Characterization of Small Isotropic Bicelles with Various Compositions

Article pubs.acs.org/Langmuir

Characterization of Small Isotropic Bicelles with Various Compositions K. S. Mineev,*,† K. D. Nadezhdin,†,‡ S. A. Goncharuk,†,§ and A. S. Arseniev†,‡ †

Shemyakin−Ovchinnikov Institute of Bioorganic Chemistry, Russian Academy of Sciences RAS, str. Miklukho-Maklaya 16/10, Moscow, 117997 Russian Federation ‡ Moscow Institute of Physics and Technology, Institutsky per., 9, 141700, Dolgoprudnyi, Russian Federation § Lomonosov Moscow State University, Leninskiye Gory, 1, Moscow, 119991, Russian Federation S Supporting Information *

ABSTRACT: Structural studies of membrane proteins are of great importance and interest, with solution and solid state NMR spectroscopy being very promising tools for that task. However, such investigations are hindered by a number of obstacles, and in the first place by the fact that membrane proteins need an adequate environment that models the cell membrane. One of the most widely used and prospective membrane mimetics is isotropic bicelles. While large anisotropic bicelles are wellstudied, the field of small bicelles contains a lot of “white spots”. The present work reports the radii of particles and concentration of the detergents in the monomeric state in solutions of isotropic bicelles, formed by 1,2-dihexanoyl-sn-glycero-3-phosphocholine (DHPC), 3-[(3-cholamidopropyl)dimethylammonio]-1propanesulfonate (CHAPS), 3-[(3-cholamidopropyl)dimethylammonio]-2-hydroxy-1-propanesulfonate (CHAPSO), and sodium cholate, as a function of lipid/detergent ratio and temperature. These parameters were measured using 1H NMR diffusion spectroscopy for the bicelles composed of lipids with saturated fatty chains of different length and lipids, containing unsaturated fatty acid residue. The influence of a model transmembrane protein (membrane domain of rat TrkA) on the properties of bicelles and the effect of the bicelle size and composition on the properties of the transmembrane protein were investigated with heteronuclear NMR and nuclear Overhauser effect spectroscopy. We show that isotropic bicelles that are applicable for solution NMR spectroscopy behave as predicted by the theoretical models and are likely to be bicelles rather than mixed micelles. Using the obtained data, we propose a simple approach to control the size of bicelles at low concentrations. On the basis of our results, we compared different rim-forming agents and selected CHAPS as a detergent of choice for structural studies in bicelles, if the deuteration of the detergent is not required.

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advantages and drawbacks.1 Organic solvents can sometimes maintain the secondary but very rarely the tertiary structure of the MP.2,3 Detergent micelles can retain the proper tertiary structure of some MPs4−7 and have a relatively small size to provide the NMR spectra with narrow lines; detergents are cheap, and some of them are commercially available in a perdeuterated form. On the other hand, micelles have a spherical shape and do not contain any patch of a planar lipid bilayer, which can severely affect the spatial structure of MPs, especially of those containing juxtamembrane regions that tend to interact with the membrane surface.8 Moreover, detergents usually tend to unfold the water-soluble domains.9 For that reason, a very thorough detergent screening procedure and functional assay are needed to find the exact detergent or combination of detergents and protein-to-detergent ratio that support the native structure of the MP under investigation,10,11

INTRODUCTION Nowadays, structural studies of membrane proteins (MPs) are of great importance and interest, with solution and solid state nuclear magnetic resonance (NMR) spectroscopy being very promising tools for that task. However, such investigations are hindered by a number of obstacles, and in the first place by the fact that MPs need an adequate environment that models the cell membrane. In the case of solution NMR spectroscopy, such an environment has to meet three major requirements: (1) soluble particles of the membrane mimetic should be of a relatively small size to provide a well resolved NMR spectra of the MP; (2) lipids or lipid analogues that constitute the membrane-like environment must to the highest possible extent reproduce the major properties of a lipid bilayer; and (3) components of the membrane mimetic should not disturb the spatial structure of the globular water-soluble domains, which may be present in some MPs. Five major classes of membrane mimetics are now available for solution NMR spectroscopy: mixtures of organic solvents, detergent micelles, lipid/protein nanodiscs (LPNs), amphipols, and bicelles, all having their © 2016 American Chemical Society

Received: March 4, 2016 Revised: May 5, 2016 Published: June 10, 2016 6624

DOI: 10.1021/acs.langmuir.6b00867 Langmuir 2016, 32, 6624−6637

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Langmuir and the existence of such an “ideal” set of conditions is not guaranteed. LPNs, formed by a membrane scaffold protein (MSP) and lipids, contain a large patch of lipid bilayer and represent an almost ideal model of cellular membrane.12 However, the size of LPNs is far too large for solution NMR spectroscopy (∼120−150 kDa), restricting their usage to application as a reference medium in the detergent screening and as an environment for the protein refolding.13−15 Recent advances, such as modified MSPs, which provide smaller LPNs,16 allow even the structural studies.17,18 Still, good NMR spectra are obtained only in relatively large LPNs (70−80 kDa),16,17 which is too large for solution NMR spectroscopy, and the properties of lipids in LPNs are thought to not correspond to the properties of lipids in fluid bilayers.19,20 Amphipols, short amphipatic polymers, are also a promising membrane mimetic, and were shown to be applicable for solution NMR spectroscopy;21 however, they are expensive, form relatively large particles, and have a nonlipid nature, which questions their adequacy as the model of a cell membrane. Bicelles are a compromise between the detergent micelles and LPNs. Bicelles are produced as a mixture of phospholipid and detergent, and are believed to contain the patch of a planar bilayer, surrounded by the rim of detergent.22 Self-assembly of bicelles was first reported in 1984,23,24 and since the beginning of the 1990s, they were actively used as an anisotropic environment to measure the NMR residual dipolar couplings, due to the ability of large bicelles to orient spontaneously in a strong magnetic field.25 First, magnetically aligned bicelles employed the bile salt analogue, 3-[(3-cholamidopropyl)dimethylammonio]-2-hydroxy-1-propanesulfonate (CHAPSO), as a rim-forming surfactant, but later, when the small doublechain lipid 1,2-dihexanoyl-sn-glycero-3-phosphocholine (DHPC) was also shown to support the formation of magnetically orientable particles,26 a mixture of DHPC and 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC) became a bicellar system of reference. The size of the bicelle varies in a wide range (40−... kDa) and is controlled by the ratio between long-chain and short-chain lipid or, in general, between the lipid and rim-forming agent, q. Bicelles with large q have a large size and are aligned in the strong magnetic field. Bicelles with small q are not affected by the magnetic field and can be used as a membrane mimetic in solution NMR studies. Low q bicelles are referred to as “isotropic”. Isotropic bicelles have been used in structural studies since the mid 2000s, which resulted in a number of resolved spatial structures, both of helical27−33 and β-barrel MPs,34 and even yielded some information on the physics beyond the helix−helix interactions in the membrane domains of integral MPs.35 The DMPC/DHPC system has been studied quite thoroughly. Several models describing the geometry of such bicelles were suggested and tested,22,36,37 the morphology of bicelles was studied by cryo-electron (cryo-EM) and freezefraction electron microscopy,38,39 small angle neutron scattering (SANS),40,41 NMR diffusion,42−44 and dynamic light scattering (DLS),45 which allowed the complex phase diagrams for the DMPC/DHPC and DMPC/CHAPSO systems to be plotted.38,41 However, most of the listed studies refer to the case of large and magnetically aligned species. Despite the publication of several important works in the past three years, there is still a lack of data in the field of small isotropic bicelles, which are used in solution NMR experiments. It is known that there is a patch of lipids around the protein in small bicelles;34 q = 0.5 DMPC/DHPC bicelles have a hydrodynamic radius of 3−4.5

