Skip to main content

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

  • Letter
  • Published:

The role of interfacial lipids in stabilizing membrane protein oligomers

Nature volume 541, pages 421–424 (2017)Cite this article

Subjects

Abstract

Oligomerization of membrane proteins in response to lipid binding has a critical role in many cell-signalling pathways1 but is often difficult to define2 or predict3. Here we report the development of a mass spectrometry platform to determine simultaneously the presence of interfacial lipids and oligomeric stability and to uncover how lipids act as key regulators of membrane-protein association. Evaluation of oligomeric strength for a dataset of 125 α-helical oligomeric membrane proteins reveals an absence of interfacial lipids in the mass spectra of 12 membrane proteins with high oligomeric stability. For the bacterial homologue of the eukaryotic biogenic transporters (LeuT4, one of the proteins with the lowest oligomeric stability), we found a precise cohort of lipids within the dimer interface. Delipidation, mutation of lipid-binding sites or expression in cardiolipin-deficient Escherichia coli abrogated dimer formation. Molecular dynamics simulation revealed that cardiolipin acts as a bidentate ligand, bridging across subunits. Subsequently, we show that for the Vibrio splendidus sugar transporter SemiSWEET5, another protein with low oligomeric stability, cardiolipin shifts the equilibrium from monomer to functional dimer. We hypothesized that lipids are essential for dimerization of the Na+/H+ antiporter NhaA from E. coli, which has the lowest oligomeric strength, but not for the substantially more stable homologous Thermus thermophilus protein NapA. We found that lipid binding is obligatory for dimerization of NhaA, whereas NapA has adapted to form an interface that is stable without lipids. Overall, by correlating interfacial strength with the presence of interfacial lipids, we provide a rationale for understanding the role of lipids in both transient and stable interactions within a range of α-helical membrane proteins, including G-protein-coupled receptors.

This is a preview of subscription content, access via your institution

Access options

Buy this article

  • Purchase on Springer Link
  • Instant access to full article PDF
Buy now

Prices may be subject to local taxes which are calculated during checkout

Figure 1: Plot of buried surface area and number of salt bridges for oligomeric α-helical membrane proteins and native mass spectra.
Figure 2: Schematic of the high-energy MS/MS platform, mass spectra and molecular dynamics simulations of the lipid-bound LeuT dimer.
Figure 3: Mass spectrum recorded for SemiSWEET and the effect of cardiolipin on the monomer–dimer equilibrium.
Figure 4: Comparison of mass spectra of NhaA with NapA and plot of transporter stabilities with assorted G-protein-coupled receptors.

Similar content being viewed by others

References

  1. Arkhipov, A. et al. Architecture and membrane interactions of the EGF receptor. Cell 152, 557–569 (2013)

    Article  CAS  Google Scholar 

  2. Wu, H. et al. Structure of a class C GPCR metabotropic glutamate receptor 1 bound to an allosteric modulator. Science 344, 58–64 (2014)

    Article  ADS  CAS  Google Scholar 

  3. Duarte, J. M., Biyani, N., Baskaran, K. & Capitani, G. An analysis of oligomerization interfaces in transmembrane proteins. BMC Struct. Biol. 13, 21 (2013)

    Article  CAS  Google Scholar 

  4. Piscitelli, C. L., Krishnamurthy, H. & Gouaux, E. Neurotransmitter/sodium symporter orthologue LeuT has a single high-affinity substrate site. Nature 468, 1129–1132 (2010)

    Article  ADS  CAS  Google Scholar 

  5. Xu, Y. et al. Structures of bacterial homologues of SWEET transporters in two distinct conformations. Nature 515, 448–452 (2014)

    Article  ADS  CAS  Google Scholar 

  6. Yeagle, P. L. Non-covalent binding of membrane lipids to membrane proteins. Biochim. Biophys. Acta 1838, 1548–1559 (2014)

    Article  CAS  Google Scholar 

  7. Hansen, S. B., Tao, X. & MacKinnon, R. Structural basis of PIP2 activation of the classical inward rectifier K+ channel Kir2.2. Nature 477, 495–498 (2011)

