Membrane protein reconstitution in nanodiscs for luminescence spectroscopy studies

Maria E. Zoghbi 1  and Guillermo A. Altenberg 2
  • 1 School of Natural Sciences, University of California, Merced, 4225 N. Hospital Road, Atwater, CA 95301, USA
  • 2 Department of Cell Physiology and Molecular Biophysics, Texas Tech University Health Sciences Center, 3601 4th Street, Lubbock, TX 79430-6551, USA
Maria E. Zoghbi
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  • Maria E. Zoghbi is an Assistant Professor in the Department of Natural Sciences at the University of California, Merced, CA, USA. She was born in Venezuela, where she studied biology at the Central University of Venezuela, and received her PhD degree from the Venezuelan Institute for Scientific Research. She joined Roger Craig’s laboratory at University of Massachusetts Medical School in 2001 for her doctoral thesis and first postdoctoral position. Between 2007 and 2016, she was a senior postdoctoral associate in Guillermo Altenberg’s laboratory at Texas Tech University Health Sciences Center. Her main research interest is to understand the function of proteins, especially membrane proteins, from a biochemical and structural perspective.
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and Guillermo A. Altenberg
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  • Guillermo A. Altenberg has been the head of the Department of Cell Physiology and Molecular Biophysics at Texas Tech University Health Sciences Center, Lubbock, TX, USA since 2014. He received his MD degree from the School of Medicine of the University of Buenos Aires in Argentina, and his PhD degree from the same university in 1987, working in the Institute for Cardiological Research. He has spent most of his academic career as faculty in the School of Medicine of The University of Texas Medical Branch at Galveston, and has been at Texas Tech University Health Sciences Center since 2007. His research interest is the structure-function of membrane transport proteins, with a focus on spectroscopy and nanotechnology applications.
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Abstract

ATP-binding cassette (ABC) exporters transport substrates across biological membranes using ATP hydrolysis by a process that involves switching between inward- and outward-facing conformations. Most of the structural studies of ABC proteins have been performed with proteins in detergent micelles, locked in specific conformations and/or at low temperature. In this article, we present recent data from our laboratories where we studied the prototypical ABC exporter MsbA during ATP hydrolysis, at 37°C, reconstituted in a lipid bilayer. These studies were possible through the use of luminescence resonance energy transfer spectroscopy in MsbA reconstituted in nanodiscs. We found major differences between MsbA in these native-like conditions and in previous studies. These include a separation between the nucleotide-binding domains that was much smaller than previously thought, and a large fraction of molecules with associated nucleotide-binding domains in the nucleotide-free apo state. These studies stress the importance of studying membrane proteins in an environment that approaches physiological conditions.

1 Introduction

Membrane proteins (MPs) represent ~30% of the currently sequenced genomes and are targets of at least 60% of the drugs currently in the market [1], [2]. Understanding the structure and function of MPs not only helps identify the causes and mechanisms of disease but also aids in the design of treatments. Integral MPs are characterized by a hydrophobic membrane-spanning domain that interacts with lipids, and hydrophilic extramembrane domains that interact with aqueous solutions. Many studies of MPs involve extraction of the proteins from cell membranes with amphiphilic molecules. Detergents are the most frequently used MP-supporting platforms because of simplicity and availability, but detergent micelles have very different physicochemical properties compared with lipid bilayers and the characteristics of micellar kinetics, with rapid detergent monomer-micelle equilibrium, reduces MP stability [3], [4], [5]. Although the importance of specific endogenous lipids on the function of some MPs has been established [6], [7], [8], [9], a major role of lipids is related to their contribution to the bulk physicochemical properties of biomembranes, such as curvature, lateral pressure profile, and thickness [10], [11], [12], [13], [14]. These properties are generally preserved in artificial lipid bilayers but are largely absent in detergents and detergent-mimicking alternatives such as peptide surfactants, amphipols, and facial amphiphiles [15], [16], [17]. Bicelles are a different kind of platform that consists of a discoidal structure formed by a mix of phospholipids and detergent, with varying sizes depending on their composition [18]. The planar region of the bicelles mimic a biomembrane; however, bicelle formation is limited to certain lipid compositions.

Reconstitution of MPs into lipid bilayers increases their stability and opens the possibility of using different configurations for different purposes. Examples of well-established bilayer configurations include small and giant unilamellar liposomes, and planar lipid bilayers [19], [20], [21]. An advantage of platforms such as liposomes is that they separate two compartments, which makes possible the study of transport of matter between the compartments. A significant drawback is the limited accessibility to the intraliposomal side for simple binding studies and to study the dynamics of structural changes in response to ligand/substrate binding. The relatively large size of liposomes also complicates optical spectroscopy measurements due to light scattering. Supported planar lipid bilayers display limited access to the extramembrane regions of the MPs at the interface between the lipid bilayer and the solid substrate, and may also present other problems, such as reduced MP stability, because of the interaction between the MPs and the solid support [22]. Nanodiscs (NDs) are proteolipid or polymer-lipid nanostructures that display a great potential as platforms for the study of MPs [23], [24], [25].

2 Nanodiscs

The NDs can replace liposomes favorably in applications in which access to both sides of MPs is advantageous [23], [24], [25]. The NDs consist of two molecules of a membrane scaffold protein (MSP) that encase a small patch of lipid bilayer (Figure 1). The MSP is a genetically modified version of apolipoprotein A1, a major component of the high-density lipoprotein complexes from serum [26]. A picture of the ND structure has emerged from a combination of studies that include size-exclusion chromatography, dynamic light scattering, analytical ultracentrifugation, electron microscopy, nuclear magnetic resonance (NMR) and electron paramagnetic resonance spectroscopy, small angle X-ray scattering, small angle neutron scattering, and molecular dynamics simulations and modeling [23], [24], [27], [28]. Most of the hydrophobic residues of the two MSPs make contact with the hydrophobic part of the lipid bilayer, whereas the hydrophilic residues face the outside, shielding water from the hydrophobic interior, and helping to maintain the NDs soluble in water. One important property of NDs is that their size can be selectively controlled in the ~8 to 17 nm range by modifying the length of the MSP [23], [25], [27], [29].

