Rapid Report pubs.acs.org/biochemistry
Specific Lipid Binding of Membrane Proteins in Detergent Micelles Characterized by NMR and Molecular Dynamics Linlin Zhao,†,⊥ Shuqing Wang,‡,⊥ Changqing Run,† Bo OuYang,*,† and James J. Chou*,†,§ †
State Key Laboratory of Molecular Biology, National Center for Protein Science Shanghai, Shanghai Institute of Biochemistry and Cell Biology, Shanghai Science Research Center, Chinese Academy of Sciences, Shanghai 200031, China ‡ School of Pharmacy, Tianjin Medical University, Tianjin 300070, China § Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School, Boston, Massachusetts 02115, United States S Supporting Information *
lipids.14 In terms of sample preparation, the detergent/lipid mixed micelle is as convenient to use as detergent micelles. An important criterion of the mixed micelle application, however, is whether the detergent is compatible with specific lipid− protein interaction that is functionally important to many membrane proteins.15 Considering this, we sought to investigate whether dodecylphosphocholine (DPC), a relatively strong and the most frequently used detergent in NMR, can preserve the binding of functionally important lipids to protein. As such, we chose the mitochondrial ADP/ATP carrier (AAC) as a case study because this transporter requires special lipids to maintain its native activity16 and, when reconstituted in DPC micelles, exhibited favorable spectroscopic properties.17 The AAC is an essential solute transporter that catalyzes the trafficking of ATP and ADP between mitochondria and cytosol;18 it has been the model system for structural and mechanistic studies of mitochondrial carriers19 because it remains, to date, the only carrier protein for which highresolution crystal structures are available.20,21 Earlier functional studies showed that cardiolipin (CL), a rich component of the mitochondrial membrane and a nonbilayer lipid, is important for the transport activity of AAC.22,23 Consistent with this finding, high-resolution crystal structures of AAC previously determined in the decyl maltoside (DM) detergent revealed several bound CLs on the periphery of the transporter.20,21 Obviously, CL binding was preserved when AAC was extracted from the mitochondrial membrane using the nonionic maltoside detergent. Recently, we developed an NMR sample of AAC reconstituted in DPC micelles to investigate the exchange dynamics of the transporter.17 As DPC as a detergent is stronger than DM, it is unclear whether its micelle system supports specific binding of CL. In this study, we performed thorough NMR measurements of CL−AAC interaction and molecular dynamics (MD) simulation to address the question posed above. Full-length yeast AAC carrier 3 (yAAC3) was expressed in Escherichia coli cells and purified by Ni-NTA affinity, ion exchange, and size-exclusion chromatography as previously described17 (Figure S1a). During NMR sample preparation,
ABSTRACT: Many membrane proteins bind specifically to lipids as an integral component of their structures. The ability of detergents to support lipid binding is thus an important consideration when solubilizing membrane proteins for structural studies. In particular, the zwitterionic phosphocholine (PC)-based detergents, which have been widely used in solution NMR studies of channels and transporters, are controversial because of their strong solubilization power and thus perceived as more denaturing than nonionic detergents such as the maltosides. Here, we investigate the ability of the mitochondrial ADP/ATP carrier (AAC) to specifically bind cardiolipin, a mitochondrial lipid important for the carrier function, in dodecylphosphocholine (DPC) micelles. We found that in DPC, the AAC specifically binds cardiolipin in a manner consistent with the bound cardiolipins found in the crystal structures of the AAC determined in n-decyl β-Dmaltoside. Our results suggest that PC detergent is compatible with specific lipid binding and that PC detergent mixed with the relevant lipid represents a viable solubilization system for NMR studies of membrane proteins.