nm (depending on the measurement approach);22,43,45 low q DMPC/DHPC bicelles grow in size upon the dilution;44,46 according to MD simulation, q = 0.25 DMPC/DHPC bicelles demonstrate a high extent of detergent/lipid mixing and absence of the planar region,46 and the size of DMPC/DHPC bicelles at some q (0.5, 0.75, and 1) and at certain total lipid concentrations is strangely dependent on the ambient temperature.45 One can also make bicelles, containing lipids with glycerol,47 ethanolamine or serine48 headgroups, cholesterol,49 ceramide,50 and gangliosides,51 to model the composition of a real membrane. This all is definitely not enough to understand and predict the properties of isotropic bicelles used in structural studies. When choosing the membrane mimetic and q ratio, we would like to know the size of the bicelle particle, the number of lipids in the planar part of the disc (whether it is enough or not to surround the protein/absorb the juxtamembrane region), and the properties of lipids around the protein, to control the size of the bicelles at any possible concentration of lipids. It would also be useful to know how exactly the q ratio may affect the structure of the protein inside the bicelle and how the presence of the protein alters the size of the bicelle particle. There is no data regarding the size and shape of low-q bicelles formed by CHAPSO or its analogue 3-[(3cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS), while our recent work shows that CHAPS needs to be considered as a possible rim-forming agent. CHAPS in the rim of the bicelles can support the native structure of the extramembrane domains of large membrane proteins, and DHPC causes the denaturation of such domains.18 Thus, the difference between CHAPS (CHAPSO) and DHPC in bicelles needs to be understood. In this respect, the objectives of the present work are (1) to compare different rim-forming surfactants in isotropic bicelles; (2) to determine the limits of applicability of various bicellar systems for solution NMR spectroscopy; (3) to study the effect of the length and saturation of fatty chains of lipids on the properties of bicelles; and (4) to investigate the properties of bicelles bearing the transmembrane proteins. For this purpose, we measure the radii of particles and concentration of the detergents in the monomeric state in solutions of isotropic bicelles formed by DHPC, CHAPS, CHAPSO, and sodium cholate as a function of lipid/detergent ratio, concentration, and temperature by 1H NMR diffusion spectroscopy. The influence of a model transmembrane protein (membrane domain of the rat TrkA receptor) on the properties of bicelles and the effect of the bicelle size and composition on the structural and dynamic properties of the transmembrane protein are also investigated.

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EXPERIMENTAL SECTION

Materials. Lipids and detergents (DMPC, 1-palmitoyl-2-oleoyl-snglycero-3-phosphocholine (POPC), 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC), DHPC, CHAPS, CHAPSO, sodium cholate) were the product of Avanti Polar Lipids Inc., USA. Stable isotope derivatives (D2O, mixture of 15N/2H-labeled amino acids) were the product of Cambridge Isotope Laboratory, USA. The structures of all used lipids and detergents are shown in Figure S1. Protein Expression. The TrkA-TM (residues 410−447, UniprotID P04629) construct was derived from the gene of the TrkA from Rattus norvegicus (NM_021589.1), kindly provided by M. Vilar by adding point mutation C442A. The TrkA-TM was expressed by a continuous exchange cell free (CF) system via the previously described protocol.18 To obtain 15N,2H-labeled TrkA-TM, the 15N/2H amino acid mixture was used. The precipitate of the reaction mixture was washed three times by the 50 mM Tris pH 8.0, 250 mM NaCl buffer, 6625

DOI: 10.1021/acs.langmuir.6b00867 Langmuir 2016, 32, 6624−6637

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Langmuir solubilized in 50 mM Tris pH 8.0, 50 mM NaCl, 1% laurylsarcosinate buffer, and finally purified by size-exclusion chromatography in 50 mM Tris pH 8.0, 50 mM NaCl, 0.2% laurylsarcosinate buffer. The protein fractions were combined, and TrkA-TM was extracted via the trichloroacetate/acetone protocol.52 Sample Preparation. To prepare the protein-free bicelles, ∼3 mg of lipids in dry powder was dissolved by the necessary amount of 10% stock solution of CHAPS, CHAPSO, or DHPC and then the total volume of the sample was adjusted to 200 ul by the addition of 30 mM NaPi buffer, pH 6.0. The resulting mixture was frozen and then heated to 40 °C in an ultrasonic bath for three to four times. A new sample was prepared for each studied q ratio. To assemble the bicelles, containing the TrkA-TM, protein precipitate was dissolved in trifluoroethanol/H2O 2:1 mixture with necessary amounts of DMPC and detergents (the overall lipid/protein ratio was taken to be equal to 120, and the initial lipid/detergent ratio q was taken to be equal to 0.5 for DMPC/DHPC and 1.0 for DMPC/CHAPS). Then, the resulting solution was diluted by pure water until the trifluoroethanol/H2O ratio reached 1:1, frozen in liquid nitrogen, and dried on the lyophilic drier. The obtained powder was dissolved in 500 ul of 30 mM NaPi buffer, pH 6.0, and put into the ultrasonic bath for 5−6 min at 40 °C. To vary the q ratio, the solution was added to the necessary amount of lipid/ detergent in dry powder and put into the ultrasonic bath for 5−6 min at 40 °C. For the dilution experiments, a 200 mM solution of bicelles at necessary q was prepared. To vary the concentration of bicelles, necessary portions of the initial solution were diluted by either pure water or the 3.5/5.5 mM solution of CHAPS and 8 mM solution of DHPC, depending on q and the type of studied bicelles. To prepare the LPN samples, necessary amounts of MSP1D1 or MSP1ΔH516 were added to the mixture of DMPC and sodium cholate (1:1) dissolved in the ND-buffer (20 mM Tris pH 8.0, 100 mM NaCl) and incubated for 1 h. The total lipid/MSP ratio was adjusted to 65 in the case of MSP1D1 and 35 in the case of MSP1ΔH5. To assemble LPNs, the Biobeads SM-2 (Biorad, USA) (1 g per 70 mg of detergent) were added and the suspension was shaken at room temperature for 12−16 h. The resulting solution was filtered (to remove Biobeads) and centrifuged at 14000g for 20 min, and the supernatant was concentrated to 500 ul. The concentration of lipids was assessed by 1 H NMR in chloroform/methanol 1:1 mixture, and the concentrations of MSPs were measured by UV spectroscopy. NMR Experiments. All diffusion 1H NMR spectra were recorded on a Bruker Avance 700 spectrometer with working frequency on protons equal to 700 MHz, equipped with a room-temperature triple resonance TXI probe. Diffusion was measured using the double LED pulse sequence53 with constant delay and varied gradient pulse power with the presaturation of water resonance in thick-wall NMR tubes (to avoid the convection effects). Diffusion experiments were recorded with 8 scans and 32 increments of the encoding gradient pulse (gradient pulse strength was varied linearly from 1 to 52 G/cm). The summary length of the encoding gradient pulse was 3.0 ms. The gradients were calibrated using the diffusion coefficient of residual water in 100% D2O at 25 °C. Experiments were run at 27 and 40 °C; for some samples, the temperature dependence in the range 10−50 °C was recorded. To take into account the possible nonequilibrium effects and convection, several (4−5) repetitions of the LED experiment were performed with varied diffusion delay (Δ). Typical durations of the diffusion delay were 100−300 ms. In case the diffusion coefficient was found to be dependent on the Δ magnitude, the convection was supposed to occur in the sample, and the real diffusion coefficient was calculated, assuming the linear dependence of the apparent diffusion coefficient on Δ. At least a 30 min interval was used after the change of the temperature to ensure the equilibrium state of the system. In the case of CHAPS/CHAPSO/cholate-based bicelles, the diffusion coefficients of lipids were measured using signals at 0.85, 1.25, 3.25, and 3.68 ppm. Diffusion coefficients of CHAPS/CHAPSO were measured using signals at 0.68, 1.04, 2.98, 3.13, 3.37, and 3.49 ppm. Diffusion coefficients of sodium cholate were measured on the basis of the signals at 0.76, 0.96, 1.07, and 3.52 ppm. In DMPC/DHPC mixtures, signals at 0.85 and 1.25 ppm were used to determine the lipid diffusion rate, while the signals at 0.95 and 1.34 ppm were taken