Figure 1:
Figure 1:

Assembly of nanodiscs. Schematic representation of the self-assembly of nanodiscs from a mixture containing an MSP, phospholipid/detergent mixed micelles (yellow circles: hydrophilic head; blue: phospholipid hydrophobic chains; orange circles: hydrophilic detergent moiety; red: hydrophobic detergent moiety) and detergent-solubilized MP upon detergent removal.

Citation: Nanotechnology Reviews 6, 1; 10.1515/ntrev-2016-0078

NDs assemble spontaneously from mixtures containing MSP and detergent-solubilized lipids upon detergent removal (Figure 1) [23], [24], [25], [26], [30]. A wide variety of phospholipids has been used to produce NDs, and the specific choice depends on several factors, including the requirements of a specific MP [23], [24], [26], [27], [30]. Different MPs have been successfully incorporated in NDs of varied lipid compositions, including synthetic and natural phospholipids [4], [31], [32], [33], [34], [35], [36], [37], [38], [39] and, in some cases, the effects of lipid composition on the MP stability, structure, and/or function was evaluated [24], [36], [37], [38], [39], [40]. However, systematic studies to evaluate potential differences between the effects of lipid composition in liposomes versus NDs are not available. In one study, increasing cholesterol displaced lipids and resulted in a slight ND swelling with the formation of a lens-shaped bilayer [41].

The phospholipid:MSP molar ratio is important, and it depends on the length of the MSP (which determines the ND diameter) and the absence (empty NDs) or presence of MP (MP-loaded NDs) [23], [24], [27]. An efficient reconstitution and homogeneous ND population depends on the use of the proper phospholipid:MSP ratio and incorrect ratios can lead to low reconstitution efficiency, MSP aggregation, and/or formation of large proteolipid complexes [27], [38]. Generally, we consult the available information to estimate the ideal MSP : lipid ratio, but perform experiments with ratios around the ideal value to determine empirically the best reconstitution conditions.

The removal of detergent initiates the formation of the NDs. This can be accomplished in a variety of ways that include dialysis, size-exclusion chromatography, and use of detergent-absorbing columns or beads [23], [24], [25], [26], [27], [30]. We generally use Bio-Beads SM2 resin beads (Bio-Rad, Hercules, CA, USA), which bind a variety of detergents [34]. Details on the expression and purification of MSPs and the ATP-binding cassette (ABC) exporter MsbA have been published [34], [42], and an overview is presented below.

3 Polymer-encased NDs

The most recent variant of the NDs are the polymer-encased NDs known as styrene-maleic acid (SMA)-lipid particles (SMALPs) or Lipodisqs [43]. In these NDs, the MSP is replaced by amphipathic SMA copolymers [44], [45]. There are two obvious and related benefits of using SMALPs. One is that no detergents are used to solubilize the MPs from the membranes. The other is that the resulting SMALP nanostructure contains the native lipids, although this could represent a disadvantage for studies of the effects of specific lipids on MP structure and function. Significant disadvantages of SMALPs are the poor control of their diameters, and the incompatibility of SMA with low pH solutions and some cationic solutes [43]. The latter includes potential problems due to the coordination of SMA carboxylates with the immobilized Ni2+ or Co2+ used for His tag-based MP purification, and SMA precipitation that follows electrostatic association of the carboxylates with cations such as Ca2+ and Mg2+. As a result, functional ATPases that require Mg2+, such as ABC proteins, cannot be studied in a functional state in SMALPs.

4 Uses of NDs in structural and functional studies of MPs

Many MPs, including transport proteins and receptors, have been successfully reconstituted in NDs for structural and functional studies using a variety of biochemical and biophysical techniques [24], [25], [27], [39], [46]. In some studies, including our recent work on MsbA [34], significant improvements in activity and/or structural differences were found in NDs versus detergent micelles. For example, the G protein-coupled β2 adrenergic receptor binds agonists and antagonists and can induce binding of GTP analogs to the G protein G when reconstituted in NDs, but not in detergent micelles [47]. The use of NDs has also allowed for new approaches to investigate difficult problems such as membrane fusion. Intracellular membrane fusion typically involves the zippering of SNARE proteins to facilitate the merger of juxtaposed bilayers. The process involves coiled-coil interactions between SNARE motifs of vesicle-anchored SNAREs (v-SNARE) and target membrane-anchored SNAREs (t-SNARE) that lead to the formation of a complex [29]. This complex is believed to assemble in multiple steps that are mechanically coupled to membrane remodeling, but the details of SNARE zippering have been hard to dissect. However, a transient SNARE complex intermediate has been recently trapped in a ND sandwich [48]. Combining NDs containing either v-SNAREs or t-SNAREs results in the formation of a SNARE complex that can be studied [48]. The rigid structure of the NDs prevents membrane fusion, allowing for the analysis of the trapped transient SNARE intermediate using techniques such as electron paramagnetic resonance spectroscopy in combination with site-directed spin labeling, and single molecule Förster (or fluorescence) resonance energy transfer (FRET). These studies have located a structural hinge at a conserved region near the center of the complex, and have demonstrated equilibrium between half-zippered and fully zipped states [48]. This novel ND-based approach provides the opportunity to probe the role of auxiliary proteins and other regulatory factors in the formation of SNARE complexes.