T
he past two decades have seen an increasing frequency of application of solution NMR spectroscopy to de novo structural characterization of transmembrane (TM) and membrane-associated protein domains as well as intact fulllength integral membrane proteins.1−3 In addition to structure determination, solution NMR affords a convenient means of investigating ligand binding and conformational dynamics when relevant membrane protein sample systems are being developed.4,5 Unlike solid-state NMR, solution NMR requires membrane proteins to be solubilized in membrane-mimetic media, and these media are mostly detergent micelles,5−8 less commonly bicelles,9−11 and very rarely lipid nanodiscs.12 The use of PC detergents, however, has been controversial, as these detergents have in some cases performed poorly in preserving the native activities of membrane proteins, especially for G-protein-coupled receptors.13 While new media for solution NMR that more closely resemble the natural lipid bilayer are still being developed, a system more suitable than detergent alone consists of detergent micelles mixed with © XXXX American Chemical Society
Received: August 12, 2016 Revised: September 11, 2016
A
DOI: 10.1021/acs.biochem.6b00836 Biochemistry XXXX, XXX, XXX−XXX
Biochemistry
Rapid Report
Given the long acyl chains of CL, it is not surprising to see widespread chemical shift changes because the lipid partition in the DPC micelle could have indirectly induced chemical shift changes by altering the micelle properties. To test whether the chemical shift perturbations are specific, we performed the same titration using 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine (POPE), another lipid rich in the mitochondrial inner membrane.24 The results showed that at a 3:1 lipid:yAAC3 ratio, only CL induced obvious chemical shift perturbations while POPE had essentially no effect (Figure 1c). The combined titration results indicate that CL binds specifically to yAAC3. We next applied nuclear Overhauser enhancement (NOE) spectroscopy to directly identify the CL binding sites on yAAC3 using proteins that are 15N-labeled and perdeuterated such that the NOE between the protein backbone amide protons and CL protons could be measured unambiguously. DPCs were also perdeuterated to avoid resonance overlap with CL. A 3D 15N-separated NOESY spectrum was recorded with a NOE mixing time of 200 ms at 900 MHz (1H frequency), for assigning residue-specific CL NOE cross-peaks. Consistent with the NMR titration results, many residues showed distinct CL NOEs, including those of the headgroup (3.90 ppm), the methylene groups of the acyl chain (1.29, 1.40, and 1.61 ppm), and the terminal methyl group (0.84 ppm) (Figure 2a). In particular, NOEs to the methylene groups of the hydrocarbon chain are widespread becuse of the large number of these methylene protons in each CL molecule.
DPC was systematically reduced to reach the minimal concentration required to generate a good NMR spectrum (Figure S1b). We first employed the NMR titration approach to examine CL-induced chemical shift perturbation of AAC. For this purpose, we used cardiolipin with 18 carbons. High-resolution three-dimensional (3D) TROSY-HNCO spectra were recorded at CL:yAAC3 ratios of 0:1, 1:1, 2:1, 3:1, and 4:1. The application of the 3D experiment during NMR titration greatly improved the completeness of peak analysis of the otherwise highly crowded HSQC spectrum (Figure S2a). Addition of CL caused very substantial chemical shift perturbations; e.g., residue I25, L92, and A145 showed shifts of as much as 0.15 ppm (Figure S2b,c). Moreover, the titration curves showed that the majority of the specific changes nearly plateaued after the 3:1 CL:yAAC3 ratio (Figure 1a). The estimated apparent
Figure 1. Characterization of cardiolipin (CL) binding by NMR titration. (a) Example of residue-specific chemical shift changes upon CL titration as monitored in the 3D HNCO spectrum. (b) Mapping onto the crystal structure of AAC. Sphere colors indicate the following: gray for −0 < Δδ < 0.05, light blue for −0.05 < Δδ < 0.10, and blue for Δδ > 0.10. (c) Comparison between the chemical shift perturbations of AAC by CL (left) and POPE (right). Red and blue peaks are from protein (0.8 mM) in the absence and presence of 2.4 mM lipid, respectively. Figure 2. Definition of cardiolipin (CL) binding sites by NOE measurements. (a) Sample strips from the 3D 15N-edited NOESYTROSY spectrum (200 ms NOE mixing time) recorded using a sample containing 0.8 mM 15N- and 2H-labeled yAAC3 and 2.4 mM CL, showing protein NOEs to three regions of the lipid, including the headgroup (blue), the bulk acyl chain (orange), and the terminus of the acyl chain (red). (b) Mapping CL NOEs onto the yAAC3 crystal structure for backbone amides that could be unambiguously analyzed (indicated by spheres). Sphere colors indicate the following: gray for amides that showed no lipid NOEs and other colors for amides that showed NOEs to lipid regions defined in panel a.