to calculate the diffusion coefficient of DHPC. Diffusion coefficients, obtained for the different signals in three to four repetitions, were averaged, and their root-mean-square deviations were taken as errors of the determined magnitudes. 1H chemical shifts were determined, using trimethylsilyl propionate (0.0 ppm) as an internal reference standard. The concentration of lipids in the mixtures was additionally controlled by 1H NMR after dissolving the 10 ul aliquot of the sample in 1:1 chloroform/methanol mixture, containing 1 mM trimethylsilyl propionate (nine protons at 0.0 ppm). All protein NMR spectra were recorded on a Bruker Avance III spectrometer with working frequency on protons equal to 800 MHz, equipped with a cryogenic triple resonance TCI probe. For each sample, two temperatures, 27 and 40 °C, were investigated. Transverse relaxation optimized spectroscopy (TROSY)54 and a TROSY-based experiment for the measurement of cross-correlated relaxation rate55 were recorded to estimate the chemical shift differences and correlation time of rotational diffusion, τa. To study the protein− lipid interaction, TROSY-based 3D 15N-NOESY-HSQC was recorded with a 100 ms mixing time. Protein signals were assigned on the basis of the HNCA spectra and NH−NH contacts in NOESY. Protein− lipid NOE contacts were integrated and normalized by the intensities of diagonal peaks to take into account the alteration in the transverse relaxation of different amide groups. Bicelle Models. To consider the behavior of bicelles, one needs to first define the “ideal” bicelle and then study the deviations of the real system. In general, one could state that, for the ideal bicelle, the radius of the particle is determined by the ratio q*22

q* =

V S [Lip] = disc λ = disc k Vrim Srim [Det] − [Det]free

(1)

where [Lip] and [Det] are the concentrations of lipid and rim-forming detergent and [Det]free is the concentration of the free detergent, analogously to the critical micelle concentration for micellar solutions. Vdisc and Sdisc are the volume and surface area of the lipid bilayer disc, while Vrim and Srim are the volume and surface area of the detergent rim; λ is the ratio of volumes, detergent over lipid, and k is the ratio of surface areas per headgroup, detergent over lipid. Here and below the effective lipid/detergent ratio, as determined by eq 1, will be referred to as q*, while the nominal ratio will be referred to as q. There are several models suggested for DMPC/DHPC particles that rely on the assumption that DMPC forms a disk, surrounded by the rim of DHPC, shape and parameters of the DHPC rim are different for different models.22,36,37 The most adequate and predictive model was suggested by Triba et al.,36 and states that DHPC forms the rim with the shape of an elliptic semitoroid with thickness r⊥. In that case, the radius of the bicelle can be expressed from eq 1 as

⎡ ⎛ 2 32λ ⎞1/2 ⎤ r⊥q* ⎢ R = r⊥ + π + ⎜π + ⎟ ⎥ 4λ ⎢⎣ 3q* ⎠ ⎥⎦ ⎝

(2)

Here r⊥ is taken to be equal to 1.1 nm (length of a DHPC molecule), R is the radius of the bicelle, and λ is the volume ratio of DHPC over DMPC (0.6136). In a similar manner, we can construct the model for an ideal bicelle, composed of the lipid and a bile salt analogue (CHAPS or CHAPSO). Bile salt is a flat molecule with hydrophobic and hydrophilic faces; therefore, it is likely to form the rim, shaped as a cylindrical layer. The simplest equation can be written for the DMPC/ CHAPS(O) bicelle, if we assume that the volumes of both DMPC in the bilayer and CHAPS(O) in the rim are independent of q*:

⎡ q* R = r⊥⎢1 + + ⎢⎣ λ

q*(q* + λ) ⎤ ⎥ ⎥⎦ λ

(3)

Equation 3 contains only two unknownsthe thickness of the CHAPS(O) layer, r⊥, and the ratio of the CHAPS volume over the DMPC volume, λ. Equations 2 and 3 can be modified to take into account the presence of an incorporated protein. This is done by solving the modified eq 1 with correct expression for the shape of the rim for the specified bicelle (elliptic semitoroid or cylindrical layer): 6626

DOI: 10.1021/acs.langmuir.6b00867 Langmuir 2016, 32, 6624−6637

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Langmuir

Figure 1. q*-Dependent parameters of various bicelles plotted as a function of the q* ratio at 27 (squares) and 40 °C (circles). (A) Radii of DMPC/ DHPC bicelles. “Ideal” R(q*) dependence is plotted by a solid line (eq 2 with r⊥ = 1.15). (B) Radii of DMPC/CHAPS bicelles. “Ideal” R(q*) dependence is plotted by a solid line (eq 3). Data measured at 15 °C are shown by triangles. (C) Radii of DMPC/CHAPSO (blue) and DMPC/ cholate (red) bicelles. Radii of DMPC/CHAPS bicelles at 27 °C are shown by black squares for reference. (D) Radii of POPC/CHAPS (red) and DPPC/CHAPS (blue) bicelles. Radii of DMPC/CHAPS bicelles at 27 °C are shown by black squares for reference. “Ideal” R(q*) dependence for the DPPC/CHAPS system is plotted by a solid line (eq 3). (F, E) Concentrations of free detergent in DMPC/DHPC (F) and DMPC/CHAPS (E) bicelles, formed at different q*.

q* =

Vdisc − Sproth [Lip] = λ [Det] − [Det]free Vrim

where Dlip is a measured coefficient, D0lip is a single-particle diffusion coefficient at infinite dilution, and λ = 1 for hard-sphere noninteracting particles. D0lip can be used to calculate the hydrodynamic radius of the particle RH:

(4)

Sprot is the area, taken by the protein on the surface of the bicelle, and h is the bilayer thickness. Interpretation of the Measured Diffusion Coefficients. In the present work, we relied on the NMR-measured translational diffusion coefficients to describe the size of isotropic bicelles in solution. This approach has a major advantage in comparison to other available techniques, such as DLS, SANS, and small-angle X-ray scattering (SAXS). In addition to the hydrodynamic radii of particles under investigation, NMR diffusion measurement allows the direct identification of the [Det]free term in eq 1, and therefore, both q* and R are determined independently in a single experiment from the diffusion coefficients of lipid (Dlip) and rim-forming detergent (Ddet)

Ddet = αDf + (1 − α)D lip

0 RH = kT /6πηDlip

Equation 8 is correct for spherical particles, to find the radius of a bicelle, which may be considered as a cylinder with height h and radius R, one needs to use the shape factor58

f /f0 = 1.009 + 1.395 × 10−2(ln p) + 7.880 × 10−2(ln p)2 + 6.040 × 10−3(ln p)3

2 1/3

and f 0 = 6πηh(3/16p ) . where p is the aspect ratio, h/2R, f = Equation 9 can be solved numerically for any given diffusion coefficient, assuming that h = 4.0 nm for DMPC or POPC and 4.4 nm for DPPC bicelles.59 In addition to the protein-free bicelles, we have studied bicelles with an incorporated model transmembrane α-helix. Since it is impossible to achieve a uniform distribution with one protein residing in every bicelle, we relied on the rotational diffusion measurements to assess the size of protein-bearing bicelles. Rotational diffusion of a bicelle is anisotropic, and three correlation times are normally needed to describe it. However, we use N−H vectors as reporters of the protein mobility, and in the single α-helix, all N−H vectors are directed along the helix axis, which, in turn, is directed along the main axis of the bicelle inertia tensor. Therefore, in an experiment that we use here (rate of cross-correlated 1H,15N relaxation55), only one correlation time is contributing to the measured effect, which is τa = 1/6D⊥. The radius of the bicelle can be calculated from the τa magnitude via an equation similar to eq 9

(5)

(6)

where Df is the diffusion coefficient of the free detergent at a given bicelle concentration, D0f is the diffusion coefficient of pure detergent below the CMC, and Φ is the volume fraction of bicelles in solution. In our calculations, we defined Φ as the ratio of the mass of lipids and detergents (subtracting the total mass of free detergent in solution) over the total mass of the solution. In a similar manner, measured diffusion coefficients of lipids need to be corrected to obtain the real diffusion coefficient at infinite dilution,42,57 which can be converted to the hydrodynamic radius of bicelles 0 Dlip = Dlip (1 − 3.2λΦ)

(9) kt/D0lip,

where α is a fraction of the detergent in a water-soluble form, [Det]free = α[Det], and Df is a diffusion coefficient of monomeric detergent. Df can be measured in a separate experiment, on the sample, containing the pure detergent below the CMC and corrected to take into account the obstruction of the solvent diffusion, caused by the presence of bicelles56 Df = (Df0)/(1 + Φ /2)

(8)

τa/τ0 = 1.18 + 0.1744(ln p + 0.2877)2 − 0.2417 (ln p + 0.2877)3 − 3.882 × 10−2(ln p + 0.2877)4 , p < 0.75

