Solid-State NMR Spectra of Lipid-Anchored Proteins under Magic

Article pubs.acs.org/JPCB

Solid-State NMR Spectra of Lipid-Anchored Proteins under Magic Angle Spinning Kaoru Nomura,* Erisa Harada, Kenji Sugase, and Keiko Shimamoto Bioorganic Research Institute, Suntory Foundation for Life Sciences, 1-1-1 Wakayamadai, Shimamoto-Cho, Mishima-Gun, Osaka 618-8503, Japan S Supporting Information *

ABSTRACT: Solid-state NMR is a promising tool for elucidating membranerelated biological phenomena. We achieved the measurement of high-resolution solid-state NMR spectra for a lipid-anchored protein embedded in lipid bilayers under magic angle spinning (MAS). To date, solid-state NMR measurements of lipid-anchored proteins have not been accomplished due to the difficulty in supplying sufficient amount of stable isotope labeled samples in the overexpression of lipid-anchored proteins requiring complex posttranslational modification. We designed a pseudo lipid-anchored protein in which the protein component was expressed in E. coli and attached to a chemically synthesized lipid-anchor mimic. Using two types of membranes, liposomes and bicelles, we demonstrated different types of insertion procedures for lipid-anchored protein into membranes. In the liposome sample, we were able to observe the crosspolarization and the 13C−13C chemical shift correlation spectra under MAS, indicating that the liposome sample can be used to analyze molecular interactions using dipolar-based NMR experiments. In contrast, the bicelle sample showed sufficient quality of spectra through scalar-based experiments. The relaxation times and protein−membrane interaction were capable of being analyzed in the bicelle sample. These results demonstrated the applicability of two types of sample system to elucidate the roles of lipid-anchors in regulating diverse biological phenomena.

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INTRODUCTION Protein lipidation occurs in many kinds of proteins, including cell adhesion molecules, receptors, scaffolding proteins, enzymes, etc. These proteins are anchored to membranes through a covalent link to fatty acids, isoprenoid, sterol, or glycosylphosphatidylinositol (GPI).1,2 The main purpose of protein lipidation is to tether proteins near the membrane surface.3 Lipid anchoring is an important process for adhesion molecules and scaffolding proteins to function.4−6 The trafficking and sorting of proteins to specific domains, namely the apical side of epithelial cells or microdomains called lipid rafts, is also an important role played by protein lipidation.7−9 The localization of lipidated protein depends on the degree of fatty acid saturation.8 Dynamic switching of the lipidation state by enzymes also regulates protein−protein interactions. For example, the activation/inactivation cycles of some G proteincoupled receptors (GPCRs) are regulated by conversion of the lipidated/nonlipidated states of the G protein α and γ subunits, respectively.10,11 Furthermore, lipid moieties have a role in disease pathogenesis.12−14 For example, defects in GPIanchoring of CD59 cause paroxysmal nocturnal hemoglobinuria (PNH) due to a loss of function in the inhibition of the formation of the membrane attack complex (MAC), which consists of membrane complement components.12 A conformational change in the prion protein after membrane insertion is known to be one of the important triggers for fatal © 2014 American Chemical Society

neurodegenerative diseases such as transmissible spongiform encephalopathies (TSEs).13,14 Thus, protein lipidation plays crucial roles for regulation of many biological events, primarily cellular signal transduction, by controlling protein localization, protein−membrane and protein−protein interactions, and their structure on the membranes. To reveal the molecular mechanisms and functions of lipidanchored proteins in membranes, it is indispensable to determine their 3D structures, dynamics, and membrane interactions at the atomic level. To this end, solid-state NMR spectroscopy is a promising technique. Although solid-state NMR studies of integral membrane proteins have previously been performed,15−19 entire lipid-anchored proteins have not been studied yet,20−22 because of technical difficulties in supplying sufficient amounts of stable isotope-labeled samples for NMR measurements of lipid-anchored proteins. First, because the protein lipidation is a posttranslational modification, overexpression of lipidated proteins in E. coli is difficult. Second, although certain acylated proteins could be produced by coexpression with transferase in E. coli, not all enzymes are available for biosynthesis of the lipid-anchor component. Finally, the protein component as well as the anchor Received: December 18, 2013 Revised: February 11, 2014 Published: February 11, 2014 2405

