density. The mass of the intact membrane complex together with the lipidomics enables us to delineate these possibilities and to confirm that 10 CLs are located in the orifice of the ring [34]. It should be noted, however, that without the X-ray structure, the location of the 10 CLs would not have been possible from mass spectrometry experiments alone. For Tt ATPase, 12 L subunits each with two transmembrane helices occupy the membrane rotor. Six PE molecules per intact Tt ATPase were identified using quantitative proteomics and lipidomics as well as knowledge of the mass of the intact ATPase and its subcomplexes. Results from the intact mass and the lipidomics were consistent with a stoichiometry of 2:1 for L subunit:lipid interactions. This substoichiometric binding suggests that the 12 subunits undergo rotation to form six dimeric L subunit pairs, each stabilised by specific lipid binding, consequently with four transmembrane helices per subunit (Figure 3 left). This changes the proton:ATP ratio in favour of proton pumping rather than ATP synthesis. Interestingly, this configuration is in accord with the 6fold electron density of the L ring observed previously [36]. This 6-fold symmetric state is also of interest mechanistically because, previously it was established that Tt ATPase can operate both as a proton pump and to synthesise ATP [37]. This led to the proposal that this specific lipid binding switches this rotary ATPase from an ATP synthase to an ion-pumping V-type ATPase [34]. In this mode, pairs of subunits interact and are stabilised by specific lipid binding to produce subunits with four transmembrane helices in line with all other membrane rotors of the V-type ATPases examined to date [35]. Interestingly, the central subunit within all membrane rotors of the rotary ATPases makes remarkably similar contact with subunits in the central stalk to propagate rotation from the membrane rotor to the soluble head. The size of this central subunit in the centre of the rotor is largely conserved, suggesting that the orifices of the membrane rotors should be similar. However, this is clearly not the case. The number of subunits in the ring appears to be fixed for a given species, but this stoichiometry varies
25+
15 0
5
11.49
21+ 22+ 20 35 23+
10
5
21+
m/z
5000 6000 7000 8000 9000
5000 6000 7000 8000 9000 TRENDS in Cell Biology
Figure 3. Ion mobility (IM) mass spectrometry of the Thermus thermophilus (Tt) ATPase membrane-containing subcomplexes. IM arrival time distribution (ATD) (blue) reveals a broadening of the distribution for ICL12 (a) compared with CL12 (b). This broadening is also observed in the extracted ATD peaks (green and blue base, respectively). Models of the 6-fold symmetry (left) stabilised by the binding of six phosphatidylethanolamine (PE) lipids (red) and the membrane rotor with 12-fold symmetry (right). Homology models for the 12-fold and 6-fold symmetry L12 rings were built using the L subunit model described previously [34]. Hybrid atomic and coarsegrained (dark green spheres) models for the I subunit were built based on both previous I subunit modelling and the recent electron microscopy density map [34,39].
6
TICB-903; No. of Pages 8
Review between species [38]. How, then, are the different membrane rings adapted for rotation of the central shaft? Consider the two examples examined here, where in situ lipid binding was identified and molecular modelling of the 12-fold L12 ring based on homology to two transmembrane helices from the Eh K ring structure [35] gave rise to a ˚ . Rotation to form six L dimers (L2)6, central orifice of 43 A with each dimer stabilised by one lipid, reduces the inner ˚ . This compares with the orifice diameter of (L2)6 to 39 A calculated for the K10 membrane rotor of Eh ATPase, in which 10 bulky CLs bind to the internal cavity, reducing it ˚ in the absence of lipids to 39 A ˚ in the presence of from 54 A these lipid brushes. This implies that lipid binding to both membrane rotors reduces the internal cavities to similar values for Eh and Tt ATPases, although the effects are much more marked in the case of the Eh ATPase. To investigate conformational heterogeneity in the rotary ATPases, we used IM mass spectrometry, as outlined above, to probe the collision cross-section (CCS) of various subcomplexes. The broadening of the arrival time distribution peaks for the membrane-embedded Tt ICL12, compared with the CL12 peaks, is consistent with enhanced conformational dynamics of subunit I (Figure 3a,b). Subunit