Because the size of the NDs depends largely on the MSP, under optimal reconstitution conditions MP-loaded NDs constitute a homogeneous and monodisperse population of proteolipid nanostructures that can be treated as soluble proteins. This is particularly advantageous for luminescence spectroscopy, NMR spectroscopy, and single-particle electron cryo-microscopy (cryo-EM) studies. Our studies using luminescence spectroscopy are discussed below. Here, we mention a few examples to highlight the potential of NDs in solution NMR spectroscopy and cryo-EM studies. Several MPs in NDs have been studied by solution NMR, including ion channels (KcsA, KvAP, VDAC-1, and VDAC-2), G protein-coupled receptors (CXCR1 and μ-opioid receptors), bacteriorhodopsin, bacterial outer MPs (OmpA and OmpX), Integrin αIIb, viral proteins (Pf1, p7), the endoplasmic reticulum Ca2+ sensor protein STIM1, and the Cytb5-CytP450 complex [39]. For solution NMR, the use of smaller NDs (6- to 8-nm diameter) reduces the rotational correlation time and increases spectral resolution and sensitivity [39]. The molecular weight of MP-ND complexes is relatively high (generally <200 kDa) but, in many cases, still suitable for solution NMR spectroscopy studies. The first structure solved by NMR spectroscopy was that of the outer membrane OmpX [29]. Compared with the structure in detergent, each strand of OmpX in NDs was shorter, a difference attributed to the slightly denaturing effect of the detergent at the water boundary and differences in hydrophobic coverage. Another example of a relevant difference between the structures of a protein in NDs versus detergent is the SNARE protein synaptobrevin [48]. Unstructured and flexible, and therefore accessible, SNARE motif residues were found with the protein in NDs, but not in dodecylphosphocholine micelles, which points to NDs as more physiologic platforms than micelles.

Recent advances in the development of direct electron detectors and data analysis have made it possible to obtain atomic-resolution structures of proteins by cryo-EM [49]. A clear example of the usefulness of combining NDs and cryo-EM is the recent structure of the heat- and capsaicin-activated ion channel TRPV1 in different states [50]. This recent study shows the location of annular and regulatory lipids, and led to the proposal of a mechanism by which phosphatidylinositides regulate the activity of the channel. Cryo-EM of MP-ND complexes is undoubtedly one of the most promising areas in MP research. A recent review by Denisov and Sligar [25] includes other examples of the use of NDs for structural and functional studies of MPs. Here, we will emphasize how spectroscopic studies of ABC transporters reconstituted in NDs can contribute to answering important questions in the field with repercussions for MP in general.

5 ABC proteins

ABC proteins constitute one of the largest families of MPs, and are found in all domains of life [51], [52], [53]. Most are transporters that use the energy of ATP hydrolysis to translocate substrates across membranes [51], [52]. Most prokaryotes’ ABC proteins are importers, whereas most ABC proteins from eukaryotes are exporters [51], [52]. The basic functional and structural unit of ABC exporters consists of two predominantly helical transmembrane domains that include the substrate-binding pocket and form the translocation pathway, and two nucleotide-binding domains (NBDs) [51], [52], [53]. The NBDs bind and hydrolyze ATP, are structurally conserved, and provide the name of the protein family [51], [52], [53]. ATP binding leads to the formation of an NBD dimer where the NBDs are in a head-to-tail orientation, and hold two ATP molecules at the interface (Figure 2, right side) [54], [55]. Each of the two nucleotide-binding sites is formed by residues from both NBDs, making the formation of this dimer essential for ATP hydrolysis [51], [54], [55]. Recent studies suggest that occupation of the two nucleotide-binding sites is necessary to form a stable dimer that can perform hydrolysis [56], but dissociation of the dimers can take place following ATP hydrolysis at only one of the two sites [57]. ATP binding/hydrolysis by the NBDs, and the consequent association/dissociation of the NBDs, is coupled to rearrangements of the transmembrane helices that switch the transporters from the inward- to the outward-facing conformation needed to accomplish the translocation of substrates (Figure 2) [51], [53].

Figure 2:
Figure 2:

Alternate accessibility model with large conformational changes. Inward- and outward-facing MsbA X-ray structures. Binding of nucleotide (orange spheres) produces an association of the nucleotide-binding domains, which results in the rotation of transmembrane helices and switching of the opening of the substrate-binding pocket from one side of the membrane to the other. Each MsbA monomer is shown in a different color and the positions of the C561 residues are shown as red spheres. The approximate boundaries of the membrane are shown by the lines, and a transported substrate is shown as a red oval. In the case of MsbA, the substrate is lipid A, which the protein translocates from the inner to the outer leaflet of the membrane.

Citation: Nanotechnology Reviews 6, 1; 10.1515/ntrev-2016-0078

The bacterial ABC transporter MsbA has been widely used as a model for studies of the molecular mechanism of ABC exporters [58], [59], [60], [61], [62], [63], [64], [65], [66], [67], [68], [69]. MsbA is an ABC exporter that operates as a lipid flippase, using the energy of ATP hydrolysis to translocate lipid A from the inner to the outer leaflet of the inner membrane of Gram-negative bacteria [70], [71], [72]. Lipid A is a component of the endotoxin responsible for the toxicity of Gram-negative bacteria [73]. MsbA is a homologue of the multidrug resistance protein P-glycoprotein, also known as MDR1 or ABCB1, which has a role in the resistance of some forms of cancer to chemotherapeutic agents [53]. The polyspecific P-glycoprotein is present in the small intestine, kidneys, liver, and blood-brain barrier, where it transports a wide variety of chemically unrelated drugs [53], [74]. Therefore, P-glycoprotein is also an important determinant of the pharmacokinetics of many drugs [53], [75], [76], [77], [78], [79], [80]. The crystal structures of MsbA and P-glycoprotein in the absence of lipids and double electron-electron resonance experimental distances of reconstituted MsbA support the prevalent view that suggests large conformational changes when the exporter switches between the inward- and outward-facing conformations (Figure 2) [66], [67], [68], [69], [81], [82], [83]. Other data suggest instead that the NBDs remain in contact at all times during the hydrolysis cycle, and that such large conformational changes might not be physiological [84], [85], [86], [87], [88]. Many of the discrepancies can be explained by the nonphysiological conditions of the experiments (absence of bilayer, low temperatures, and protein locked in a particular conformational state). We have been using MsbA to develop a more physiologically relevant experimental approach to study ABC exporters [34], [42].