dissociation constants (KD) for A91, A148, and E268 in Figure 1a are approximately 100, 100, and 20 μM, respectively (Figure S2d). The AAC architecture resembles an open-top barrel formed by three structurally similar domains (I−III) in a parallel orientation. Each domain consists of two TM helices separated by an amphipathic helix. Overall, the chemical shift perturbation is widespread on the periphery of the carrier (Figure 1b), and the titration data suggest a 3:1 CL:yAAC3 binding stoichiometry. B
DOI: 10.1021/acs.biochem.6b00836 Biochemistry XXXX, XXX, XXX−XXX
Biochemistry
Rapid Report
On the basis of the observed lipid NOE cross-peaks, the protein residues were assigned to be in close contact with each of the three regions of CL shown in Figure 2a, including (1) the headgroup region (cross-peak 1 only), (2) the core regions of the acyl chains (any combination of cross-peak 2, 3, or 4), and (3) the terminal regions of the acyl chain (cross-peaks 4 and 5). The residues in contact with the headgroup are located mainly on the matrix side of the carrier in and near the amphipathic helices (h12, h34, and h56); they are mostly charged or polar residues such as R141, Y179, and K275 in regions I, II, and III, respectively (Figure 2b and Figure S3). Most of the peripheral residues along the six TM helices show NOE cross-peaks to the acyl chain protons. As expected, among these TM residues, those close to the matrix side showed both methylene and headgroup NOEs (e.g., W76, F182, and A277) and several residues near the cytosol side showed methylene and terminal methyl group NOEs (e.g., I14, I100, and S292). Finally, the middle sections of the TM helices showed mostly methylene NOEs. The majority of the TM residues in H4 of domain II were not assigned (Figure S2a), and thus, the NOE data are not available for this region of the protein. The NOE data are overall consistent with CL positions observed in the crystal structures. The yAAC3 crystal structure determined in DM showed electron densities corresponding to the headgroups of tetramyristoyl (C14) CL, one for each of the three domains.21 The locations of these densities near the amphipathic helices, where several lysines and arginines reside, are consistent with our NOE data (Figure 2b) and also consistent with the hypothesis that electrostatic interactions between the negatively charged CL headgroup and basic residues on the matrix side of the carrier mediate CL recognition.25 In the yAAC3 crystal structure, however, most of the electron density for the CL acyl chains is missing. We thus performed MD simulation to gain insights into the position and orientation of the acyl chains. The simulation system consists of yAAC3 and three CLs embedded in the POPC lipid bilayer (Methods in the Supporting Information). The CLs were initially positioned such that their headgroups coincide with those in the crystal structure. Two MD stages were used for this system. The first stage was restrained MD in which the protein backbone and the CL headgroup atoms were restricted with a force constant of 100 kJ mol−1 Å−2 whereas the protein side chains and CL acyl chains were allowed to move. Upon reaching equilibrium, the system was subjected to the second stage, unrestrained simulation (40 ns). The simulation and NOE data are overall in agreement (Figure 3a); i.e., during the simulation, the distances between the acyl chain methyl carbons and protein backbone nitrogens (4.5−7.0 Å) were on average consistent with the corresponding NOE-derived distances between the protons (3−5 Å) (Figure S4). The combined NMR and MD results indicate that CL acyl chains interact with the hydrophobic residues along the six TM helices. Because CL has four acyl chains, one CL is sufficient for covering the two TM helices of each domain, and thus three CLs are sufficient for wrapping the entire carrier. It is interesting to note that hydrophobic interaction and geometry matching are the key factors governing acyl chain placement. In particular, for CLs in regions I and II, one of the acyl chains has its terminal methyl group anchored to a hydrophobic pocket in the middle of the TM helices, which provided explanation for the rather unexpected methyl NOEs observed in these regions of the protein (Figure S5).
Figure 3. Positions of CLs (C18) bound to yAAC3 after MD simulation for 40 ns in the POPC bilayer. The blue and red spheres indicate analyzed residues that showed NOEs to the CL headgroup and terminal methyl group, respectively. Key NOEs to the CL terminal methyl group are marked by a black dashed arrow. Simulation details are described in the text and Supporting Information.
In summary, we have shown that in DPC micelles, AAC is able to interact specifically with CL. All three structurally similar domains of AAC interact with a CL molecule, while domain I (H1 and H2) appeared to show the strongest contact based on NOE analysis. The CL binding is not a nonspecific effect of hydrophobic partitioning in micelles, as the POPE phospholipid induced far less chemical shift perturbation on AAC than CL did at the same detergent and lipid concentrations. Another important observation is that AAC can bind CL in similar manner in two completely different environments, DPC versus DM and solution versus crystal. Given the high abundance of CL in the inner mitochondrial membrane (∼17%),24 our results strongly suggest that AAC exists in complex with CL in the native environment, though the precise role, structural or functional, of this lipid in AAC remains to be investigated. The combined NMR/MD experiments also provided the structural information about interaction between the CL acyl chain and yAAC3 that was largely missing in the crystal structure. Finally, our study provides a solid example in which detergent micelles formed with a PC detergent are compatible with specific lipid binding by a membrane protein, while suggesting the use of a DPC/ lipid mixed micelle as a viable solution to NMR studies of membrane proteins.
■
ASSOCIATED CONTENT
S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.biochem.6b00836. Detailed text description and figures relating to sample preparation (Figure S1), NMR data (Figure S2), MD simulation (Figure S4), and structural details of protein− lipid interaction (Figures S3 and S4) (PDF)
■
AUTHOR INFORMATION
Corresponding Authors
*E-mail:
[email protected]. *E-mail:
[email protected]. C
DOI: 10.1021/acs.biochem.6b00836 Biochemistry XXXX, XXX, XXX−XXX
Biochemistry
Rapid Report
Author Contributions ⊥
L.Z. and S.W. contributed equally to this work. L.Z. and J.J.C. conceived the study. L.Z. and C.R. prepared the samples and collected NMR data. L.Z. and B.O. analyzed NMR data. S.W. performed MD analysis. J.J.C. and L.Z. wrote the paper, and all authors contributed to editing of the manuscript. Funding
This work was supported by CAS Grant XDB08030301 and National Natural Science Foundation of China grant no. 31570746 to J.J.C. Notes
The authors declare no competing financial interest.