dx.doi.org/10.1021/jp4124106 | J. Phys. Chem. B 2014, 118, 2405−2413

The Journal of Physical Chemistry B

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Figure 1. (a) Schematic representation of a lipid-anchored protein and of the sample preparation of anchored proteins in liposomes (b, top) and bicelles (b, bottom). (c) HPLC purification of the anchored proteins from the reaction mixture of anchor and GB1. (d) MALDI-TOF-MS spectra of the anchorless GB1 (top) and anchored GB1 (bottom). (e) Gel filtration profiles of reaction mixtures (top) and the eluted fraction containing the anchored-protein in bicelles (bottom). (f) 31P NMR spectrum of anchored GB1 in bicelles.

component will be isotope labeled, which complicates NMR spectra. Moreover, the production of a GPI-anchor, which consists of a phosphoethanolamine linker, glycan core, and a phospholipid tail, in eukaryotic cells results in heterogeneous mixtures with variations in the glycan core and lipid moieties.23,24 To overcome these problems in the present study, we chemically synthesized anchor-like substitute and attached it to a protein that was expressed in E. coli. This strategy would be useful for proteins with C-termini lipidated by a highly functionalized anchor, such as GPI. Because the sample preparation for NMR analysis of GPI-anchored protein is more difficult than that of other lapidated proteins, we used a new type of anchor possessing a diacylglycerol tail (Figure 1a and Scheme S1 in the Supporting Information) to resemble the GPI-anchor. This is the first solid-state NMR study to show the feasibility of a methodology for the analyses of the protein component of lipidated proteins embedded in membranes.

Although several techniques to produce proteins with anchorlike substitutes have been developed to examine the effect of membrane-anchoring on their 3D structure, distribution, and diffusion kinetics in membranes using circular dichroism (CD) and fluorescence observations,25−28 analyses at the atomic level are difficult using these measurement techniques. Because NMR studies require higher concentrations of proteins than CD and fluorescent measurements, we verified the applicability of this NMR measurement methodology using the B1 domain of the IgG-binding protein G (GB1) as a model protein. Even though GB1 is not an actual lipid-anchored protein, it is suitable for validation because it has been well characterized as a standard sample for NMR studies.29−34 We first provided evidence showing high resolution solid-state NMR spectra for a lipid-anchored protein embedded in membranes under magic angle spinning (MAS).35 2406