6 Luminescence resonance energy transfer

Luminescence resonance energy transfer (LRET) is a highly sensitive spectroscopic technique based on energy transfer from a donor to an acceptor that can be used to study MPs during their functional operation [34], [42], [56], [89], [90], [91]. LRET is somewhat similar to the traditional FRET, where an excited donor transfers energy to a nearby acceptor by a process that is highly dependent on the donor-acceptor distance (Figure 3A). Both FRET and LRET can be used as molecular rulers, but LRET has some advantages for MP studies. These advantages depend on the distinctive properties of the luminescent lanthanide (Tb3+ or Eu3+) that is used as an LRET donor [91]. First, the light emitted by the LRET donors is atomic-like, with sharp peaks separated by dark regions (Figure 3B), which allows for easy isolation of the acceptor emission without contamination from the donor emission [34], [42], [91], [92], [93]. This is clearly different in FRET, where there is an unavoidable overlap of the broad spectra of the donor and acceptor. Second, the most interesting and useful property of Tb3+ or Eu3+ is the long lifetime of their excited state; whereas traditional fluorophores display nanosecond lifetimes (e.g. ~3 ns for fluorescein), the luminescent lanthanides excited-state lifetimes are 105 to 106 times longer (1–2 ms). This property allows for delayed acquisition of the emission of the LRET fluorophore acceptor where the detector can be gated with a delay of tens to hundreds of microseconds after the excitation pulse (Figure 4) [91]. In LRET, the luminescent lanthanide is excited with a short pulse from a laser or a flash lamp, and the long-lifetime emission from the LRET donor and the acceptor sensitized emission can be recorded 20 to 200 μs after the excitation pulse [34], [42], [56], [89], [91], [93]. The sensitized emission is the long-lifetime emission from the LRET acceptor that can only arise from energy transfer, e.g. fluorescein emission with approximately millisecond-lifetimes, as opposed to its ~3-ns lifetime in the absence of LRET [91]. During the delay period after the excitation pulse, all the light that arises from processes with lifetimes in the nanosecond range decay to negligible values, yielding a very low background signal [91]. These processes include sample autofluorescence, emission from the donor due to direct excitation, and scattering of the excitation pulse. The latter is frequently the most problematic in MP spectroscopic research because of the size of structures such as liposomes and detergent micelles, and a much lower extent MP-loaded NDs.

Figure 3:
Figure 3:

(A) Schematic representation of energy transfer between donor and acceptor probes. The process of energy transfer is highly dependent on the donor-acceptor distance. Energy transfer results in decreased donor emission, increased acceptor emission and shorter lifetime of the excited state of the donor molecules that participate in energy transfer. In LRET, the latter is essentially identical to the long lifetime emission of the fluorophore acceptor (sensitized emission). (B) Emission spectra of LRET donor (Tb-only) and donor-acceptor (Tb-fluorescein). When the emission is measured in gated mode, acceptor emission (fluorescein in this case) due to direct excitation is very low (acceptor only, black trace). This is expected from the short duration of the excitation pulse and the short lifetime of the fluorescein excited state (~3 ns). Tb3+ emits in sharp peaks with interposed dark regions (donor only, blue trace), and therefore a band-pass emission filter easily isolates the acceptor sensitized-emission (emission with long lifetime) from the luminescence of the lanthanide complex. The emission peak at ~520 nm (red trace) is the sensitized emission from fluorescein (long-lifetime acceptor emission arising from energy transfer), and exhibits microsecond lifetimes. Intensity and lifetime measurements that we usually perform are listed. Modified from Bao et al. [93].

Citation: Nanotechnology Reviews 6, 1; 10.1515/ntrev-2016-0078

Figure 4:
Figure 4:

Advantages of LRET and basic experimental protocol. Representative illustration of typical LRET data showing gated emission spectra with a microsecond delay after a 337-nm excitation pulse. Major features of LRET and the associated advantages are listed.

Citation: Nanotechnology Reviews 6, 1; 10.1515/ntrev-2016-0078

In LRET, the relevant long-lifetime emissions are those from the donor Tb3+ or Eu3+ and the sensitized emission of the LRET acceptor fluorophore [91]. Distances between the donor and acceptor probes are calculated from

E=1τDA/τD

R=R0(E11)1/6

where E is the efficiency of energy transfer, R0 is the Förster distance (the distance at which E=0.5), and τD and τDA are the lifetimes of the donor in the absence and presence of the acceptor, respectively. Because the sensitized acceptor emission arises from energy transfer, it can be used as a surrogate of the lifetime of the molecules of the donor that participate in energy transfer, making τDA equal to the sensitized emission lifetime [91]. In addition, the use of sensitized emission makes the calculation of donor-acceptor distances from LRET lifetimes independent of the labeling stoichiometry, i.e. the more LRET donor-acceptor pairs, the higher the sensitized emission intensity, but lifetimes stay the same [89], [91], [93]. Another advantage of LRET is that the emission from the donor and acceptor is unpolarized, minimizing problems caused by orientation factors, which introduce uncertainties in FRET-based calculations [42], [89], [91], [93]. Although both FRET and LRET can monitor distances with atomic resolution, the LRET features described previously allow for more accurate calculation of distances over a wide range, from 25 to 120 Å, depending on the choice of the fluorescent acceptor, and make it more suitable for the kinetic analysis of moving domains in MPs [42], [56], [91].