■ ■
ACKNOWLEDGMENTS We thank Zhijun Liu and Bin Wu from the NMR facility of NCPSS for their assistance with NMR data collection. REFERENCES
(1) Oxenoid, K., and Chou, J. J. (2013) Curr. Opin. Struct. Biol. 23, 547. (2) Kim, H. J., Howell, S. C., Van Horn, W. D., Jeon, Y. H., and Sanders, C. R. (2009) Prog. Nucl. Magn. Reson. Spectrosc. 55, 335. (3) Kang, C., and Li, Q. (2011) Curr. Opin. Chem. Biol. 15, 560. (4) Oxenoid, K., and Chou, J. J. (2016) Protein Sci. 25, 959. (5) Jaremko, L., Jaremko, M., Giller, K., Becker, S., and Zweckstetter, M. (2014) Science 343, 1363. (6) Schnell, J. R., and Chou, J. J. (2008) Nature 451, 591. (7) Oxenoid, K., Dong, Y., Cao, C., Cui, T., Sancak, Y., Markhard, A. L., Grabarek, Z., Kong, L., Liu, Z., Ouyang, B., Cong, Y., Mootha, V. K., and Chou, J. J. (2016) Nature 533, 269. (8) Hiller, S., Garces, R. G., Malia, T. J., Orekhov, V. Y., Colombini, M., and Wagner, G. (2008) Science 321, 1206. (9) Fu, Q., Fu, T. M., Cruz, A. C., Sengupta, P., Thomas, S. K., Wang, S., Siegel, R. M., Wu, H., and Chou, J. J. (2016) Mol. Cell 61, 602. (10) Morrison, E. A., DeKoster, G. T., Dutta, S., Vafabakhsh, R., Clarkson, M. W., Bahl, A., Kern, D., Ha, T., and Henzler-Wildman, K. A. (2011) Nature 481, 45. (11) Lau, T. L., Kim, C., Ginsberg, M. H., and Ulmer, T. S. (2009) EMBO J. 28, 1351. (12) Hagn, F., Etzkorn, M., Raschle, T., and Wagner, G. (2013) J. Am. Chem. Soc. 135, 1919. (13) Serrano-Vega, M. J., Magnani, F., Shibata, Y., and Tate, C. G. (2008) Proc. Natl. Acad. Sci. U. S. A. 105, 877. (14) Seddon, A. M., Curnow, P., and Booth, P. J. (2004) Biochim. Biophys. Acta, Biomembr. 1666, 105. (15) Yeagle, P. L. (2014) Biochim. Biophys. Acta, Biomembr. 1838, 1548. (16) Claypool, S. M., Oktay, Y., Boontheung, P., Loo, J. A., and Koehler, C. M. (2008) J. Cell Biol. 182, 937. (17) Bruschweiler, S., Yang, Q., Run, C., and Chou, J. J. (2015) Nat. Struct. Mol. Biol. 22, 636. (18) Klingenberg, M. (2008) Biochim. Biophys. Acta, Biomembr. 1778, 1978. (19) Kunji, E. R., and Robinson, A. J. (2010) Curr. Opin. Struct. Biol. 20, 440. (20) Pebay-Peyroula, E., Dahout-Gonzalez, C., Kahn, R., Trezeguet, V., Lauquin, G. J., and Brandolin, G. (2003) Nature 426, 39. (21) Ruprecht, J. J., Hellawell, A. M., Harding, M., Crichton, P. G., McCoy, A. J., and Kunji, E. R. (2014) Proc. Natl. Acad. Sci. U. S. A. 111, E426. (22) Heimpel, S., Basset, G., Odoy, S., and Klingenberg, M. (2001) J. Biol. Chem. 276, 11499. (23) Hoffmann, B., Stockl, A., Schlame, M., Beyer, K., and Klingenberg, M. (1994) J. Biol. Chem. 269, 1940. (24) van Meer, G., Voelker, D. R., and Feigenson, G. W. (2008) Nat. Rev. Mol. Cell Biol. 9, 112. (25) Beyer, K., and Klingenberg, M. (1985) Biochemistry 24, 3821. D
DOI: 10.1021/acs.biochem.6b00836 Biochemistry XXXX, XXX, XXX−XXX