    Article  ADS  CAS  Google Scholar 

  8. Gao, Y., Cao, E., Julius, D. & Cheng, Y. TRPV1 structures in nanodiscs reveal mechanisms of ligand and lipid action. Nature 534, 347–351 (2016)

    Article  ADS  CAS  Google Scholar 

  9. Whitelegge, J. P. Integral membrane proteins and bilayer proteomics. Anal. Chem. 85, 2558–2568 (2013)

    Article  CAS  Google Scholar 

  10. Savas, J. N., Stein, B. D., Wu, C. C. & Yates, J. R., III . Mass spectrometry accelerates membrane protein analysis. Trends Biochem. Sci. 36, 388–396 (2011)

    CAS  PubMed  PubMed Central  Google Scholar 

  11. Konijnenberg, A., van Dyck, J. F., Kailing, L. L. & Sobott, F. Extending native mass spectrometry approaches to integral membrane proteins. Biol. Chem. 396, 991–1002 (2015)

    Article  CAS  Google Scholar 

  12. Laganowsky, A. et al. Membrane proteins bind lipids selectively to modulate their structure and function. Nature 510, 172–175 (2014)

    Article  ADS  CAS  Google Scholar 

  13. Landreh, M., Marty, M. T., Gault, J. & Robinson, C. V. A sliding selectivity scale for lipid binding to membrane proteins. Curr. Opin. Struct. Biol. 39, 54–60 (2016)

    Article  CAS  Google Scholar 

  14. Levy, E. D., Boeri Erba, E., Robinson, C. V. & Teichmann, S. A. Assembly reflects evolution of protein complexes. Nature 453, 1262–1265 (2008)

    Article  ADS  CAS  Google Scholar 

  15. Pliotas, C. et al. The role of lipids in mechanosensation. Nat. Struct. Mol. Biol. 22, 991–998 (2015)

    Article  CAS  Google Scholar 

  16. Ilgü, H. et al. Variation of the detergent-binding capacity and phospholipid content of membrane proteins when purified in different detergents. Biophys. J. 106, 1660–1670 (2014)

    Article  ADS  Google Scholar 

  17. Brügger, B. Lipidomics: analysis of the lipid composition of cells and subcellular organelles by electrospray ionization mass spectrometry. Annu. Rev. Biochem. 83, 79–98 (2014)

    Article  Google Scholar 

  18. Taki, T. TLC-blot (far-eastern blot) and its application to functional lipidomics. Methods Mol. Biol. 1314, 219–241 (2015)

    Article  Google Scholar 

  19. Cox, J. & Mann, M. Quantitative, high-resolution proteomics for data-driven systems biology. Annu. Rev. Biochem. 80, 273–299 (2011)

    Article  CAS  Google Scholar 

  20. Barrera, N. P., Di Bartolo, N., Booth, P. J. & Robinson, C. V. Micelles protect membrane complexes from solution to vacuum. Science 321, 243–246 (2008)

    Article  ADS  CAS  Google Scholar 

  21. Audet, M. & Bouvier, M. Restructuring G-protein-coupled receptor activation. Cell 151, 14–23 (2012)

    Article  CAS  Google Scholar 

  22. Huang, W. et al. Structural insights into μ-opioid receptor activation. Nature 524, 315–321 (2015)

    Article  ADS  CAS  Google Scholar 

  23. Sounier, R. et al. Propagation of conformational changes during μ-opioid receptor activation. Nature 524, 375–378 (2015)

    Article  ADS  CAS  Google Scholar 

  24. Dorsch, S., Klotz, K. N., Engelhardt, S., Lohse, M. J. & Bünemann, M. Analysis of receptor oligomerization by FRAP microscopy. Nat. Methods 6, 225–230 (2009)

    Article  CAS  Google Scholar 

  25. Drew, D. & Boudker, O. Shared molecular mechanisms of membrane transporters. Annu. Rev. Biochem. 85, 543–572 (2016)

    Article  CAS  Google Scholar 

  26. Herz, K., Rimon, A., Jeschke, G. & Padan, E. β-sheet-dependent dimerization is essential for the stability of NhaA Na+/H+ antiporter. J. Biol. Chem. 284, 6337–6347 (2009)