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C were mixed and vortexed for 15 min at 4 °C. The sample was then placed in a water bath at 38 °C for 30 min. The resulting mixture was extremely viscous but became fluid after shaking in an ice bath. Repeating the heating (38 °C) and cooling (4 °C) cycles four times provided a solution containing 25% w/v bicelles composed of DMPC/anchor/DHPC-C7 at a molar ratio of 3:0.06:1 in buffer C (550 μL). The coupling reaction and purification were carried out at 4 °C. To reduce dimers at cysteine residues, 7.2 mg of GB1 was reacted with an equimolar amount of TCEP (0.26 mg) in 10 mM HEPES (133 μL) for 1 h at room temperature. This was mixed with 267 μL of bicelle solution (25% w/v) containing the anchor mimic and shaken at 1500 rpm at 4 °C for about 16 h. Uncoupled GB1 was removed by loading 200 μL of the reaction mixture onto a gravity-flow Biospin column (37 mm working bed height, 1.2 mL bed volume, BioRad) containing Sephadex G-75 (Pharmacia) with buffer C. Each fraction was checked by gel filtration chromatography (Superdex 75, 5 × 150 mm) using the Ä KTA purifier fast performance liquid chromatography (FPLC) system (GE Healthcare) (Figure S1 in the Supporting Information). From these analyses, we selected the fraction that included the anchored GB1 in bicelles without the anchorless GB1. The two fractions from the gravity-flow column were then concentrated using an Amicon Ultra centrifugal filter unit (3.5 kDa molecular weight cutoff, Millipore). Finally, a sample containing 3.7 mg of anchored GB1 and 22% lipids was obtained in a 100 μL solution. Preparation of Paramagnetic Ion Containing Bicelle Samples. All lipids (0.814 mg of 18:1 DGS-NTA, 109 mg of DMPC, 25.8 mg of DHPC-C7, and 2.5 mg of anchor) were mixed at a molar ratio of [DMPC]:[DHPC-C7]:[anchor]:[18:1 DGS-NTA] = 1:0.33:0.02:0.005 and dissolved in chloroform. After solvent evaporation under vacuum for 16 h, the lipid film was hydrated with 550 μL of buffer C and vortexed. Thereafter, heating (38 °C)/cooling (4 °C) cycles were repeated four times. The coupling reaction between GB1 and bicelles was the same as described in the previous paragraph except that 0.318 mg of MnCl2 was added to 267 μL of bicelle sample after mixing with protein and shaken at 1500 rpm at 4 °C for about 1 h before uncoupled GB1 was removed using a gravity-flow Biospin column. The paramagnetic ions that did not reside on the membrane surface were also removed by the column. Solid-State NMR Measurements. All solid-state NMR spectra were acquired on a Bruker Avance III 600 spectrometer (Bruker Biospin, Billerica, MA) equipped with a narrow-bore magnet operating at a resonance frequency of 150.13 MHz for 13 C, 60.81 MHz for 15N, 242.93 MHz for 31P, and 600.13 MHz for 1H. Experiments were recorded with an E-free tripleresonance and VTN double-resonance 4 mm MAS probe. 1D 31 P NMR spectra were acquired under static conditions. In the 2D 13C−13C DARR experiments, the MAS rate was 13.5 kHz. The other measurements were performed under 5 kHz MAS. The lengths of 90° pulses on 13C, 15N, 31P, and 1H were 4.9, 6.2, 5.0, and 4.9 μs, respectively. For the 1D 13C, 2D 13C−1H, and 2D 15N−1H refocused INEPT experiments, 2.5 kHz GARP decoupling was applied during the acquisition. In the 1D 31P, 13 C CPMAS, and 2D 13C−13C DARR experiments, 51 kHz SPINAL-64 1H decoupling39 was performed during acquisition. The 1D 13C cross-polarization (CP) MAS spectra were acquired using a ramped (from 50 to 100%) spin-lock pulse on the 1H channel and square contact pulse on the 13C channel during a 2 ms contact time. In the 2D 13C−13C DARR