If one wants to study how different parts of a protein move with respect of each other, it is essential to control the location of the probes within the protein with high accuracy. For LRET, this can be accomplished in different ways. The most common is to use recombinant proteins with two residue side chains that can be labeled selectively with the donor and acceptor probes. This is what we regularly do for our MsbA experiments [34], [42]. We first generated a Cys-less MsbA, in which all native Cys residues were replaced with Ala, and then introduced a single Cys in the position of interest [42]. Because the structural/functional MsbA unit is a homodimer, we have two equivalent residues, one in each MsbA monomer, available for labeling. We have shown that the Cys-less MsbA and several single Cys MsbA mutants display normal ATPase activity even when the Cys are labeled with LRET probes [42]. Several thiol-reactive fluorophores suitable as LRET acceptors are commercially available. As a donor, we use a Tb3+ chelate DTPA-cs124-EMPH developed by Ge and Selvin [94]. The compound contains a maleimide for covalent reaction with the Cys thiol, a carbostyril (cs124, 7-amino-4-methyl-2(1H)-quinolinone) antenna chromophore that absorbs well at 335 nm, and the high-affinity diethylenetriaminepentaacetic acid (DTPA) chelator. Luminescent lanthanides have low extinction coefficients, but the cs124 antenna absorbs the excitation light and transfers it to the lanthanides efficiently, whereas the chelator shields the lanthanides from the quenching effect of water [91], [94]. This compound (thiol-reactive Lanthascreen, Thermo Fisher, Bellfonte, PA, USA) is water soluble, displays millisecond-lifetimes, high-quantum yields, large Stokes shifts, and emission spectra with sharp peaks [94]. One potential problem is the size of the probe that places the lanthanide away from the Cys side chain. This, however, is a minor problem because the distances determined in several systems are very close to those estimated from crystal structures [34], [42], [89], [90], [95], and the position of the probe can also be modeled [96], [97], [98]. Genetically encoded lanthanide-binding sequences are an enticing alternative to chemical labeling. Sequences derived from the EF-hand motifs of Ca2+-binding proteins with high affinity for lanthanides have been developed by Imperiali and collaborators and have been introduced in proteins for LRET studies [99], [100], [101], [102], [103]. These tags include acidic coordination residues and a Trp residue that functions as an antenna [99], [100]. Although these tags are not as efficient as the chemical probes for LRET, they are genetically encoded and therefore they can be used as specific lanthanide binders that minimize the need to purify the proteins. In principle, since the LRET donor binds only to the recombinant protein and the donor-acceptor distance for efficient LRET is rather short, the use of the genetically encoded tags allows work with unpurified preparations, including cells, providing that the lanthanides can access the tag [101], [102], [103].

7 Preparation of MsbA-loaded NDs for LRET measurements

Although we have studied many MsbA mutants, in our published studies, we mostly used the Salmonella typhimurium single-Cys MsbA mutant T561C [34], [42]. This Cys is located near the C-terminal of MsbA, in a region of the NBDs that is expected to undergo a large conformational change during the ATP hydrolysis cycle (Figure 2) [82]. Cys561 is on the outside of the NBD and away from the active site [82] and it is, therefore, accessible to labeling, and its modification does not affect MsbA ATPase activity [42]. We have incorporated a His tag sequence at the N-terminal end of MsbA, expressed the protein in Escherichia coli, solubilized it from the membrane with 1% dodecylmaltoside, and then purified it by affinity chromatography based on the affinity for Co2+ of the His tag [34], [42]. We generally store aliquots of purified MsbA at −80°C in 100 mm NaCl, 20 mm Tris/HCl pH 7.5, 0.065% dodecylmaltoside, 0.04% sodium cholate, and 15% glycerol, in the presence of the reducing agent tris(2-carboxyethyl)phosphine (TCEP; 0.2 mm). In most studies, we label the purified T561C simultaneously with a 2× molar excess of the thiol-reactive Tb3+-chelate DTPA-cs124-EMPH as donor, and N-(2-aminoethyl)maleimide (Bodipy FL maleimide) as acceptor. With this labeling protocol, random labeling seems reasonable because the reaction of both probes is based on the same chemistry, and the same residue is labeled in each T561C MsbA monomer. Therefore, it is expected that 50% of the MsbA homodimers will be labeled with one donor and one acceptor, whereas 25% will be labeled only with donors or acceptors. The LRET signal will then arise only from the MsbA homodimers labeled with one donor and one acceptor. After labeling the protein in detergent, we remove the free unreacted probes by gel filtration on Superdex 200 10/300 GL column (GE Healthcare, Marlborough, MA, USA).