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Langmuir where p is the aspect ratio, h/2R, τa is a measured parameter, and τ0 is a correlation time for a sphere with the same volume: πh3η/4p2kT.58 Analogously to translational diffusion, rotational diffusion is also obstructed at higher concentrations;60 therefore, measured τa needs to be corrected prior to solving eq 10:

τa0 = τa(1 − 0.63Φ − 0.74Φ2)

by the presence of rod-like or band-like particles in the solutions of high-q* isotropic bicelles above the phase transition temperature of DMPC, as was found by cryo-EM spectroscopy.41,45 Apart from the bicelle radius, the concentration of free DHPC was also found to be dependent on both temperature and q* (Figures 1F and 2C). The q* dependence is more pronounced: [Det]free gradually decreased from 9.7 mM at q* = 0.31 to 7.3 mM at q* = 0.82 at 40 °C. Reported [Det]free, measured at both 27 and 40 °C, is always slightly higher than obtained with 31P NMR.46 However, we need to point out that in the case of the 31P studies only one parameter is measured, and titration is performed, assuming that [Det]free is constant. We show here that [Det]free is dependent on q*; therefore, the parameter decreases upon dilution of bicelles, which results in the small systematic error in [Det]free, measured in dilution experiments. To summarize the paragraph, we conclude that at q* = 0.3− 0.8 DMPC/DHPC bicelles follow the “ideal” model at temperatures, normally used in solution NMR spectroscopy; however, some temperature-dependent nonideal behavior is observed. DMPC/CHAPS Bicelles. A different pattern is observed for DMPC/CHAPS and DMPC/CHAPSO bicelles. First of all, both CHAPS and CHAPSO form bicelles, which are much smaller than DMPC/DHPC particles with the same q* (Figure 1B, Table S2). For example, DMPC/CHAPS form bicelles with a radius equal to 3.0−3.2 nm when q* is in the range 0.87−1.0, and this is equivalent to q* ∼ 0.5−0.6 for DMPC/DHPC solutions. According to model (3), R(q*) dependence has to be linear or almost linear. Analysis of the obtained curves reveals that DMPC/CHAPS bicelles follow the model in the range 0.6 < q* < 1.1 at 27 and 40 °C. Parameters of the fit to eq 3 are as follows: r⊥ = 1.10 ± 0.06 nm and λ = 1.34 ± 0.15, which seem rather reasonable, taking into account that the volume of the CHAPS molecule has to include the portion of the solvent shell around the particle. Both at q* < 0.6 and at q* > 1.1 bicelles are formed with larger radii than expected from specified r⊥ and λ magnitudes. The concentration of free detergent is lower for CHAPS than for DHPC and is also dependent on q*: at q* = 0.4, there was 6.3 mM of free CHAPS and its quantity decreased gradually to 2.3 mM at q* = 1.5 (Figure 1E). [Det]free is also almost not dependent on the temperature: it decreases from 4 to 2.3 mM at 10−20 °C and then gradually increases to 2.8 mM at 50 °C at q* = 1.55. At q* = 1.05, the character of the dependence is similar (Figure 2D). Analysis of 1 H NMR spectra reveals that properties of lipids are very much alike in DMPC/DHPC and DMPC/CHAPS bicelles with q* > 0.6 (Figure S2). In both cases, the signal of the lipid methyl group was very narrow and split into two triplets, indicating the different environment of two fatty chains. The splitting was dependent on q* and is maximal for larger bicelles. Unlike DMPC/DHPC, in DMPC/CHAPS bicelles with q < 0.6, signals of two chains merge, implying that the packing of the bilayer (if one is present) becomes different from that observed in DMPC/DHPC particles. This may be caused by the mismatch between the equilibrium thickness of the lipid bilayer and the geometric properties of the CHAPS rigid core; and it could explain the deviation of the q*(R) dependence from linear at low q*, observed in DMPC/CHAPS solutions. Almost no effect of the ambient temperature is observed for DMPC/CHAPS mixtures at q* < 1.1; the bicelle radius is constant at q* = 1.05 in the range 10−50 °C (Figure 2B). In

(11)

Altogether, eqs 5−11 provide a self-consistent set of bicelle radii and free detergent concentrations, which, as we will show below are in agreement with the data, obtained by other techniques, such as phosphorus NMR, light scattering, and SAXS/SANS experiments.

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RESULTS AND DISCUSSION DMPC/DHPC Bicelles. To begin, we have measured the radius of q = 0.5 DMPC/DHPC bicelles, which were studied by a variety of techniques, to establish the reliability of our approach. Use of eqs 5−9 resulted in the following parameters: diffusion coefficient at infinite dilution, Do = (111 ± 1) × 10−12 m2 s−1, bicelle radius, R = 3.1 ± 0.1 nm, [Det]free = 8.3 ± 0.3 mM at q* = 0.53 and 40 °C (Table S1). According to DLS measurements, q = 0.5 bicelles have radii in the range 2.9−3.3 nm, depending on concentration45 and phosphorus NMR yields [Det]free in the range 7−8 mM for this particular q ratio.22,46 However, it is noteworthy that the results of other NMR diffusion studies are very controversial. Recent works report the following radii of q = 0.5 DMPC/DHPC species: RH = 4.3 nm at 25 °C43 and 6−8 nm, depending on temperature.61 The analysis of the listed works reveals that in one case (6−8 nm) the measured diffusion was not corrected for the concentration of bicelles and in the other case (4.3 nm) the correction was applied but authors did not take into account that bicelles grow in size upon dilution, due to the fact that the presence of free detergent in solution increases the effective q* ratio, and the effect is much more pronounced at low concentrations (authors diluted bicelles to ∼1%). This could result in the incorrect fit and reduced coefficient λ in eq 7, which was not reported in the paper. If we take the reported diffusion coefficient D = 56 × 10−12 m2 s−1 at 25 °C and 200 mM of total lipids43 and apply eq 7 with λ = 1, we will obtain R = 2.96 nm, which is consistent with DLS experiments and the radius measured here. Therefore, we conclude that our approach yields data that are in agreement with DLS measurements and phosphorus NMR, while it is quite difficult to reconcile our results with other NMR diffusion studies. To continue, we recorded the R(q*) dependence for several types of isotropic bicelles at two temperatures, 27 and 40 °C, starting from classic DMPC/DHPC mixtures (Table S1). At 40 °C, the radius of these bicelles ranged from 2.5 nm at q* = 0.31 to 4.3 nm at q* = 0.82 and almost perfectly followed the “ideal” model (eq 2) with r⊥ between 1.1 and 1.2 nm (Figure 1A). It is noteworthy that the bicelle radius was depending on the ambient temperature. At 27 °C, bicelles were slightly (0.1−0.2) greater for q* < 0.6, while q* = 0.8 bicelles were 0.3 nm greater at 40 °C. According to the infrared spectroscopy measurements,46 the phase transition takes place below 27 °C in isotropic bicelles. Therefore, the effect cannot be explained by the increase of the average volume of lipid molecules in the planar region of a bicelle caused by the phase transition of lipids. Most likely, the size of the particles is increased at high q* due to the temperature-dependent mixing of lipids and detergents in the lipid bilayer region of bicelles, which was shown to occur in magnetically aligned species;36 or, alternatively, the observed temperature dependence is caused 6628