EXPERIMENTAL SECTION Materials. DMPC (1,2-dimyrystoyl-sn-glycero-3-phosphocholine), DHPC-C6 (1,2-dihexyanoyl-sn-glycero-3-phosphocholine), DHPC-C7 (1,2-diheptanoyl-sn-glycero-3-phosphocholine), and DGS-NTA (1,2-dioleoyl-sn-glycero-3-[(N-(5amino-1-carboxypentyl)iminodiacetic acid)succinyl]) were purchased from Avanti Polar Lipids (Alabaster, AL) and used without further purification. All chemicals were purchased from Nacalai Tesque (Kyoto, Japan). Sample Preparation Method for Liposome-Bound GB1. To obtain completely free sulfhydryl groups, 8 mg of GB1 was reacted with TCEP (10 mM) in 10 mM HEPES (25 mL) for 1 h at room temperature. TCEP was removed using a desalting column (PD-10) with 10 mM HEPES buffer. The concentration of the resulting reduced GB1 was about 25 μM. A solution of lipid-anchor mimic 3 (Scheme S1 in the Supporting Information) (0.84 mM) in DMSO (6 mL) was added to the GB1 solution (in 10 mM HEPES buffer, 30 mL). The mixture was then shaken at 250 rpm at 37 °C for about 16 h. To remove the uncoupled GB1 and lipid-anchor mimic from the reaction mixture, the sample was applied to a C4 packed HPLC column (46 × 150 mm (analytical), 200 × 150 mm (preparative), Waters) and eluted with an acetonitrile/water gradient (10% to 95%) in the presence of 0.1% (v/v) TFA at 50 °C. The flow rate is 1 mL/min for analytical and 15 mL/min for preparative. The uncoupled GB1, anchored protein, and uncoupled anchor were eluted at 23, 48, and 61 min, respectively. Because the acyl bonds at sn-1 and sn-2 of the anchor were easily hydrolyzed at pH < 2, the eluted anchoredprotein was immediately neutralized with aqueous ammonia. To remove ammonium trifluoroacetate, the sample was applied to a second C4 packed HPLC column and eluted with a water/ acetonitrile gradient in the presence of 1% (v/v) formic acid at 50 °C. Eluted fractions were neutralized with aqueous ammonia and subsequently lyophilized. The lyophilization process was repeated three times to completely remove ammonium formate. The structures of the obtained samples were confirmed by MALDI-TOF-MS. Anchored GB1 was reconstituted into liposomes as follows. The liposomes were prepared as described previously.36−38 A suspension of DMPC liposomes (12.5 mM, 1 mL) was solubilized with 200 μL of 10% n-dodecyl β-D-maltoside (DDM). To a solution of DMPC liposomes was added 2 mg of the anchored protein solubilized in 500 μL of buffer B (10 mM HEPES, 100 mM KCl, pH 7.2) to produce a protein:DMPC ratio of 1:50. The mixture was then equilibrated at room temperature for 2 h with gentle agitation. To remove the detergent, 40 mg (wet weight) BioBeads SM2 absorbent (BioRad) was added to the mixture, and the suspension was gently shaken at room temperature. The BioBeads were replaced every 12 h for a period of 60 h (five changes) and liposomes containing the anchored protein were collected by pipetting. The liposome suspension was ultracentrifuged (at 165 000 g for 1 h) and the pellet was washed twice with 1 mL of buffer B. The protein concentration was determined using a CBQCA Protein Quantitation Kit (Molecular Probes). Approximately 1.2 mg of the anchored protein in 50 μL of pellet was obtained. Sample Preparation Method for Anchored GB1 in Bicelles. A suspension of 109 mg of DMPC/2.5 mg of anchor in 275 μL of buffer C (10 mM HEPES, 50 mM KCl, pH 7.2) and a suspension of 25.8 mg of DHPC-C7 in 275 μL of buffer 2407