For reconstitution of purified MsbA into NDs, we combined lipids with MSP and labeled MsbA (Figure 1). We have used MSP1E3D1, a MSP variant that produces NDs with a diameter of ~11 nm, which fit one MsbA per ND. The presence of one MsbA homodimer per ND was confirmed by the combination of size estimations by size-exclusion chromatography [34] and dynamic light scattering, a MSP/MsbA molar ratio of 2, and direct observation by transmission electron microscopy (unpublished data). These NDs do not constrain the conformational changes of MsbA [34]. MSP1E3D1 is expressed in Escherichia coli and contains a His tag sequence for purification by affinity chromatography, essentially as described previously for MsbA. The MSP1E3D1 has a tobacco etch virus (TEV) protease site used to remove the His tag after purification. We have been successful using a mix of MsbA, E. coli total lipid extract solubilized in a standard salt buffer with 100 mm sodium cholate, and MSPE3D1 at a MsbA : MSP and MSP : lipid molar ratios of 1 : 8 and 1 : 110, respectively. The mix is incubated for 1 h at 4°C with gentle rotation, and the detergent is removed by overnight incubation with Biobeads SM-2 (Figure 1) [34]. This procedure yields a mixture of empty and MsbA-loaded NDs. An enriched fraction of MsbA-loaded NDs with a hydrodynamic radius of 13.2±0.7 nm (mean±SEM; measured by dynamic light scattering) can be obtained by size-exclusion chromatography on a Superdex 200 10/300 GL column [34]. This process also removes free unreacted probes when labeling is performed after reconstitution. When desired, MsbA-loaded NDs can be separated more efficiently from empty NDs by affinity chromatography based on the interaction of Co2+ or Ni2+ with the His tag of MsbA [34].

LRET measurements are performed using ~0.5 μm MsbA in 3 mm pathlength quartz cuvettes [34], [42]. Early on, we performed the measurements using a nitrogen laser-based system [93], but these days we use simpler instruments [34], [42], [57], [89]; an Optical Building Blocks phosphorescence lifetime photometer (EasyLife L, OBB, Birmingham, NJ, USA) for the intensity and lifetime measurements, and a Photon Technology International spectrometer (QM3SS, PTI, London, Ontario, Canada) to record LRET spectra. We generally use a 200-μs delay from the beginning of an ~1-μs excitation pulse from a Xe flash lamp. For the intensity measurements, we use narrow band-pass filters for the 335-nm excitation and we collect the Tb3+ and Bodipy FL emitted light with centers at 490 and 520 nm, respectively. We have measured changes in LRET intensity to determine steady-state kinetics (e.g. affinity for ATP), time course of changes in LRET intensity to follow conformational changes in real time (e.g. to follow conformational changes during the hydrolysis cycle), and lifetimes of the donor in the absence of acceptor and lifetime of the Bodipy FL sensitized emission to determine movements of the NBDs during the hydrolysis cycle with angstrom resolution (Figure 3B) [34], [42], [57], [89], [93]. Combined with a stopped-flow device, LRET can provide dynamic information on conformational changes in the millisecond-to-minute time frame [34], [42], [57].

8 Activity and structural changes measured in an ABC protein reconstituted in NDs

MsbA has been studied using a variety of approaches, and crystal structures in the inward- and outward-facing conformations are available (Figure 2) [58], [59], [60], [61], [62], [63], [64], [65], [66], [67], [68], [69], [82]. Because of technical limitations and simplicity, most of the structural studies of ABC exporters have been performed with the proteins solubilized in detergent, locked in specific conformations, and/or at low temperature [61], [62], [63], [64], [65], [66], [67], [68], [69]. Studies under more physiological conditions are, however, essential to understand the structure and function of MPs. These include reconstitution into lipid bilayers (the native MP environment), use of physiological temperatures (37°C for most mammalian MPs), and studies of the MP while it is functioning, in addition to the usual measurements with the MP locked in specific conformations. For ABC proteins, the latter are in the nucleotide-free (apo), ATP-bound, and posthydrolysis state (obtained by inhibition with orthovanadate). To accomplish the goal of studying ABC exporters under more physiological conditions than usual, we have assayed the structure, function, and conformational changes of MsbA reconstituted in NDs using LRET, while the protein performs hydrolysis at 37°C [34].

One advantage of our LRET studies in NDs is that they allow for a straightforward comparison of the same MsbA preparation at 37°C in the lipid bilayer versus detergent. Not surprisingly, the ATPase activity of MsbA in NDs was higher than that of MsbA in detergent, and was also more stable and less sensitive to temperature changes [34]. Increased ATPase activity of MsbA reconstituted in NDs compared with the values in detergent was originally reported by Kawai et al. [104]. However, the quality of the preparation was far from ideal, with reported activities ~30-fold lower than ours. Using LRET, we have been able to determine changes in donor-acceptor distances not only with the protein locked in particular states during the hydrolysis cycle (as it is usually done) but also while MsbA was hydrolyzing ATP. We have also been able to assess the distribution of distances in any particular condition and the percentage of molecules in each conformation [34], [42]. Figure 5 shows representative sensitized emission LRET decays of MsbA reconstituted in NDs (MsbA-NDs, main panel) and MsbA in detergent (MsbA-detergent, inset) in the absence of nucleotide (apo), after the addition of NaATP (ATP-bound; promotes formation of NBD dimers that cannot hydrolyze ATP), and after the addition of Mg2+ (MgATP; Mg2+ is a cofactor required for ATP hydrolysis by ABC proteins). The most obvious difference between the decays measured from MsbA in ND versus detergent is that the increase in LRET intensity produced by switching from the apo state (black) to the ATP-bound state (red) was much smaller in the MsbA-NDs. Also, during hydrolysis (blue), the sensitized emission decay is close to that of the apo state in the MsbA-NDs (black), which is not the case for MsbA-detergent. Analysis of the LRET decays from MsbA-ND and MsbA-detergent indicates that each decay can be fit by a two-exponential function; i.e. the decays can be accounted by two donor-acceptor distances in every experimental condition. These two distances stay essentially the same during the ATP hydrolysis cycle (Figure 6A) [34], [42]. The shorter distance (~36 Å) is very similar in MsbA-ND and MsbA-detergent [34], [42], and corresponds to the outward-facing, nucleotide-bound closed conformation (Figure 2) [82]. The longer distance in the apo state is shorter in MsbA-ND than in MsbA-detergent (47±1 Å vs. 53±1 Å) [34], [42], and is much shorter than the corresponding distance in the inward-facing MsbA apo structure based on X-ray crystallography (>80 Å; Figure 2) [82]. Because the extent of the NBD movements in MsbA is smaller in NDs when compared with MsbA in detergent, an obvious question is whether this difference depends on constraints imposed on the MP by the ND size. This possibility was ruled out because similar results were obtained in MsbA reconstituted in ~100-nm diameter unilamellar liposomes [34], [42]. Therefore, the large difference between the NBD-NBD distances in MsbA-NDs and the apo crystal structure [82] is clearly not the result of MsbA confinement in the NDs.