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the case of larger q*, the size of bicelles begins to grow with the temperature and the magnitude of the effect is greater in larger bicelles. At q* = 1.55, the radius of the particles is increased from 4.2 nm at 15 °C to 9.5 nm at 50 °C. Like in the case of DMPC/DHPC species, these changes also cannot be in a simple way explained by the gel−liquid crystalline phase transition. According to the literature, the volume of lipid molecules is increased by 10−20% upon such a transition; therefore the size of a bicelle is not expected to grow greater than by 20%.62 It is noteworthy that below the critical temperature of DMPC (24 °C) large DMPC/CHAPS bicelles are formed with the radii, predicted by model (3) with parameters, reported above. Therefore, we state that below the phase transition temperature DMPC/CHAPS bicelles behave “ideally”, while at higher temperatures large bicelles (q* > 1.1) begin to grow in size, which is, most likely, caused by the temperature-dependent mixing of lipids and detergents in the lipid bilayer region of bicelles or by the change in the bicelle shape and presence of the band-like structures in solution, as reported in previous works.41,45 In other words, DMPC/ CHAPS bicelles are applicable as membrane mimetics for solution NMR spectroscopy at q* < 1.1. CHAPSO and Sodium Cholate as an Alternative to CHAPS. To go ahead, we tested the sodium cholate and CHAPSO as rim-forming agents for isotropic bicelles. CHAPSO behaved almost identically to CHAPSthe R(q*) dependence was similar and large bicelles also grew in size with the temperature (Figure 1C, Table S3). In 1H NMR spectra, the shape and width of signals from DMPC and detergents were completely the same, indicating the similar parameters of lipid packing and mobility. This is expected, taking into account that CHAPS and CHAPSO have almost identical structure and similar CMC values (6 and 8 mM, respectively). Nevertheless, at q* ∼ 1, CHAPSO formed bicelles that are 5−7% greater in comparison to the DMPC/CHAPS particles; however, this difference does not exceed the two sums of errors of measured magnitudes, and can be deemed negligible. We were not able to find in the literature the reason why CHAPSO was initially preferred to CHAPS in the case of magnetically aligned bicelles,25 but it is clear that CHAPS and CHAPSO are almost equivalent for the solution NMR applications. In contrast, mixtures of DMPC and sodium cholate did not behave as expected for an ideal bicelle. At q* < 1, radii of DMPC/cholate bicelles were up to 20% greater than was observed for DMPC/ CHAPS mixtures with the same q*. Moreover, at q* > 1.25, the size of bicelles stopped growing with the increase of q* (Table S4). Third, signals from the detergents and lipids in 1H NMR spectra were relatively broad at all tested q*, indicating that the environment of both lipid and cholate molecules is heterogeneous (Figure S2). Altogether, these observations suggest that DMPC and sodium cholate form mixed micelles rather than bicelles: the detergent may be present in the planar area (if such an area exists), and lipid could migrate to the rim. Use of Other Lipids for Bicelle Formation. Previous studies already demonstrated that lipids with longer and shorter fatty chains (DPPC and 1,2-dilauroyl-sn-glycero-3-phosphocholine61), unsaturated chains (POPC42), and anionic headgroups (DMPG47) can form bicelles with DHPC, while no R(q*) dependencies were measured for those cases. To further investigate the bicellar systems, we tested the possibility of bicelle formation of lipids with longer fatty tails and unsaturated chains with CHAPS (Tables S5 and S6). Most commonly used lipids, POPC and DPPC, were selected for that purpose.

Figure 2. (A) Temperature dependence of the disk radius R, measured for DMPC/DHPC bicelles with q* = 0.63 (red) and 0.82 (blue). (B) Temperature dependence of the disk radius R, measured for DMPC/ CHAPS bicelles with q* = 1.55 (red circles), 1.40 (green triangles), 1.29 (magenta triangles), and 1.05 (blue squares). The solid blue line in panel B corresponds to R = 3.2 nm. (C) Temperature dependence of free detergent concentration in DMPC/DHPC solutions at q* = 0.63 (red) and 0.82 (blue). (D) Temperature dependence of free detergent concentration in DMPC/CHAPS solutions at q* = 1.55 (red) and 1.05 (blue). 6629

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Following the same methodology, the radius of the MSPΔH5 LPNs was found to be equal to 4.1 ± 0.1 nm, which is comparable to q* = 1.2 DMPC/CHAPS bicelles (3.9 nm) or q* = 0.8 DMPC/DHPC bicelles (4.2 nm) at 40 °C and corresponds to 85 molecules of DMPC in the bilayer. This value surprisingly coincides exactly with the reported size of MSPΔH5 LPNs measured by cryo-EM and DLS (4.1 and 4.2 nm16). As two bicelle samples with similar size were obtained and described, we can compare spectroscopic properties of lipid signals in bicelles and LPNs. Major distinction can be found, looking at the appearance of the methyl groups of lipid fatty tails. In bicelle solutions, signals from methyl groups are extremely narrow and split due to the different environment of two fatty chains of DMPC, as discussed above (Figures 3 and

Together with DMPC, these two sample all possible cases with phase transition temperature: POPC will always be above the phase transition (−2 °C), while DPPC will be below or close to the phase transition (41 °C) under experimental conditions. First experiments revealed that POPC/CHAPS bicelles are extremely large at q* = 1 (R = 5.4 and 5.7 nm at 27 and 40 °C), while at q* = 0.6 their size was comparable to the size of DMPC/CHAPS particles (2.9 nm, Figure 1D). Probably, the temperature-dependent increase in the size of a bicelle is relevant at lower q* for POPC mixtures. Since the lipid is always far above the phase transition, the effect of temperature is almost not observed at 27−40 °C, while the size is already increased. DPPC in its turn always formed the ideal bicelles: the R(q*) dependence was linear at 27 °C on the whole range of tested q* (0.4−1.3); some deviations from the ideal behavior were observed for the q* = 1.3 DPPC/CHAPS mixture at 40 °C. Parameters of model (3) that describe the behavior of DPPC/CHAPS bicelles are the following: r⊥ = 1.31 ± 0.05 nm and λ = 1.61 ± 0.15. In other words, CHAPS forms a 0.2 nm thicker rim around DPPC than around DMPC, and the volume of a single CHAPS molecule in the rim is increased by 20%. Despite the “ideal” behavior of DPPC/CHAPS bicelles in a wide range of q*, they are scarcely applicable for solution NMR experiments: DPPC/CHAPS start to form bicelles only after several freeze−thaw cycles with heating to 40−50 °C in the ultrasonic bath, so very accurate temperature control is needed and problems with incorporation of proteins into the DPPCbased bicelles are expected. Altogether, data obtained for DMPC, DPPC, and POPC in bicelles reveal that CHAPS-based bicelles behave as predicted by model (3) at low q* and at some point (q* = 1.1−1.2 for saturated fatty chains) the size of the bicelle begins to depend on the temperature. Below the temperature of phase transition, large bicelles also behave “ideally”, while, above the phase transition temperature, their size is much greater than can be expected from the volume and surface area per headgroup of lipids in fluid bilayers. Most likely, the state of lipids in small bicelles is always the same as the state of lipids in large bicelles below the phase transition temperature. Lipid−Protein Nanodiscs. Since lipid−protein nanodiscs (LPNs) are starting to be widely used as membrane mimetics for NMR structural studies in solution,16−18 we decided to characterize LPNs using the NMR diffusion experiments with the same probe calibration and data processing approach given in the theory section. For that purpose, we selected the LPNs formed by two variants of membrane scaffold protein (MSP): MSPD1, which is the full-length construct, providing the largest LPN particles, and MSPΔH5, which is usually selected from the recently introduced shorter variants of MSP17 to perform structural studies. Our data show that DMPC/MSPD1 LPNs have a radius of 4.9 ± 0.1 nm, which corresponds to 150 molecules of lipid in the disc, assuming the average surface area per lipid headgroup is equal to 0.65 nm2.59 The size of LPNs is not dependent on the temperature in the range 27−40 °C within the experimental error, and corresponds to an ideal DMPC/CHAPS bicelle with q* = 2.1 and DMPC/DHPC bicelle with q* = 1.23. The obtained radius is consistent with the LPN parameters, measured by other techniques, such as SAXS (5.1−5.2 nm63), SANS (4.7 nm64), cryo-EM (4.75 nm16), and DLS (4.7 nm not implementing the shape-factor16). This once again confirms the applicability and reliability of our approach.

Figure 3. Comparison of signals from the lipid methyl groups, observed in 1H NMR spectra of MSPD1 and MSPΔH5 LPNs, with line shapes observed in DMPC/DHPC and DMPC/CHAPS bicelles of similar size.

