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 Católica 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
Förster 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
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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
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
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(a)
Trends in Cell Biology January 2013, Vol. 23, No. 1
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 Förster 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
move 6 Å and become tethered to the transmembrane
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].
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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
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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
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,
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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].
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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
central orifice of 43 Å. Rotation to form six L dimers (L2)6,
with each dimer stabilised by one lipid, reduces the inner
diameter of (L2)6 to 39 Å. This compares with the orifice
calculated for the K10 membrane rotor of Eh ATPase, in
which 10 bulky CLs bind to the internal cavity, reducing it
from 54 Å in the absence of lipids to 39 Å in the presence of
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
9.7 Å was reported. This technological breakthrough shows
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
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 Cientı́fico y Tecnológico
(FONDECYT) regular grant #1120169 and the Millennium Scientific
Initiative (Ministerio de Economı́a, 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.
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