    Article  CAS  Google Scholar 

  27. Anderluh, A. et al. Single molecule analysis reveals coexistence of stable serotonin transporter monomers and oligomers in the live cell plasma membrane. J. Biol. Chem. 289, 4387–4394 (2014)

    Article  CAS  Google Scholar 

  28. Zhen, J. et al. Dopamine transporter oligomerization: impact of combining protomers with differential cocaine analog binding affinities. J. Neurochem. 133, 167–173 (2015)

    Article  CAS  Google Scholar 

  29. Coleman, J. A., Green, E. M. & Gouaux, E. X-ray structures and mechanism of the human serotonin transporter. Nature 532, 334–339 (2016)

    Article  ADS  CAS  Google Scholar 

  30. Kilic, F. & Rudnick, G. Oligomerization of serotonin transporter and its functional consequences. Proc. Natl Acad. Sci. USA 97, 3106–3111 (2000)

    Article  ADS  CAS  Google Scholar 

  31. Tan, B. K. et al. Discovery of a cardiolipin synthase utilizing phosphatidylethanolamine and phosphatidylglycerol as substrates. Proc. Natl Acad. Sci. USA 109, 16504–16509 (2012)

    Article  ADS  CAS  Google Scholar 

  32. Yamashita, A., Singh, S. K., Kawate, T., Jin, Y. & Gouaux, E. Crystal structure of a bacterial homologue of Na+/Cl−-dependent neurotransmitter transporters. Nature 437, 215–223 (2005)

    Article  ADS  CAS  Google Scholar 

  33. Coincon, M. et al. Crystal structures reveal the molecular basis of ion translocation in sodium/proton antiporters. Nat. Struct. Mol. Biol. 23, 248–255 (2016)

    Article  CAS  Google Scholar 

  34. Lee, C. et al. Crystal structure of the sodium–proton antiporter NhaA dimer and new mechanistic insights. J. Gen. Physiol. 144, 529–544 (2014)

    Article  CAS  Google Scholar 

  35. Laganowsky, A., Reading, E., Hopper, J. T. & Robinson, C. V. Mass spectrometry of intact membrane protein complexes. Nat. Protocols 8, 639–651 (2013)

    Article  CAS  Google Scholar 

  36. Hernández, H. & Robinson, C. V. Determining the stoichiometry and interactions of macromolecular assemblies from mass spectrometry. Nat. Protocols 2, 715–726 (2007)

    Article  Google Scholar 

  37. Marty, M. T. et al. Bayesian deconvolution of mass and ion mobility spectra: from binary interactions to polydisperse ensembles. Anal. Chem. 87, 4370–4376 (2015)

    Article  CAS  Google Scholar 

  38. Gault, J. et al. High-resolution mass spectrometry of small molecules bound to membrane proteins. Nat. Methods 13, 333–336 (2016)

    Article  Google Scholar 

  39. Abraham, M. J. et al. GROMACS: High performance molecular simulations through multi-level parallelism from laptops to supercomputers. SoftwareX 1–2, 19–25 (2015)

    Article  ADS  Google Scholar 

  40. Stansfeld, P. J. et al. MemProtMD: automated insertion of membrane protein structures into explicit lipid membranes. Structure 23, 1350–1361 (2015)

    Article  CAS  Google Scholar 

  41. de Jong, D. H. et al. Improved parameters for the martini coarse-grained protein force field. J. Chem. Theory Comput. 9, 687–697 (2013)

    Article  CAS  Google Scholar 

  42. Bussi, G., Donadio, D. & Parrinello, M. Canonical sampling through velocity rescaling. J. Chem. Phys. 126, 014101 (2007)

    Article  ADS  Google Scholar 

  43. Berendsen, H. J. C., Postma, J. P. M., Vangunsteren, W. F., Dinola, A. & Haak, J. R. Molecular dynamics with coupling to an external bath. J. Chem. Phys. 81, 3684–3690 (1984)