Figure 5:
Figure 5:

Comparison of MsbA in nanodiscs with MsbA in detergent. The MsbA T561C mutant was used in the experiments. Sensitized Bodipy FL emission decays during the ATP-hydrolysis cycle from MsbA in nanodiscs (MsbA-NDs; main panel) and detergent (MsbA-detergent; inset). Black, apo; red, ATP-bound; blue, MgATP (continuous hydrolysis). Intensities are normalized to the ATP intensity at 200 μs. Modified from Zoghbi et al. [34].

Citation: Nanotechnology Reviews 6, 1; 10.1515/ntrev-2016-0078

Figure 6:
Figure 6:

Comparison of MsbA in nanodiscs with MsbA in detergent. (A) Donor-acceptor distances of MsbA in nanodiscs during the hydrolysis cycle. Distances calculated in different states during the hydrolysis cycle from the LRET sensitized Bodipy FL emission decays in MsbA-NDs (means±SD). Possible MsbA structures associated with each distance are shown on the right with the monomers in the MsbA homodimer shown in blue and cyan; the position of the labeled Cys-561 is shown in red. (B) Percentage of MsbA molecules with associated nucleotide-binding domains during the hydrolysis cycle (MsbA molecules displaying the shorter distance; ~36 Å). Data are means±SD. The experimental conditions were: apo, nucleotide-free state obtained in nominally divalent cation-free buffer with 1 mM EDTA to chelate trace divalent cations and prevent ATP hydrolysis; ATP-bound, ATP-bound state obtained after the addition of 1 mm NaATP; MgATP, continuous hydrolysis state obtained after the addition of 10 mm MgSO4; and MgATP + Vi, high-energy posthydrolysis state obtained by the addition of 0.25 mm orthovanadate in the presence of MgATP. Modified from Zoghbi et al. [34].

Citation: Nanotechnology Reviews 6, 1; 10.1515/ntrev-2016-0078

The differences in LRET intensity decays observed during the hydrolysis cycle (Figure 5) are not caused by changes in distances (i.e. different conformations), but are mainly due to changes in the percentage of molecules in each conformation. To illustrate this point, Figure 6B shows the proportion of molecules adopting the 36-Å conformation during the ATP hydrolysis cycle for MsbA-NDs (black) and MsbA-detergent (red). The percentage of molecules in each conformation can be determined from the fit pre-exponential terms corresponding to each donor-acceptor decay and the rate of energy transfer [34], [42]. In the apo state, the proportion of molecules in the 36-Å conformation was much higher in MsbA-ND (48% vs. 7%). This higher proportion of molecules displaying the short donor-acceptor distance in the apo state explains most of the higher LRET intensity in MsbA-ND versus MsbA-detergent (Figure 5) [34]. As expected, ATP shifts the equilibrium toward a larger proportion of molecules adopting the 36-Å conformation (Figure 6B) [34], [42]. When MsbA is performing ATP hydrolysis in the presence of MgATP, the percentage of MsbA molecules displaying the shorter distance decreased to ~50% (Figure 6B). During hydrolysis, NBDs dissociate and reassociate, yielding a steady-state with approximately half of the molecules in each conformation [34], [42], [89].

The LRET intensity decays can also be analyzed by an exponential series method that can recover distance distributions. This type of analysis can be used to discriminate between discrete distributions of donor-acceptor distances, and continuous distributions between conformations [105], [106]. The clear peaks around the distances calculated from the multiexponential fits confirm the presence of discrete conformations in MsbA-ND in the apo state (Figure 7, black) [34]. This is also the case in all other states studied, except for MsbA-ND during ATP hydrolysis (Figure 7A, red), in which the distance distribution was broader [34]. This broadening suggests that during hydrolysis MsbA adopts a wider range of conformations around the 36-Å distance. This behavior was not present in MsbA-detergent (Figure 7B, red), which also showed MsbA sampling much longer distances in the apo state (Figure 7B, black), suggesting large flexibility of the apo MsbA in detergent micelles [34]. From the observation of the distance distributions in Figure 7, it is also apparent that MsbA is much more compact in NDs versus detergent.

Figure 7:
Figure 7:

Distance distributions in the apo state and during hydrolysis. Distance distributions calculated from the lifetime distributions of LRET sensitized emission intensity decays in the apo state and during ATP hydrolysis. Black, apo; red, continuous ATP hydrolysis in the presence of Mg-ATP. (A) Distance distributions in MsbA-NDs. (B) Distance distributions in MsbA-detergent.

Citation: Nanotechnology Reviews 6, 1; 10.1515/ntrev-2016-0078

In summary, the differences in LRET intensities and decays between MsbA-ND and MsbA-detergent are the result of (1) a much larger fraction of NBDs associated or very near to each other in the apo MsbA-ND, and (2) the replacement of the longer distance of fully dissociated NBDs in MsbA-detergent by a shorter distance compatible with loosely associated NBDs or NBDs dissociated but very close to each other. Our conclusion that the catalytic cycle of MsbA in a lipid bilayer proceeds with small conformational changes agrees with a FRET study of P-glycoprotein in liposomes at 37°C, as well as with an electron-microscopy study of P-glycoprotein in a lipid bilayer that showed the NBDs near the apo state, as well as with several computational studies [84], [85], [86], [87], [88].