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experiments, a rotary resonant 1H field (13.5 kHz) was applied during the DARR mixing period. The actual temperature in the sample under 5 kHz MAS was calibrated using chemical shift differences between the water and methyl peaks in the low hydrated DMPC sample.40 All temperatures quoted are calibrated temperatures. Chemical shifts were externally referenced to sodium 2, 2-dimethyl-2-silapentane-5-sulfonate (DSS) using the downfield 13C resonance of adamantane (40.48 ppm).41 To prevent drying of the sample, the NMR tube was capped with a drive chip with a sealing function (Phi Creative).

mixture must contain certain levels of organic solvent (20% DMSO in this study). This solvent system might produce undesirable denaturing of the protein.48,49 In addition, labile proteins may decompose during HPLC purification. In contrast, anchors incorporated into bicelles can react with proteins in a purely aqueous buffer system without the need for organic solvents. Therefore, proteins need not to be exposed to severe environments. 1D Spectra of Anchored GB1 in the Presence and Absence of Membrane. Solid-state NMR were measured for the aforementioned preparations under 5 kHz MAS. 13C refocused INEPT50 (Figure 2a at 32 °C) and cross-polarization

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RESULTS AND DISCUSSION Preparation of Anchored GB1 in Membranes. The GB1 protein was attached to a decapeptide linker that included a reactive cysteine residue at the C-terminus to adjust the distance from the phosphate of the anchor to that of the GPI anchor as a representative of lipid anchor. 13C- and 15N-labeled GB1 was easily obtained by overexpression in E. coli and purified according to a previous report.42 To anchor the protein, we prepared two different types of membranes, liposomes (Figure 1b, top) and bicelles (Figure 1b, bottom). For the first method (Figure 1b, top), purified GB1 was reacted with the maleimide group in the lipid anchor mimic ([anchor]:[GB1] = 6.7:1) prior to reconstruction into liposomes. The HPLC profile for the reaction mixture indicated that approximately one-half of the GB1 reacted with the anchors (Figure 1c). Figure 1d shows MALDI-TOF MS spectra of the anchorless GB1 (top) and the anchored protein (bottom). The difference in the measured mass values was 785, which corresponds to the mass of the anchor. The doublet peaks in both spectra originated from heterogeneity arising from the partial cleavage of the N-terminal methionine in GB1 during overexpression in E. coli.43 Subsequently, we reconstituted the anchored protein into DMPC liposomes using a detergent mediated method.44 In the second method (Figure 1b, bottom), after the anchorcontaining bicelles ([DMPC]:[DHPC-C7]:[anchor] = 3:1:0.06) were formed, they were reacted with GB1 ([anchor]:[GB1] = 2:1). The uncoupled GB1 was removed using a gravity-flow column (Sephadex G-75). The absence of unbound GB1 was evaluated by Superdex 75 gel filtration chromatography of each fraction (Figure 1e, see also Figure S1 in the Supporting Information). Bicelle morphology was confirmed by 31P NMR spectra under static conditions (Figure 1f). Each method has certain advantages. The liposomes in the first method are more stable to hydration than the bicelles in the second method. The bicelle samples are prone to excessive hydration, which may induce morphological changes that produce mixtures of multilamellar vesicles and micelles.45 Therefore, we used DHPC-C7 to produce bicelle samples that are stable over a wider range in hydration. Bicelle samples produced with DMPC/DHPC-C7 were more resistant to a hydration than bicelles produced with DMPC/DHPC-C6 because of the lower critical micelle concentration (cmc) for DHPC-C7 (1.6 mM) compared to that for DHPC-C6 (15 mM).46,47 On the other hand, the conditions for the coupling reaction between the anchor and the protein in the second method are milder than in the first method. The first method requires the protein to be reacted with the anchor before reconstruction into the membranes. Because the anchor was unable to be dispersed in the aqueous buffer, the reaction

Figure 2. (a) 13C refocused INEPT spectrum of anchored GB1 in liposomes (containing 1.2 mg of anchored protein), (b) CP spectra, and (c) CP spectra at 4 °C. A total of 7000−14000 scans were coadded for liposome samples. (d) 13C refocused INEPT of the anchorless GB1 dissolved in buffer, 6000 scans. (e) 13C refocused INEPT spectrum of the anchored GB1 in bicelles (containing a 3.7 mg anchored GB1), (f) CP spectra, and (g) CP spectra at 4 °C. A total of 1200−2400 scans were coadded for bicelle samples. All spectra were acquired under 5 kHz MAS at 32 °C, except for (c) and (g), which were obtained at 4 °C. The peaks from lipids are marked as *.

(CP)51 (2b at 32 °C and 2c at 4 °C) spectra were successfully obtained for the anchored protein in liposomes. It is generally known that magnetizations of mobile components are selected by scalar transfer using INEPT, whereas magnetizations of rigid components are observed using CP by dipolar transfer. At 32 °C, both GB1 and lipid peaks appeared in the INEPT spectrum (Figure 2a), whereas almost all resonances in the CP spectrum (Figure 2b) were attributed to the lipids. These results suggest that the lipids in liposome are rigid, whereas the protein is flexible to the extent that 1H−13C dipolar couplings required 2408



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Figure 3. 2D 13C−13C chemical shift correlation spectra of liposome anchored GB1 with 10 and 50 ms dipolar assisted rotational resonance (DARR) mixing times acquired under 13.5 kHz MAS at 4 °C. The data size was 200 points with a 26 μs dwell time in t1 and 2048 complex points with a 15 μs dwell time in t2. The pulse delay was 2 s. The data were Fourier transformed using zero filling to 4096(ω2) × 1024(ω1) complex points with SINE = 3 apodization and 100 Hz exponential line broadening in the t1 and t2 dimensions, respectively. Color circles overlaid are artificial correlations referring the solution NMR chemical shifts of GB1 reported by Wilton et al. (BMRM entry 7280)34 for aliphatic carbons and by Franks et al. (BMRM entry 15156)31 for carbonyl carbons in both main and side chains (one-bond correlation in blue, two-bond correlation in red, three-bond correlation in orange, and four-bond correlation in yellow).