    Article  ADS  CAS  Google Scholar 

  44. Hess, B. P-LINCS: A parallel linear constraint solver for molecular simulation. J. Chem. Theory Comput. 4, 116–122 (2008)

    Article  CAS  Google Scholar 

  45. Tironi, I. G., Sperb, R., Smith, P. E. & Vangunsteren, W. F. A generalized reaction field method for molecular dynamics simulations. J. Chem. Phys. 102, 5451–5459 (1995)

    Article  ADS  CAS  Google Scholar 

  46. Pall, S. & Hess, B. A flexible algorithm for calculating pair interactions on SIMD architectures. Comput. Phys. Commun. 184, 2641–2650 (2013)

    Article  ADS  CAS  Google Scholar 

  47. Stansfeld, P. J. & Sansom, M. S. Molecular simulation approaches to membrane proteins. Structure 19, 1562–1572 (2011)

    Article  CAS  Google Scholar 

  48. Jefferys, E., Sands, Z. A., Shi, J., Sansom, M. S. & Fowler, P. W. Alchembed: a computational method for incorporating multiple proteins into complex lipid geometries. J. Chem. Theory Comput. 11, 2743–2754 (2015)

    Article  CAS  Google Scholar 

  49. Michaud-Agrawal, N., Denning, E. J., Woolf, T. B. & Beckstein, O. MDAnalysis: a toolkit for the analysis of molecular dynamics simulations. J. Comput. Chem. 32, 2319–2327 (2011)

    Article  CAS  Google Scholar 

  50. DeLano, W. L. The PyMOL Molecular Graphics System. (2002) http://www.pymol.org/

  51. Krissinel, E. & Henrick, K. Inference of macromolecular assemblies from crystalline state. J. Mol. Biol. 372, 774–797 (2007)

    Article  CAS  Google Scholar 

  52. Sturt, H. F., Summons, R. E., Smith, K., Elvert, M. & Hinrichs, K. U. Intact polar membrane lipids in prokaryotes and sediments deciphered by high-performance liquid chromatography/electrospray ionization multistage mass spectrometry–new biomarkers for biogeochemistry and microbial ecology. Rapid Commun. Mass Spectrom. 18, 617–628 (2004)

    Article  ADS  CAS  Google Scholar 

  53. Hopper, J. T. et al. Detergent-free mass spectrometry of membrane protein complexes. Nat. Methods 10, 1206–1208 (2013)

    Article  CAS  Google Scholar 

  54. Barrera, N. P. et al. Mass spectrometry of membrane transporters reveals subunit stoichiometry and interactions. Nat. Methods 6, 585–587 (2009)

    Article  CAS  Google Scholar 

  55. Wang, S. C. et al. Ion mobility mass spectrometry of two tetrameric membrane protein complexes reveals compact structures and differences in stability and packing. J. Am. Chem. Soc. 132, 15468–15470 (2010)

    Article  CAS  Google Scholar 

  56. Reading, E. et al. The role of the detergent micelle in preserving the structure of membrane proteins in the gas phase. Angew. Chem. Int. Edn Engl. 54, 4577–4581 (2015)

    Article  CAS  Google Scholar 

Download references

Acknowledgements

We thank K. Giles (Waters Corporation) and J. Benesch for development of the high-energy source and T. Allison, M. Degiacomi and J. Gault for many helpful discussions. The Robinson group is funded by a Wellcome Trust Investigator Award (104633/Z/14/Z), an ERC Advanced Grant ENABLE (641317) and an MRC programme grant (MR/N020413/1). K.G. is a research fellow of the Royal Commission for the Exhibition of 1851 and a Junior Research Fellow at St Catherine’s College, Oxford. J.A.C.D. is supported by an EPSRC studentship, held at the Life Sciences Interface Doctoral Training Centre. M.L. holds an ERC Marie Curie Career Development Fellowship and is a Junior Research Fellow at St Cross College, Oxford. D.D. acknowledges support from the EMBO Young Investigator Program, Vetenskapsrådet and the Knut and Alice Wallenberg foundation. A.J.B. acknowledges a BBSRC David Phillip’s Fellowship, BB/J014346/1. The authors are also grateful for plasmids from E. Gouaux (LeuT), W. Frommer and L. Feng (SemiSWEET).