9 General conclusions and perspectives

In general, our results suggest that in MsbA reconstituted in a lipid bilayer and at physiological temperature, the conformational changes during the ATP hydrolysis cycle are smaller (NBDs separated at most by ~10 Å) than those expected from prior studies. The increased sampling distance of MsbA in detergent in the apo state (Figure 7B, black) suggests that under certain experimental conditions (e.g. crystallization conditions), it may be possible to lock the protein in rare conformations where the NBDs are far apart from each other.

Our studies of MsbA in NDs assess structural changes in an ABC exporter under native-like conditions, and show major differences with the widely open inward-facing crystal structure and also with other studies performed under less-physiological conditions, including our LRET studies of MsbA in detergent. The results stress the importance of performing structural/functional studies of MPs under native-like conditions that include insertion into lipid bilayers and normal temperatures.

Acknowledgements

This work was supported by the Cancer Prevention & Research Institute of Texas grant RP101073.

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    Assembly of nanodiscs. Schematic representation of the self-assembly of nanodiscs from a mixture containing an MSP, phospholipid/detergent mixed micelles (yellow circles: hydrophilic head; blue: phospholipid hydrophobic chains; orange circles: hydrophilic detergent moiety; red: hydrophobic detergent moiety) and detergent-solubilized MP upon detergent removal.

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    Alternate accessibility model with large conformational changes. Inward- and outward-facing MsbA X-ray structures. Binding of nucleotide (orange spheres) produces an association of the nucleotide-binding domains, which results in the rotation of transmembrane helices and switching of the opening of the substrate-binding pocket from one side of the membrane to the other. Each MsbA monomer is shown in a different color and the positions of the C561 residues are shown as red spheres. The approximate boundaries of the membrane are shown by the lines, and a transported substrate is shown as a red oval. In the case of MsbA, the substrate is lipid A, which the protein translocates from the inner to the outer leaflet of the membrane.

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    (A) Schematic representation of energy transfer between donor and acceptor probes. The process of energy transfer is highly dependent on the donor-acceptor distance. Energy transfer results in decreased donor emission, increased acceptor emission and shorter lifetime of the excited state of the donor molecules that participate in energy transfer. In LRET, the latter is essentially identical to the long lifetime emission of the fluorophore acceptor (sensitized emission). (B) Emission spectra of LRET donor (Tb-only) and donor-acceptor (Tb-fluorescein). When the emission is measured in gated mode, acceptor emission (fluorescein in this case) due to direct excitation is very low (acceptor only, black trace). This is expected from the short duration of the excitation pulse and the short lifetime of the fluorescein excited state (~3 ns). Tb3+ emits in sharp peaks with interposed dark regions (donor only, blue trace), and therefore a band-pass emission filter easily isolates the acceptor sensitized-emission (emission with long lifetime) from the luminescence of the lanthanide complex. The emission peak at ~520 nm (red trace) is the sensitized emission from fluorescein (long-lifetime acceptor emission arising from energy transfer), and exhibits microsecond lifetimes. Intensity and lifetime measurements that we usually perform are listed. Modified from Bao et al. [93].

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    Advantages of LRET and basic experimental protocol. Representative illustration of typical LRET data showing gated emission spectra with a microsecond delay after a 337-nm excitation pulse. Major features of LRET and the associated advantages are listed.

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    Comparison of MsbA in nanodiscs with MsbA in detergent. The MsbA T561C mutant was used in the experiments. Sensitized Bodipy FL emission decays during the ATP-hydrolysis cycle from MsbA in nanodiscs (MsbA-NDs; main panel) and detergent (MsbA-detergent; inset). Black, apo; red, ATP-bound; blue, MgATP (continuous hydrolysis). Intensities are normalized to the ATP intensity at 200 μs. Modified from Zoghbi et al. [34].

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    Comparison of MsbA in nanodiscs with MsbA in detergent. (A) Donor-acceptor distances of MsbA in nanodiscs during the hydrolysis cycle. Distances calculated in different states during the hydrolysis cycle from the LRET sensitized Bodipy FL emission decays in MsbA-NDs (means±SD). Possible MsbA structures associated with each distance are shown on the right with the monomers in the MsbA homodimer shown in blue and cyan; the position of the labeled Cys-561 is shown in red. (B) Percentage of MsbA molecules with associated nucleotide-binding domains during the hydrolysis cycle (MsbA molecules displaying the shorter distance; ~36 Å). Data are means±SD. The experimental conditions were: apo, nucleotide-free state obtained in nominally divalent cation-free buffer with 1 mM EDTA to chelate trace divalent cations and prevent ATP hydrolysis; ATP-bound, ATP-bound state obtained after the addition of 1 mm NaATP; MgATP, continuous hydrolysis state obtained after the addition of 10 mm MgSO4; and MgATP + Vi, high-energy posthydrolysis state obtained by the addition of 0.25 mm orthovanadate in the presence of MgATP. Modified from Zoghbi et al. [34].

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    Distance distributions in the apo state and during hydrolysis. Distance distributions calculated from the lifetime distributions of LRET sensitized emission intensity decays in the apo state and during ATP hydrolysis. Black, apo; red, continuous ATP hydrolysis in the presence of Mg-ATP. (A) Distance distributions in MsbA-NDs. (B) Distance distributions in MsbA-detergent.