S2). In LPNs, the signal of the methyl group is broad, indicating that either the motions of lipid chains are slowed down or the environment of lipid molecules is heterogenic. At high temperatures (50 °C), the triplet shape of the methyl groups becomes resolved, and comparison to the spectra of large bicelles demonstrates clearly that two chains of DMPC are equivalent in MSPΔH5 LPNs, like they are in low-q* DMPC/ CHAPS particles. Thus, a different state of lipids is observed in LPNs in comparison to various bicelles. This is in agreement with the observation that in LPNs lipids are packed tightly and are less mobile than in fluid liposomes.19,20,64 On the other hand, it was shown that LPNs experience phase transitions at temperatures close to the critical temperatures found for lipid bilayers.63 Therefore, it is quite difficult to understand what the biologically relevant analogy of the lipid bilayer in bicelles and LPNs iswhether it is a classic gel, liquid ordered, or liquid crystalline states or we observe some different states with altered mobility, volume, surface area per headgroup, and other parameters of lipids. Controlling the Bicelle Size upon Dilution. A recent work reports that DMPC/DHPC bicelles grow in size upon dilution.46 This effect was attributed to the presence of a certain amount of free DHPC in solution, which does not depend on the concentration of bicelles and increases the effective q* ratio. With this regard, authors introduced the concentration boundaries for the application of bicelles, so that the q* does not deviate from the initially planned q by more than 10%. For instance, they stated that 120 mM is the minimal concentration for q = 0.5 DMPC/DHPC bicelles. Despite the amount of 6630

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Langmuir other significant data reported in the work,46 we deem this particular suggestion as impractical for two major reasons. First, rotational diffusion of bicelles is impeded at high concentrations (see eq 1160), which leads to line broadening and a decrease in sensitivity in NMR spectra. According to an experimental NMR study,65 the effect of the increased lipid concentration can be much more severe than predicted by eq 11, when the concentration of lipids exceeds 10% (corresponding to ca. 200 mM DMPC/DHPC bicelles). Therefore, one would like to minimize the total amount of lipids in solution as much as possible. Second, sometimes it is necessary to maintain the certain lipid-to-protein ratio (LPR) (e.g., when studying the dimerization of membrane proteins). The amount of protein is usually limited, and one cannot increase the concentration of lipids to the desired 120 mM. The normal NMR concentration of a protein for the screening of experimental conditions is 0.1 mM, and in some cases, one would like to have as little as 4−6 mM of lipids in bicelles to maintain the LPR 40−60. Thus, instead of using the lower limit for the lipid concentration, it would be much more practical to control the size of bicelles at any required amount of lipid in solution. It can be easily accomplished if the concentration of free detergent is known for the given q*. To show that, we performed the series of dilution experiments with q* = 0.5 DMPC/DHPC, q* = 0.5 DMPC/CHAPS, and q* = 1.0 DMPC/CHAPS bicelles (Figure 4). In all three cases, initial samples containing 200 mM of lipids were diluted either by the pure water or by the solution, containing 8 mM DHPC, 3.3 mM of CHAPS (for q* = 1), or 6 mM of CHAPS (for q* = 0.5), which correspond to the concentration of free detergent for respective q*. As expected, the size of bicelles is retained even at very low concentrations, when the excess of detergent is added to the solution (DMPC/ CHAPS q* = 1 bicelles were diluted to 1.3 mM), while bicelles grow in size dramatically, when diluted by pure water. Moreover, all three data sets fit almost perfectly to eq 7, with the parameter λ equal to 0.98−1.00, while the best and most accurate fit, obtained for q* = 1 DMPC/CHAPS bicelles, resulted in λ = 1.00 ± 0.02. Therefore, our approach, assuming λ = 1 to determine the single-particle diffusion coefficients, is rather precise and will not result in systematic errors that exceed 2%. Analysis of 1H NMR spectra reveals that both the line width and line shape of the signal, corresponding to the DMPC methyl group, are retained upon the dilution with [Det]free solution, while they change dramatically when the initial sample is diluted by pure water (Figure S3). Still, we need to admit that at low concentration the size of bicelles is controlled worse, because when [Det] − [Det]free ≪ [Det], small errors that can take place during weighting of the detergent may lead to large errors in the obtained q*. Nevertheless, it is rather easy to adjust the size of the bicelles to the desired value in the case of such error via a titration procedure. To sum up, the recipe of the size control is simple: calculate the amounts of lipid and detergent for the desired q and then add an extra amount of detergent, corresponding to the [Det]free, reported in the present work for the specified q*, detergent, and ambient temperature (see Tables S1−S6 in the Supporting Information). Effect of the Protein on the Bicelle Radius. Apart from the size of protein-free bicelles, parameters of bicelles with incorporated proteins are also of great importance for structural studies of membrane proteins in solution. According to our model (eq 4), bicelles are expected to grow in size with embedded protein, presence of a transmembrane protein is

Figure 4. (A) Diffusion coefficients of q* = 0.53 DMPC/DHPC bicelles at 40 °C plotted as a function of the volume fraction of lipids and detergents in bicelles upon dilution by 8 mM DHPC solution (blue) or by pure water (red). A fit of the obtained data to eq 7 is shown by the blue line. (B) Dependence of the effective q* ratio on the total lipid and detergent concentration in DMPC/DHPC q = 0.53 upon dilution either by pure water (red) or by the solution, containing 8 mM DHPC (blue). The blue line denotes the desired q* = 0.53. (C) 6631

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with TrkA-TM, while at q* > 0.5 the size of the bicelles with protein is always lower than the size of protein-free bicelles with the same q*, which is unexpected (Figure 5A). This apparent decrease of bicelle size, when the TM protein is present, may be interpreted in two ways. First, interpretation is straightforward and suggests that the size of bicelles indeed does decrease; however, we cannot find any physical explanation for this possibility. On the other hand, we use cross-correlated relaxation rates to measure the coefficients of rotational diffusion, and the hydrodynamic radii of particles in solution. This approach assumes that the TM protein is rigid; if the protein is mobile, the effective hydrodynamic radius can be reduced. Therefore, the increased mobility of the model TM helix in DMPC/DHPC bicelles in the ns time scale can explain the observed effects. This assumption is additionally supported by the fact that, according to translational diffusion, the average radius of q* = 0.5 bicelles is 0.3 nm greater than the size, derived from the NMR relaxation parameters. Thus, we can conclude that DMPC/CHAPS bicelles behave as predicted, and grow in size with incorporated model TM α-helix. We can say nothing on the size and shape of protein-bearing DMPC/ DHPC particles; however, unlike DMPC/CHAPS, the model TM helix is not static in such an environment and is mobile in the ns time scale. Effect of the Bicelle Size and Type of a Rim-Forming Agent on the Properties of a TM α-Helix. As it is clear from the previous paragraph, not only the embedded protein can affect the size of bicelles, but bicelle type can also affect the properties of a TM protein. To further investigate such an

Figure 4. continued The same as panel A for q* = 0.5 (green) and q* = 0.99 (blue, red) DMPC/CHAPS bicelles diluted by 6 mM (green) and 3.3 mM (blue) CHAPS solution or by pure water (red). (D) Dependence of the effective q* ratio on the lipid and detergent concentration in DMPC/ CHAPS bicelles upon dilution either by pure water (red) or by the solution, containing 3.3 mM CHAPS (blue). The blue line denotes the desired q* = 0.99.

equivalent to the addition of extra amount of lipid to the solution. To study the effect of proteins on bicelles, we used the model transmembrane (TM) helix, the TM domain of rat TrkA (TrkA-TM, residues 410−447). The 2H,15N-labeled TrkA-TM was incorporated into DMPC/DHPC and DMPC/CHAPS bicelles, and the size of bicelle particles with protein was assessed on the basis of the cross-correlated relaxation rates (eqs 10 and 11). Experiments with DMPC/CHAPS mixtures reveal that, when q* was in the range 0.7−1.0, bicelles with TrkA-Tm behaved as predicted by the “ideal” model with Sprot = 1.5 nm2, their radius was increased by ca. 0.3 nm. Analogously to the protein-free bicelles, at low q*, the size of bicelles is slightly greater than expected and is increased by more than 0.6 nm at q* = 0.45. At q* larger than 1.1, a temperature-dependent size increase is observed. At q* = 1.31, the size of the TrkA-TM-containing bicelles exceeded 5.5 nm (Figure 5B). In the case of DMPC/DHPC bicelles, similar radii were observed at q* = 0.3−0.4 for protein-free bicelles and bicelles

Figure 5. Radii of DMPC/DHPC (A) and DMPC/CHAPS (B) bicelles plotted as a function of q* in the presence (red) and in the absence (blue) of TrkA-TM at 27 (squares) and 40 °C (circles). Blue lines represent model 2 with r⊥ = 1.15 nm (A) and model 4 with r⊥ = 1.10 nm and λ = 1.34 (B). Red dashed lines represent predicted radii of bicelles in the case when the protein is present, which takes the area on the surface of the particle (Sprot) equal to 1.5 nm2 (1 helix) and 9 nm2 (6−7 helices). (C) The line broadening of signals in 1H,15N-TROSY spectra of deuterated TrkA-TM in DMPC/CHAPS q* = 0.73 (blue bars) and DMPC/DHPC q* = 0.58 (red bars) bicelles at 40 °C. The ratio between the intensity of the signal from the C-terminal residue (I39) and the intensity of other signals is shown. The gray rectangle denotes the TM region, and the red dashed line indicates the average ratio for the C-terminal part of the TM helix. 6632

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Figure 6. (A) Differences in amide chemical shift ΔCS = ΔδH2 + ΔδN 2/36 , observed in DMPC/DHPC (q* = 0.58) and DMPC/CHAPS (q* = 0.73) bicelles, plotted versus the amino acid sequence of TrkA-TM. (B) Root-mean-square deviations of amide chemical shifts of TrkATMσ = σ 2(δH ) + σ 2(δN )/36 , observed in CHAPS- (blue) and DHPC-based (red) bicelles, formed at different q*. (C) Secondary structure of TrkA-TM in bicellar environment. (D) Normalized intensities of lipid-CH2/protein-NH cross-peaks in 3D 15N-edited NOESY spectra of TrkA-TM in DMPC/DHPC and DMPC/CHAPS bicelles.

samples demonstrated very similar relative intensities in the Cterminal part of the TM segment (residues 16−32), significant line broadening was observed in the first two turns of the αhelix, and the effect was much more pronounced in DMPC/ DHPC bicelles. Only three residues gave rise to broad signals in DMPC/CHAPS environment, while almost all eight Nterminal residues had very broad signals in the DMPC/ DHPC mixture (Figure 5C), and the difference was even more dramatic at 27 °C. Thus, in DMPC/DHPC bicelles, the model TM helix is destabilized in several ways: it performs fast motions in the ns time scale, that affect the rotational diffusion of the protein, and the N-terminus of the helix performs slow motions in the μs−ms time scale, which results in line broadening in NMR spectra. These effects are either not observed or weak in DMPC/CHAPS solution. Another factor that needs to be considered is the properties of protein-containing bicelles, formed with various q*. With this respect, we studied the chemical shift changes that take place in NMR spectra of TrkA-TM upon the variation of q* in DMPC/ DHPC and DMPC/CHAPS bicelles. Again, we observed that changes in generalized chemical shifts with q* are subtle and occur mainly at the edges of the TM helix (residues 8−15 and 27−35); the pattern of changes is very similar in DMPC/ CHAPS and DMPC/DHPC (Figure 6B). It is noteworthy that chemical shifts of almost all amide protons in the TM region shift in the same direction in both types of bicelles. When q*

influence, we monitored the intensities of cross-peaks and amide chemical shifts of deuterated TrkA-TM in DMPC/ DHPC and DMPC/CHAPS bicelles in 1H,15N-TROSY spectra. Chemical shifts reveal that NMR spectra of TrkA-TM in two types of bicelles are very similar; changes in generalized chemical shifts (ΔCS = ΔδH2 + ΔδN 2/36 ) are subtle and are observed mainly at the edges of a TM helix. Almost no changes in chemical shifts are detected for residues 15−28 (ΔCS < 0.02 ppm), the secondary structure is the same in DHPC- and CHAPS-based bicelles, and the transmembrane helix is formed on the region 8−34, according to NH−NH NOE contacts and rotational diffusion correlation times (Figure 6A). As the concentration of protein is different in the two samples, we used the intensity of the cross-peak from the Cterminal residue (I39) as a reference and used the ratio of intensities II39/Ix as an indicator of slow motions in the μs−ms time scale. I39 is tumbling independently from the TM helix, and the intensity of its signal in the NMR spectrum almost does not depend on the type and size of the bicelle. If slow motions are absent, all transmembrane peaks should have similar intensities, and if the slow motions occur, signal lines become broader and the ratio is increased. For this comparison, we selected conditions with almost identical rotational diffusion parameters: DMPC/CHAPS q* = 0.73 (τc = 18.5 ± 1 ns) and DMPC/DHPC q* = 0.58 (τc = 18.6 ± 1 ns). While two 6633

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bicelles. Beaugrand et al.46 introduced the lowest q, equal to 1, when the segregation between lipids and detergents was still present. Here we need to point out that the phase transition temperature does not report on the mixing between the lipid and detergent in the planar area of bicelles. According to the 31 P NMR studies of magnetically aligned DMPC/DHPC and DMPC/CHAPSO bicelles, 15−30% of detergent in the planar area does not substantially affect the character of phase transition in such systems.36,68 Phase transition is a cooperative event, and most likely, there is a minimal number of lipids in a bicelle, necessary for the phase transition to occur. Besides, the observed effects can be caused by the proximity of mobile DHPC molecules, which reside in the rim of the small bicelle. We need to admit that it is rather difficult to establish a criterion to distinguish between the mixed micelle and bicelle. However, several possible approaches are suggested. First, the structures of the surface-associated or transmembrane proteins can be investigated in various bicelles. In this manner, the work by Chou et al. demonstrated the presence of a flat surface in q = 0.25 DMPC/DHPC bicelles,8 based on the structure of the surface-bound peptide from the HIV-1 gp41 protein. In the present study, we also report the changes in NMR chemical shifts of the model TM protein in DMPC/DHPC and DMPC/ CHAPS bicelles (Figure S4). Slight q*-dependent changes are observed, but they can be interpreted in terms of stretching of the TM helix in response to the altered equilibrium thickness of lipid bilayer in bicelles, formed at different q*. Alternatively, lipid−protein interactions can be investigated. Like in the work by Lee et al.,34 we observe only the lipid−protein interactions in NMR spectra of TrkA-TM in DHPC- and CHAPS-based bicelles and interactions between the protein and rim-forming agents were not detected. This does not mean the complete absence of the detergent in a lipid bilayer region of bicelles: the detergent is in fast exchange between the water-soluble and bicelle-bound states. However, this indicates that the protein is surrounded by the coat of tightly packed lipid molecules. Since small bicelles cannot be observed directly by cryo-EM or other microscopic techniques, the judgment about the shape of the particles at low q could be made on the basis of the other data, such as the R(q) dependence. First, the radius of a spherical mixed micelle cannot exceed the length of the largest of mixed compounds; in our case, it is 2.0−2.5 nm, the length of a DMPC molecule. Thus, in all of the studied solutions, not spherical mixed micelles but rather discoidal particles are formed. However, even if the shape of a particle is discoidal, some mixing can still occur between the lipid and rim-forming agent, detergent can enter the “planar” region, thus increasing the size of the bicelle, or the lipid can migrate to the rim, which is likely to decrease the size of the particle. Therefore, we suggest that, if the bicelle size follows the ideal model (which is expressed in eqs 2 and 3), then we can assume that such mixing is negligible and the isotropic bicelles indeed are composed of a patch of lipid bilayer surrounded by the rim of detergent. According to the present work, DHPC-, CHAPS-, and CHAPSO-based bicelles follow the ideal R(q*) dependence below the phase transition temperature of the lipid bilayer, formed of the corresponding lipid, in a wide range of q*, and above the phase transition temperature at q* < 1.1 for CHAPS and CHAPSO and q* < 0.8 for DHPC. This is in agreement with the study by Glover et al.,22 performed on the DMPC/ DHPC system. Small deviations from the model are observed for CHAPS and CHAPSO at q* < 0.6−0.7, which may indicate that some mixing does occur for very small particles and 10−