Author information

Author notes

  1. Kallol Gupta, Joseph A. C. Donlan and Jonathan T. S. Hopper: These authors contributed equally to this work.

Authors and Affiliations

  1. Department of Chemistry, University of Oxford, 12 Mansfield Road, Oxford, OX1 3TA, UK

    Kallol Gupta, Joseph A. C. Donlan, Jonathan T. S. Hopper, Michael Landreh, Weston B. Struwe, Andrew J. Baldwin & Carol V. Robinson

  2. Department of Biochemistry and Biophysics, Centre for Biomembrane Research, Stockholm University, Stockholm, SE-106 91, Sweden

    Povilas Uzdavinys & David Drew

  3. Department of Biochemistry, University of Oxford, South Parks Road, Oxford, OX1 3QU, UK

    Phillip J. Stansfeld

Authors
  1. Kallol Gupta
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. Joseph A. C. Donlan
    View author publications

    You can also search for this author in PubMed Google Scholar

  3. Jonathan T. S. Hopper
    View author publications

    You can also search for this author in PubMed Google Scholar

  4. Povilas Uzdavinys
    View author publications

    You can also search for this author in PubMed Google Scholar

  5. Michael Landreh
    View author publications

    You can also search for this author in PubMed Google Scholar

  6. Weston B. Struwe
    View author publications

    You can also search for this author in PubMed Google Scholar

  7. David Drew
    View author publications

    You can also search for this author in PubMed Google Scholar

  8. Andrew J. Baldwin
    View author publications

    You can also search for this author in PubMed Google Scholar

  9. Phillip J. Stansfeld
    View author publications

    You can also search for this author in PubMed Google Scholar

  10. Carol V. Robinson
    View author publications

    You can also search for this author in PubMed Google Scholar

Contributions

K.G. and C.V.R. designed the experiments. K.G. and J.A.C.D. performed protein expression and MS experiments. J.T.S.H. performed the high-energy experiments with K.G. K.G. and M.L. performed MS experiments on NapA and NhaA. P.U. expressed and purified NhaA and NapA under the guidance of D.D. J.A.C.D. purified SemiSWEET with the help of W.B.S. P.J.S. carried out molecular dynamics simulations. A.J.B. and K.G. performed theoretical calculations to determine the oligomeric strength. K.G. and C.V.R wrote the manuscript with contributions from all authors.

Corresponding author

Correspondence to Carol V. Robinson.

Ethics declarations

Competing interests

The authors declare no competing financial interests.

Additional information

Reviewer Information Nature thanks S. Bernèche, A. Lee and J. Whitelegge for their contribution to the peer review of this work.

Extended data figures and tables

Extended Data Figure 1 Mass spectra of LeuT recorded with increasing collision voltages and of a LeuT fusion protein construct.

a, Mass spectra of LeuT, liberated from octylglucoside micelles, (green/grey spheres, most abundant charge state highlighted in pale blue), show that the 7.4-kDa lipid adduct (blue/purple head groups) is retained throughout the trap collision energy range (white, blue arrow) of the mass spectrometer. b, Mass spectra of LeuT expressed as a fusion protein with eYFP (LeuT–eYFP yellow circles), liberated from octylglucoside micelles, show that the dimer is similarly associated with a 7.4-kDa adduct.

Extended Data Figure 2 Mass spectra of LeuT following incubation with delipidating detergents and E. coli polar lipids.

a, Mass spectrum of LeuT liberated from octylglucoside micelles (green head groups) shows low-abundance delipidated monomers (green spheres, 59.3 kDa) and high-abundance lipid-bound dimers (green/black spheres, 126.0 kDa). b, Mass spectrum of LeuT after incubation with neopentyl glycol (NG, orange head-groups) shows only delipidated monomers. c, Mass spectrum of LeuT in octylglucoside (OG), after incubation with neopentyl glycol, shows only delipidated monomers. d, Mass spectrum recorded after incubation of delipidated LeuT monomers, in octylglucoside, with E. coli polar lipids (blue/purple head-groups) shows delipidated monomers and lipid-bound dimers. e, Mass spectrum recorded after adding dilysocardiolipin (blue head-groups) to delipidated monomeric LeuT in octylglucoside (c) shows no dimerization in the presence of this lipid.