circles based on the chemical shift values of GB1 reported by Wilton et al. (BMRM entry 7280)34 for aliphatic carbons and by Franks et al. (BMRM entry 15156)31 for carbonyl carbons in both main and side chains. These patterns are in good agreement with the experimental data. Thus, we realized that the observed cross peaks are from intra residue dipolar interaction in GB1. The strong two- and three-bond correlation peaks appeared at 50 ms mixing, e.g., Thr Cα-Cγ, Val Cα-Cγ, Lys Cγ-Cε, Leu Cα-Cγ, and Leu Cα-Cδ, and Lys Cα-Cδ, whereas only subsidiary two-bond correlation peaks appeared at 10 ms mixing. These results suggest that almost all residues in the protein component are rigid at 4 °C. Because lipids are not 13 C-labeled, there are no correlation peaks from lipids. The spectral patterns in Figure 3a,b were practically the same as those reported for microcrystalline GB1 with mixing of 5 and 25 ms,31 with the exception of line broadening. Line broadening shown in our sample most likely originated from the larger conformational heterogeneity in GB1 than in microcrystalline samples. To obtain well resolved spectra, we plan to use sparsely 13C-labeled samples prepared with 1,3-13Cor 2-13C-glycerol.30 Franks et al. reported that longer mixing, for 500 ms, yields a strong correlation between carbons separated by 5.8−8.2 Å in microcrystalline samples.30 Therefore, using longer mixing times will probably provide correlation peaks between anchored proteins and other integral proteins. If we use labeled lipid, it could be possible to observe the correlation peaks between anchored protein and membrane. 2D 1H−13C and 1H−15N INEPT Spectra of Anchored GB1 in the Bicelles. Figure 4 shows the 2D 1H−13C (a) and 1 H−15N (b) refocused INEPT spectra of anchored GB1 in bicelles (red) acquired at 5 kHz MAS at 32 °C. In both spectra, the spectral resolution was comparable to that of the anchorless

for CP were almost averaged out. When motion became significantly slower at a lower temperature (at 4 °C), the CP peaks of the protein component were detected (Figure 2c). Compared to the refocused INEPT spectrum of the anchorless GB1 dissolved in the buffer (Figure 2d), peaks of the anchored GB1 in the liposomes were broadened by relaxation during the refocused INEPT sequence (Figure 2a), suggesting that the anchoring at the C-terminus partially restricts protein mobility. The possibility that GB1 nonspecifically binds to liposomes was ruled out because no peaks originated from GB1 were observed when we performed the same membrane reconstitution procedure using anchorless GB1 (data not shown)). On the other hand, the 13C refocused INEPT (Figure 2e) of anchored GB1 in bicelles provided spectra with higher resolution than in liposomes (Figure 2a). In the CP spectrum, only lipid peaks were observed at 32 °C (Figure 2f), similar to that in the liposome samples. The results at 32 °C indicate that the protein and lipids in bicelles in a “Swiss-cheese”-like phase above the phase-transition temperature, Tm (25 °C),45 move faster than in liposomes in the liquid crystalline phase, and that the mobility of the lipids is restricted compared to that of the protein. In contrast, both the lipid and protein peaks disappeared at 4 °C (Figure 2g) unlike in the liposome sample. The lack of peaks in the CP spectrum at 4 °C implied that the mobility in this sample is higher than in the liposomes in the gel phase, reflecting a fluid-like phase composed of disk shaped assemblies below Tm. 2D 13C−13C DARR Spectra of Anchored GB1 in Liposomes. The 2D 13C−13C chemical shift correlation spectra of anchored GB1 in liposomes were acquired with 10 and 50 ms dipolar assisted rotational resonance (DARR) mixing times at 4 °C (Figure 3). We also indicated the estimated intra-residue correlations up to 4 bonds apart in color 2409



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from conformational fluctuation between the monomer and dimer states. Utilizing the high resolution spectra, as shown in Figure 4b, we determined residue specific R1N values of anchored GB1 in bicelles using the pulse sequence in Figure S2 (Supporting Information). A series of 2D 1H−15N refocused INEPT spectra were acquired with increasing delay time, τ, in the range 0−0.5 ms. Resulting R1N values were compared to those reported in the solution NMR experiment done by Idiyatullin et al.53 in Figure S3 (Supporting Information). The average R1N value was 1.63 s−1, which is 80% of that reported in the solution NMR analysis, 2.25 s−1. These results implied that anchoring to bicelles slightly reduces protein mobility. Determination of Protein Regions Residing in Proximity to the Membrane Surface. Next, we performed a paramagnetic relaxation enhancement (PRE) experiment to determine which part of the protein is closest to the membranes.54−59 Because paramagnetic ions induce relaxation of nuclei depending on the electron-nuclei distance, it has been routinely used to obtain structural information in solution NMR60−62 and is also being applied to solid-state NMR.33,63,64 Regarding the membrane sample, using PRE effects, Tuzi et al. demonstrated the signal assignments of residues residing near the membrane surface.55 Additionally, Buffy et al. determined the depth of peptides in membranes.56 We determined residues in the vicinity of the membrane surface using a bicelle sample in which manganese ions are located on the membrane surface, by chelating them in the headgroup of the lipid components. Figure 5a shows the comparison of INEPT spectra of anchored GB1 in bicelles in the presence and absence of Mn2+ chelated to the lipid head. Appreciable pseudocontact shifts due to Mn2+ were not observed. Some peaks show remarkable loss of their intensities in the presence of Mn2+ ion. The intensity reduction, measured as the ratio I = (Ipara/Inonpara)/(Ipara/Inonpara)max between the sample in the presence and absence of Mn2+ ions, are shown in Figure 5b as a function of the residue number. In Figure 5c, the residues that lost more than 70% of their intensity are mapped on the GB1 structure in red. The region from residue 34, the terminal region of α-helix, to residue 43, the beginning of third β-strand (β3), and residue T11 are located near the membrane. These residues are geometrically close to the C-terminus. Therefore, the anchor component is probably extended nearly perpendicular to the membrane surface, whereas the protein component has no specific domains interacting with the membrane. If we apply this method to proteins that interact with the membrane, we will identify which part of a lipidated protein interacts with the membrane as well as proteins in general.

Figure 4. Overlay of the 2D 1H−13C (a) and 1H−15N (b) chemical shift correlation spectra recorded on the anchored protein in bicelles (red) and the anchorless GB1 dissolved in buffer (blue) with 5 kHz MAS. The 2D 13C−1H refocused INEPT experiments were obtained by recording 300 t1 increments with a 120 μs dwell time and 400 scans each. The 2D 15N−1H refocused INEPT experiments were obtained by recording 100 t1 increments with a 240 μs dwell time and 1000 scans each. Cross peaks in Figure 3b were assigned by referencing them to the reported GB1 chemical shift values.34 The peaks that originated from the residues of the linker and some minor peaks are surrounded by broken line rectangles. All spectra were obtained at 32 °C.

GB1 dissolved in buffer (blue). The 1H−13C spectra (Figure 4a) have pronounced 13C−13C J-splittings in the 13C dimension, indicating that the 13C−1H and 1H−1H dipolar couplings were motionally averaged throughout the protein. In Figure 4a, most of the peaks overlapped. The peak of the Cterminal cysteine was observed only in the solution sample, whereas peaks detected only in the bicelle sample were attributed to the lipids. The lipid peaks appeared at different chemical shift values from the protein peaks, and thus did not disturb the spectral analysis of the protein. In the 2D 1H−15N refocused INEPT spectrum (Figure 4b), some peaks in the bicelle sample were weakened because anchoring restricted protein mobility. Chemical shift values of the protein component were nearly identical regardless of anchoring; suggesting that binding to the membranes does not affect the protein 3D structure. The only residue that shifted after membrane anchoring is E19. GB1 has been reported to form dimers through the interface between β2 (residue 13 to 19) and β3 (residue 42 to 46) strands in the wild type crystalline state.52 GPI anchored proteins are also known to have a tendency to induce a dimer formation.9 Therefore, the shift at E19 may arise

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CONCLUSIONS In this study, we obtained solid-state NMR spectra of a pseudo lipid-anchored protein embedded in two kinds of lipid membranes. Selection of the membrane system depends on the intended purpose because each system offers different advantages. The bicelle sample showed sufficient quality of 1D and 2D INEPT spectra of the anchored protein at 32 °C (Figures 2e and 4). Furthermore, the relaxation time analyses of each backbone amide revealed that protein mobility decreases with tethering by an anchor (Figure S3, Supporting Information). In addition, PRE measurement in the bicelle sample demonstrated that the residues which reside geometrically close to C-terminus are located near the membrane (Figure 5). In this manner, we have seen bicelle samples are 2410