and the size of bicelles is increased, almost all TM amide proton signals move downfield (Figure S4). Since chemical shifts of amide protons in proteins depend mainly on the length of the corresponding hydrogen bond,66 we can state that the TM helix becomes slightly shorter at higher q*, which may indicate that the larger bicelles have a thinner lipid bilayer. Lipid−Protein Interactions in Bicelles. The last important feature of bicelles is the state of lipids and lipid− protein interactions. To study the issue, we recorded four 3D 15 N-edited NOESY spectra of deuterated TrkA-TM in DMPC/ DHPC (q* = 0.32, 0.58) and DMPC/CHAPS (q* = 0.56, 0.98) bicelles and measured the intensities of intermolecular lipid/ protein cross-peaks. Our first goal was to find out whether there is a detectable amount of rim-forming agent, or only lipids are in close proximity to the protein molecule. In all four spectra, only signals of lipid CH2 (1.25 ppm) and terminal methyl groups (0.85 ppm) were detected, suggesting that mainly lipids are surrounding the model TM domain and the mixing between the lipid and detergent in the bilayer area is relatively low or absent (Figure S5). Besides, protein−lipid NOE crosspeaks can tell us a lot about the lipid packing and mobility around the protein. The NOE itself is proportional to τC/r6, where r and τC are the length and rotational correlation time of the vector, connecting the interacting nuclei. Thus, since the protein−lipid distances are likely to be similar in all kinds of bicelles, the differences in protein−lipid NOE intensities may report on the lipid mobility around the protein. To assess the effect, we normalized the intensities of lipid−protein crosspeaks in 15N-edited-NOESY spectra by the intensities of the diagonal peaks (to take into account the varied transverse relaxation of different residues) and additionally divided the obtained values by the overall correlation time of rotational diffusion to exclude the effect of the bicelle size. As a result, we found that regardless of the q* in DHPC-containing bicelles the magnitude of lipid−protein NOE (signal to lipid CH2-group, 1.25 ppm) is 2-fold lower than in CHAPS-containing bicelles (Figure 6D). In other words, lipids are more mobile in DMPC/ DHPC than in DMPC/CHAPS. A similar observation was reported in a recent work, when the mobility of lipids was assessed on the basis of the 13C NMR relaxation parameters.67 And this goes in line with all differences in behavior of TrkATM in CHAPS- and DHPC-based bicelles: the increased mobility of the protein in DMPC/DHPC on the ns and μs−ms time scales. Above, we report the parameters of isotropic bicelles which are necessary to understand and predict the properties of small bicelles: sizes in the presence and in the absence of TM proteins, concentrations of free detergent, properties of lipids at various q*, and temperatures. Below we would like to discuss the meaning of the obtained data, and compare the described bicelles from the point of their applicability as membrane mimetics for structural studies. Shape of Small Isotropic Bicelles. A number of recent works doubt the mere existence of small isotropic bicelles. MD investigation reveals that preassembled q = 0.25 bicelles are not stable during the simulations;46 experimental works claim that parameters of small bicelles differ a lot from the parameters of magnetically oriented bicelles: they do not demonstrate any lipid phase transition, or phase transition is not expressed and have shifted critical temperature;45,46 the size of isotropic bicelles deviates from that predicted by the models used in some works. Therefore, the authors conclude that at low q DMPC/DHPC mixtures form mixed micelles rather than 6634

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SUMMARY AND CONCLUSIONS To conclude, the present study reports the NMR-based approach that was used to investigate the radii of particles and concentration of the detergents in the monomeric state in solutions of isotropic bicelles, formed by DHPC, CHAPS, CHAPSO, and sodium cholate, as a function of lipid/detergent ratio and temperature. These parameters were measured for the bicelles composed of lipids with saturated fatty chains of different length and lipids, containing unsaturated fatty acid residue. The influence of a model transmembrane protein (membrane domain of rat TrkA) on the properties of bicelles and the effect of the bicelle size and composition on the structural and dynamic properties of the transmembrane protein were investigated. We show that isotropic bicelles that are applicable for solution NMR spectroscopy behave as predicted by the theoretical models and are likely to form discoidal particles with a patch of a lipid bilayer rather than mixed micelles. Using the obtained data, we propose a simple approach to control the size of bicelles at low concentrations. On the basis of our results, we compared the different rimforming agents and selected CHAPS as a rim-forming surfactant of choice for the structural studies if the deuteration of detergents is not required.

15% of the detergent can enter the bilayer area. In contrast, bicelles formed of the sodium cholate do not follow the models, and can serve as an example of what happens when mixed micelles are present in solution. Therefore, with all aforesaid, we conclude that low-q mixtures of lipids with DHPC, CHAPS, and CHAPSO form small isotropic bicelles with a low extent of mixing between lipid and rim-forming agents, which behave as predicted by the theoretical models. Comparison of Various Bicelles. To compare the applicability of various membrane mimetics, which were under investigation in the present study, we need to formulate the requirements that determine the usefulness of bicelles in structural studies. First, we need to point out that small bicelles of CHAPS and CHAPSO are almost identical; therefore, only CHAPS and DHPC will be considered below. From the point of the solution NMR, the size of the particles is one of the major factors, affecting the quality of NMR spectra. Both CHAPS and DHPC can form bicelles with a wide range of radii, up to 2.2−2.3 nm; however, other things being equal, one would like to maximize the amount of lipids in a bilayer to enhance the adequacy of the membrane mimetic. To investigate this aspect, we calculated the amount of lipids in different bicelles and LPNs. Assuming that the density of lipids in a bilayer is the same for all membrane mimetics (which is most likely a very rough approximation), the amount of lipids in particles of the same size depends on the thickness of the rim, r⊥. This parameter equals 1.1 nm for DMPC/CHAPS, equals 1.15−1.2 nm for DMPC/DHPC, and has to exceed ∼1.5 nm in the case of LPNs. Therefore, CHAPS, out of all tested rim-forming agents, yields bicelles with the maximal contents of lipids in a bilayer region; however, the difference between DHPC and CHAPS in this regard is almost negligible. Apart from the bicelle size, the stability and nativity of the spatial structure of transmembrane and water-soluble domains of membrane protein in a bicelle are also of great importance. We show here that membrane protein is likely to be more stable in CHAPS-based environment: in DMPC/DHPC, the model TM domain had an increased mobility in both ns and μs−ms time scales. Besides, there are a number of examples when CHAPSO-containing bicelles demonstrated the better functionality of reconstituted protein,69,70 pH,71 and temperature stability70 than other bicellar mixtures. In our recent work, we showed that the globular water-soluble domain of the p75 receptor is unfolded in DMPC/DHPC and retains its native structure in DMPC/CHAPS solution.18 As we show here, CHAPS is characterized by a lower concentration of monomeric detergent in bicelles than DHPC, which facilitates the control of the bicelle size at low concentrations. Last but not least, CHAPS is three times cheaper than CHAPSO and 9 times cheaper than DHPC. Thus, according to all listed criteria, CHAPS is the detergent of choice for isotropic bicelles used in structural studies of membrane proteins. On the other hand, there is one big advantage of DHPC that is relevant mainly for solution NMR spectroscopy. Sometimes it is useful to have lipids and detergents in deuterated formto optimize the transverse relaxation and get rid of unwanted intense crosspeaks in NMR spectra. While DHPC is commercially available in deuterated form, we found no works, reporting the synthesis or other way of production of deuterated CHAPS, CHAPSO, and other sterols.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.langmuir.6b00867. Six tables with measured parameters of various bicelles and five additional figures (PDF)

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Author Contributions

The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The work is supported by the Russian Science Foundation, grant no. 14-14-00573.

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ABBREVIATIONS NMR, nuclear magnetic resonance; TM, transmembrane; MP, membrane protein; LPN, lipid−protein nanodisc; HSQC, heteronuclear single quantum correlation spectroscopy; NOESY, nuclear Overhauser effect spectroscopy; TROSY, transverse relaxation optimized spectroscopy; DLS, dynamic light scattering; SANS, small-angle neutron scattering; SAXS, small-angle X-ray scattering; cryo-EM, cryogenic electron microscopy; DMPC, 1,2-dimyristoyl-sn-glycero-3-phosphocholine; DMPG, 1,2-dimyristoyl-sn-glycero-3-phosphoglycerol; DPPC, 1,2-dipalmitoyl-sn-glycero-3-phosphocholine; POPC, 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine; DHPC, 1,2dihexanoyl-sn-glycero-3-phosphocholine; CHAPS, 3-[(3cholamidopropyl)dimethylammonio]-1-propanesulfonate; 6635

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CHAPSO, 3-[(3-cholamidopropyl)dimethylammonio]-2-hydroxy-1-propanesulfonate

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DOI: 10.1021/acs.langmuir.6b00867 Langmuir 2016, 32, 6624−6637