Extended Data Figure 3 High-energy MS/MS experiment of the 23+ charge state of dimeric LeuT, with the 7.4-kDa adduct, as a function of collision voltage.

Three satellite peaks represent the lipid-bound states arising through the dissociation of the monomer. The naked monomer is highlighted in blue, while the three satellite peaks are assigned to one phospholipid, one cardiolipin and three phospholipid-bound species (red, green and yellow, respectively). Under higher energy, only the cardiolipin-bound species remains, discounting the mathematical possibility of two phospholipid-bound species. Inset shows the isolated 23+ charge state of the lipid bound dimer. Presence of bound cardiolipin at a higher energy, over that of phospholipid, indicates a higher binding energy of cardiolipin over the latter, potentially owing to greater ionic and hydrophobic interactions.

Extended Data Figure 4 Site-directed mutagenesis of selected residues at the LeuT dimer interface, resulting mass spectra and molecular dynamics simulations.

a, Mass spectrum of LeuT F488A/Y489A, liberated from octylglucoside micelles, reveals monomeric LeuT (green spheres). Inset shows the LeuT dimer interface, with key π-stacking interactions (yellow dotted lines, distances labelled in red) and between aromatic residues (purple). When residues F488 and Y489 (orange arrows) are mutated to alanine, the π-stacking interactions are abolished and LeuT cannot dimerize. b, Molecular dynamics simulations of LeuT in an E. coli lipid bilayer reveal possible binding sites of interfacial phospholipids and cardiolipin (upper panel, viewed from cytoplasmic side of membrane). The cardiolipin phosphate groups (orange) interact closely with positively charged residues (K376, H377, R506; blue) at the dimer interface. Phosphoethanolamine (PE) and phosphatidylglycerol (PG) also bind at the dimer interface. c, Mass spectrum of LeuT expressed in a cardiolipin-deficient E. coli strain (BKT22), liberated from octylglucoside micelles, shows monomeric LeuT, implying that cardiolipin is required for LeuT dimerization. d, Mass spectrum of LeuT K376A/H377A, liberated from octylglucoside micelles, shows monomeric LeuT.

Extended Data Figure 5 CGMD simulations on LeuT and NhaA dimer.

a, Particle densities from five repeats of 1-μs CGMD simulations for cardiolipin around LeuT. The surface densities represent the most occupied positions from the simulations of the phosphate (orange), glycerol (red) and alkyl tails (purple) particles of cardiolipin. The proposed binding sites at the interface are the only places where cardiolipin shows considerable population density. b–d, Comparative particle densities of cardiolipin (b), phosphatidylglycerol (c) and phosphoethanolamine (d) at the LeuT dimeric interface, summed over the simulations show no or minimal densities for phosphatidylglycerol and phosphoethanolamine at the cardiolipin-binding site. Together, a–d show that the proposed binding sites of cardiolipin at the interface are sites of specific bindings. e, Dimeric structure of LeuT with modelled APT (aminopentanetetrol, aminophospholipids) classes of lipid present in A. aeolicus52. The lipid was drawn in ChemDraw and subsequently modelled by superimposition onto cardiolipin to give the cardiolipin-bound dimeric structure. The favourable van der Waals distances show that it is capable of bridging the dimeric entity through the same sets of residues that were found to be critical towards cardiolipin binding, in an endogenous environment lacking cardiolipins. f, Particle densities from five repeats of 1-μs CGMD simulations for cardiolipin (phosphate group in orange, glycerol in red and alkyl tails in purple) and POPG (in blue) around NhaA dimer interface. As before, the density of cardiolipin is considerably higher than that of phosphatidylglycerol. However, unlike LeuT, here the difference between the density of cardiolipin and phosphatidylglycerol is lower, suggesting this site has less exclusivity towards cardiolipin than that in LeuT. Indeed, mass spectrometry analysis shows a heterogenous distribution of lipids with dimeric NhaA, with mostly cardiolipin but some amount of bound phospholipids.