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anchor mimic. Because the structure of the GPI anchors is extremely complicated, synthesizing the entire GPI anchor is impractical for the purpose of NMR studies. Although our proposed anchor lacks a glycan core component, we compensated for this by adding a peptide linker in its place. Our anchor is a comprehensive mimic of the lipid component of lipidated proteins. By modifying this anchor, we will be able to focus on specific lipidated proteins. Anchoring to membranes enables the investigation of the behavior of lipidated proteins more correctly than when an anchorless protein in solution is used. The current results collected from the solid-state NMR method deserve further study, with the protein components being replaced with actual lipid-anchored proteins to examine the roles of protein−membrane and protein−protein interactions on membranes. These interactions are responsible for regulating diverse biological phenomena, such as embryonic development, immune responses, and diseases related to lipid-anchored proteins.

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ASSOCIATED CONTENT

S Supporting Information *

Figure 5. (a) Comparison of INEPT spectra of anchored GB1 in bicelles in the presence and absence of Mn2+. Data acquisition parameters were the same for the spectra shown in Figure 4. The peaks that originated from the residues of the linker and some minor peaks are surrounded by broken line rectangles. (b) Relative intensities of the cross peaks in the 2D 1H−15N INEPT spectra of anchored GB1 in the bicelle in the presence and absence of Mn2+ chelated to the lipid as a function of residue number. Relative intensities were calculated using the equation I = (Ipara/Inonpara)/(Ipara/Inonpara)max, where Ipara and Inonpara are the peak integrals in the presence and absence of Mn2+. The I value of residues with overlapping peaks were set to zero. (c) Residues that lost greater than 70% of their intensity were mapped to the ribbon domain representation of the GB1 structure, depicted in red.

Detailed protocol of the synthesis of lipid-anchor mimic and expression and purification of cysteine-tagged GB1; synthetic scheme of the lipid-anchor mimic; gel filtration profiles; schematics of NMR pulse sequences used for measurements of backbone amide 15N longitudinal relaxation rate constant R1N; the backbone amide 15N longitudinal relaxation rate constant R1N of GB1. This information is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*K. Nomura: e-mail, [email protected]. Notes

The authors declare no competing financial interest.

most applicable for 3D structure, dynamics, and membrane interaction studies of lipid-anchored protein samples using scalar-based NMR experiments.65 In contrast, the liposome samples enable us to observe the protein peaks in CP and 2D DARR measurements at low temperature (4 °C) (Figures 2c and 3). This results from the fact that 13C−1H and 13C−13C dipolar couplings remain when the lateral diffusion of the membrane lipid is drastically reduced in the gel phases.66 The peak detection of both the protein and lipids allowed us to analyze inter- and intramolecular interactions through dipolarbased experiments. Recently, Mazab-Jafari et al. reported solution-state NMR observations of a GTPase tethered to a nanodiscs.67 The nanodisc is a competent system used to obtain high-resolution solution-state NMR spectra of membrane-associated molecules without MAS.67−70 On the other hand, solid-state NMR provided great advantage for observing the dynamics of oligomer/complex structures in membranes because liposomes or bicelles have smaller effects on the lateral diffusion rate and no limitation in the number of molecules contained in the membrane, in contrast to nanodiscs. By combining the information from the solution NMR measurement using nanodiscs with that from the solid-state NMR measurement using liposomes or bicelles, we will be able to gain a deeper understanding of the molecular mechanism of lipid-anchored proteins in membranes. In conclusion, we succeeded in analyzing the structure and dynamics of lipid-anchored GB1 to membranes. To tether the protein domain near the membranes, we used an anchor with a diacylglycerol tail and a phosphoethanolamine linker, as a GPI-

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ACKNOWLEDGMENTS We thank Dr. Kenichi Morigaki for his helpful discussions regarding the membrane sample preparations. We also thank Dr. Makoto Suematsu for his valuable comments. This work was supported by a Grant-in-Aid for Scientific Research (#90353515 and #90353515) to KN from the Ministry of Education, Culture, Sports, Science and Technology of Japan. The authors also thank Suntory Holdings for their financial support.

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Solid-State NMR Spectra of Lipid-Anchored Proteins under Magic

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