I contains a transmembrane portion which is at 908 to the soluble arm. IM CCS data, together with molecular modelling, suggest that this soluble arm can undergo conformational dynamics from 908 to 1208. This observation, coupled with a proposed nucleotide-binding site in subunit I at the hinge region of the 908 configuration, which binds preferentially to ADP, led us to propose that ADP binding triggers a conformational change. This conformational change in turn causes subunit I to move away from the proton conductance channel formed between the membrane I and the L subunits in the ring, and disrupt the channel. Lipids from the membrane would then seal the gap left by the subunit movement when ATP levels are depleted, and in so doing preserve the proton gradient created at the expense of ATP [34]. Subsequently, a high-resolution electron cryomicroscopy (cryo-EM) structure of the intact Tt ATPase at ˚ was reported. This technological breakthrough shows 9.7 A 12-fold symmetry for the L12 ring [39]; presumably in this case, by contrast to the one above, the Tt ATPase was caught in an ATP synthesis mode (Figure 3 right). Although lipid binding could not be discerned in this cryoEM structure and the detergent micelle contributes to the overall dimensions, when compared with a sixfold model produced from our experimental data, the two rotor rings appear remarkably similar. This would imply that only minor rearrangements in the ring would be enough to trigger the switch between pumping and ATP synthesis. Isolating this rotary Tt ATPase in its two different modes of action and investigating lipid-binding patterns in the two possible forms is now a major goal. In summary, these studies of the two rotary ATPases described here highlight the importance of combining electron microscopy and X-ray crystallography with mass spectrometry. The synergy of these techniques has not only enabled the location of lipids to be defined, but also their identity and stoichiometry to be deduced. In the future, coupling this lipid-binding information with the
Trends in Cell Biology xxx xxxx, Vol. xxx, No. x
location of post-translational modifications will enable their effects on subunit interfaces and lipid binding to be understood. Ultimately, this will add to our understanding of the regulation and control of these fascinating rotary motors. Concluding remarks The diversity of lipid structures that are being uncovered and the beginnings of an understanding of their roles in organising and maintaining membrane protein interactions is sparking considerable interest from structural biologists and lipid chemists alike. Interestingly, lipids that are found in subunit interfaces are often of minimal abundance in the membrane in which the complex is embedded, as was found for the V-type ATPases [40] and for the ion channels that select (PIP)2, a minor component of the cell membrane [41]. This leads to the proposal that complexes select lipids from the available pool for specific tasks rather than using the most abundant lipids. The four transmembrane K subunits of Eh ATPase select lipids with four acyl chains, whereas the two transmembrane L subunits use PE lipids with two acyl chains. This specific tailoring of lipids is also observed for the VDAC dimer, p24, and the ADP/ATP carrier protein, which binds preferentially to PG, sphingomyelin, and CL, respectively. At this stage, it is unclear how these recognition events occur. It is clear, however, that more information is needed before definitive conclusions can be made based not only on structural recognition but also on functional requirements. Increasingly evident is the fact that lipids can no longer be ignored in the structures of membrane complexes; their ability to fine-tune interactions and to stabilise different interfaces is leading to numerous mechanistic insights. With the new biophysical approaches that are coming to the fore to uncover lipid binding, it seems likely that mechanistic understanding of the role of lipids will advance rapidly. In the near future, it is hoped that this will lead to fascinating insights into the role of the diversity of lipid structures and into their correlated binding events. Acknowledgements Funding from Fondo Nacional de Desarrollo Cientı´fico y Tecnolo´gico (FONDECYT) regular grant #1120169 and the Millennium Scientific Initiative (Ministerio de Economı´a, Fomento y Turismo) #P10-035-F (N.P.B.), an ERC advanced grant and the Wellcome Trust (M.Z.), and the Royal Society (C.V.R.) is acknowledged.
References 1 Dowhan, W. (1997) Molecular basis for membrane phospholipid diversity: why are there so many lipids? Annu. Rev. Biochem. 66, 199–232 2 Lee, A.G. (2011) Lipid-protein interactions. Biochem. Soc. Trans. 39, 761–766 3 Adamian, L. et al. (2011) Lipid-binding surfaces of membrane proteins: evidence from evolutionary and structural analysis. Biochim. Biophys. Acta 1808, 1092–1102 4 Lensink, M.F. et al. (2010) Identification of specific lipid-binding sites in integral membrane proteins. J. Biol. Chem. 285, 10519–10526 5 Contreras, F.X. et al. (2012) Molecular recognition of a single sphingolipid species by a protein’s transmembrane domain. Nature 481, 525–529 6 Betaneli, V. et al. (2012) The role of lipids in VDAC oligomerization. Biophys. J. 102, 523–531 7
TICB-903; No. of Pages 8
Review 7 Palmieri, F. (2004) The mitochondrial transporter family (SLC25): physiological and pathological implications. Pflugers Arch. 447, 689– 709 8 Pebay-Peyroula, E. et al. (2003) Structure of mitochondrial ADP/ATP carrier in complex with carboxyatractyloside. Nature 426, 39–44 9 Nury, H. et al. (2005) Structural basis for lipid-mediated interactions between mitochondrial ADP/ATP carrier monomers. FEBS Lett. 579, 6031–6036 10 Trezeguet, V. et al. (2000) A covalent tandem dimer of the mitochondrial ADP/ATP carrier is functional in vivo. Biochim. Biophys. Acta 1457, 81–93 11 Umena, Y. et al. (2011) Crystal structure of oxygen-evolving photosystem II at a resolution of 1.9 A. Nature 473, 55–60 12 Hansen, S.B. et al. (2011) Structural basis of PIP2 activation of the classical inward rectifier K+ channel Kir2.2. Nature 477, 495–498 13 Suh, B.C. and Hille, B. (2008) PIP2 is a necessary cofactor for ion channel function: how and why? Annu. Rev. Biophys. 37, 175–195 14 Watt, I.N. et al. (2010) Bioenergetic cost of making an adenosine triphosphate molecule in animal mitochondria. Proc. Natl. Acad. Sci. U. S. A. 107, 16823–16827 15 Sakata, E. et al. (2011) The catalytic activity of Ubp6 enhances maturation of the proteasomal regulatory particle. Mol. Cell 42, 637–649 16 Uetrecht, C. et al. (2008) High-resolution mass spectrometry of viral assemblies: molecular composition and stability of dimorphic hepatitis B virus capsids. Proc. Natl. Acad. Sci. U. S. A. 105, 9216–9220 17 Stengel, F. et al. (2010) Quaternary dynamics and plasticity underlie small heat shock protein chaperone function. Proc. Natl. Acad. Sci. U. S. A. 107, 2007–2012 18 Ebong, I.O. et al. (2011) Heterogeneity and dynamics in the assembly of the Heat Shock Protein 90 chaperone complexes. Proc. Natl. Acad. Sci. U. S. A. 108, 17939–17944 19 Morgner, N. et al. (2007) A novel approach to analyze membrane proteins by laser mass spectrometry: from protein subunits to the integral complex. J. Am. Soc. Mass Spectrom. 18, 1429–1438 20 Barrera, N.P. et al. (2008) Micelles protect membrane complexes from solution to vacuum. Science 321, 243–246 21 Barrera, N.P. et al. (2009) Mass spectrometry of membrane transporters reveals subunit stoichiometry and interactions. Nat. Methods 6, 585–587 22 Barrera, N.P. and Robinson, C.V. (2011) Advances in the mass spectrometry of membrane proteins: from individual proteins to intact complexes. Annu. Rev. Biochem. 80, 247–271 23 Friemann, R. et al. (2009) Molecular dynamics simulations of a membrane protein-micelle complex in vacuo. J. Am. Chem. Soc. 131, 16606–16607 24 Borysik, A.J. and Robinson, C.V. (2012) Formation and dissociation processes of gas-phase detergent micelles. Langmuir 28, 7160–7167 25 Hogan, C.J., Jr et al. (2008) Charge carrier field emission determines the number of charges on native state proteins in electrospray ionization. J. Am. Chem. Soc. 130, 6926–6927 26 Wang, S.C. et al. (2010) Ion mobility mass spectrometry of two tetrameric membrane protein complexes reveals compact structures and differences in stability and packing. J. Am. Chem. Soc. 132, 15468–15470 27 Ruotolo, B.T. et al. (2007) Ion mobility-mass spectrometry reveals longlived, unfolded intermediates in the dissociation of protein complexes. Angew. Chem. Int. Ed. Engl. 46, 8001–8004
8
Trends in Cell Biology xxx xxxx, Vol. xxx, No. x
28 Doshi, R. et al. (2010) Dissection of the conformational cycle of the multidrug/lipidA ABC exporter MsbA. Proteins 78, 2867–2872 29 Nagao, K. et al. (2010) Lipid outward translocation by ABC proteins. FEBS Lett. 584, 2717–2723 30 van Meer, G. et al. (2006) ABC lipid transporters: extruders, flippases, or flopless activators? FEBS Lett. 580, 1171–1177 31 Marek, M. et al. (2011) The yeast plasma membrane ATP binding cassette (ABC) transporter Aus1: purification, characterization, and the effect of lipids on its activity. J. Biol. Chem. 286, 21835–21843 32 Velamakanni, S. et al. (2009) A multidrug ABC transporter with a taste for salt. PLoS ONE 4, e6137 33 Hite, R.K. et al. (2010) Principles of membrane protein interactions with annular lipids deduced from aquaporin-0 2D crystals. EMBO J. 29, 1652–1658 34 Zhou, M. et al. (2011) Mass spectrometry of intact V-type ATPases reveals bound lipids and the effects of nucleotide binding. Science 334, 380–385 35 Murata, T. et al. (2005) Structure of the rotor of the V-Type Na+ATPase from Enterococcus hirae. Science 308, 654–659 36 Bernal, R.A. and Stock, D. (2004) Three-dimensional structure of the intact Thermus thermophilus H+-ATPase/synthase by electron microscopy. Structure 12, 1789–1798 37 Yokoyama, K. et al. (2000) V-Type H+-ATPase/synthase from a thermophilic eubacterium, Thermus thermophilus. Subunit structure and operon. J. Biol. Chem. 275, 13955–13961 38 Pogoryelov, D. et al. (2012) Engineering rotor ring stoichiometries in the ATP synthase. Proc. Natl. Acad. Sci. U. S. A. 109, E1599–E1608 39 Lau, W.C. and Rubinstein, J.L. (2012) Subnanometre-resolution structure of the intact Thermus thermophilus H+-driven ATP synthase. Nature 481, 214–218 40 Ray, P.H. et al. (1971) Effect of growth temperature on the lipid composition of Thermus aquaticus. J. Bacteriol. 108, 227–235 41 Jiang, Q.X. and Gonen, T. (2012) The influence of lipids on voltagegated ion channels. Curr. Opin. Struct. Biol. 22, 529–536 42 Sali, A. and Blundell, T.L. (1993) Comparative protein modelling by satisfaction of spatial restraints. J. Mol. Biol. 234, 779–815 43 Aller, S.G. et al. (2009) Structure of P-glycoprotein reveals a molecular basis for poly-specific drug binding. Science 323, 1718–1722 44 Bayrhuber, M. et al. (2008) Structure of the human voltage-dependent anion channel. Proc. Natl. Acad. Sci. U. S. A. 105, 15370–15375 45 Schneidman-Duhovny, D. et al. (2005) PatchDock and SymmDock: servers for rigid and symmetric docking. Nucleic Acids Res. 33, W363–W367 46 Fahy, E. et al. (2005) A comprehensive classification system for lipids. J. Lipid Res. 46, 839–861 47 Sud, M. et al. (2007) LMSD: LIPID MAPS structure database. Nucleic Acids Res. 35, D527–D532 48 Subramaniam, S. et al. (2011) Bioinformatics and systems biology of the lipidome. Chem. Rev. 111, 6452–6490 49 Ivanova, P.T. et al. (2009) Lipidomics: a mass spectrometry based systems level analysis of cellular lipids. Curr. Opin. Chem. Biol. 13, 526–531 50 Bligh, E.G. and Dyer, W.J. (1959) A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol. 37, 911–917 51 Harkewicz, R. and Dennis, E.A. (2011) Applications of mass spectrometry to lipids and membranes. Annu. Rev. Biochem. 80, 301–325