Extended Data Figure 6 Mass spectra of His-tagged and unmodified SemiSWEET and identification of endogenous and exogenous lipid binding.

a, Mass spectrum of unmodified SemiSWEET, liberated from tetraethyleneglycolmonooctyl ether (C8E4) micelles, reveals SemiSWEET monomers and dimers (black spheres). b, Mass spectrum of deca-His tagged SemiSWEET, liberated from C8E4 micelles, reveals SemiSWEET monomers and dimers (green spheres). c, High energy MS/MS of unmodified SemiSWEET, liberated from dodecylmaltoside (DDM) micelles, allows isolation of the 6+ charge state (black spheres) of the SemiSWEET monomer (black spheres) bound to endogenous lipids. Fragmentation of the lipid-bound species leads to loss of either cardiolipin (1,470 ± 26 Da, purple head-groups), one or two neutral phospholipids (each 756 ± 22 Da, blue head-groups), or a positively charged phospholipid. Trap collision voltages are shown in white inside the blue arrow. d, Mass spectrum of deca-His SemiSWEET, liberated from C8E4 micelles and incubated with phosphatidylglycerol (blue head-groups). phosphatidylglycerol binds to both monomers and dimers (dotted boxes highlight lipid-bound peaks) without substantial preference. e, Mass spectrum recorded after incubation in solution of an equimolar ratio of deca-His tagged and untagged SemiSWEET (green and black spheres, respectively), liberated from tetraethyleneglycolmonooctyl ether (C8E4) micelles. Plot of the percentage abundance of hetero- and homodimers over time (inset), SemiSWEET heterodimers (red trace, peaks highlighted red in mass spectrum) and homodimers (black trace), revealing the solution-phase monomer–dimer equilibrium (PDB accession number: 4QND).

Extended Data Figure 7 Mass spectrum and high-energy MS/MS of NhaA at a range of collision voltages.

a, Mass spectrum of NhaA, liberated from C8E4 micelles, reveals NhaA monomers (green spheres) bound to cardiolipin (purple head-groups) and an ensemble of NhaA dimer species in different lipidation states (highlighted in green). b, MS/MS of the 15+ charge state (green) of the NhaA dimer (green/black spheres) bound to two cardiolipin molecules liberated from C8E4 micelles. Increasing collision voltage applied to the 2× cardiolipin-bound species leads either to loss of 1 cardiolipin to form NhaA dimers bound to 1 cardiolipin (40 V) or to loss of 2 cardiolipin molecules to form delipidated NhaA dimers, with concomitant generation of NhaA monomers (70 V) and further dissociation of NhaA dimers into monomers (120 V). Trap collision voltages are depicted in white, inside the blue arrow.

Extended Data Figure 8 Sequence and structure alignment of LeuT with other eukaryotic biogenic transporters.

a, The basic residues of LeuT that are involved in lipid binding (red box) are conserved across the BATs. b, Two views of the superimposed structures of LeuT (PDB accession number: 2A65, black) and SERT (PDB accession number: 5I6Z, light blue) show the differences in the dimer interface. Dimer interface helices are highlighted with arrows and coloured (LeuT, green; SERT, red); basic residues responsible for lipid binding in LeuT are shown in yellow mesh. One of the interface helices in SERT swings away from the interface, negating the possibility of lipid-induced oligomerization, analogous to that proposed for LeuT.

Extended Data Table 1 Summary of the mass spectral analysis of membrane proteins forming strong oligomers
Full size table

Supplementary information

Supplementary Table 1

Total buried surface area and the number salt bridges for each of the oligomeric proteins. The respective PDB IDs are also provided (XLSX 22 kb)

PowerPoint slides

Rights and permissions

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Gupta, K., Donlan, J., Hopper, J. et al. The role of interfacial lipids in stabilizing membrane protein oligomers. Nature 541, 421–424 (2017). https://doi.org/10.1038/nature20820

Download citation

  • Received:

  • Accepted:

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/nature20820

This article is cited by

Comments

By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.

Search

Advanced search

Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing