Abstract
Recent applications of mass spectrometry (MS) to study membrane protein complexes are yielding valuable insights into the binding of lipids and their structural and functional roles. To date, most native MS experiments with membrane proteins are based on detergent solubilization. Many insights into the structure and function of membrane proteins have been obtained using detergents; however, these can promote local lipid rearrangement and can cause fluctuations in the oligomeric state of protein complexes. To overcome these problems, we developed a method that does not use detergents or other chemicals. Here we report a detailed protocol that enables direct ejection of protein complexes from membranes for analysis by native MS. Briefly, lipid vesicles are prepared directly from membranes of different sources and subjected to sonication pulses. The resulting destabilized vesicles are concentrated, introduced into a mass spectrometer and ionized. The mass of the observed protein complexes is determined and this information, in conjunction with ‘omics’-based strategies, is used to determine subunit stoichiometry as well as cofactor and lipid binding. Within this protocol, we expand the applications of the method to include peripheral membrane proteins of the S-layer and amyloid protein export machineries overexpressed in membranes from which the most abundant components have been removed. The described experimental procedure takes approximately 3 d from preparation to MS. The time required for data analysis depends on the complexity of the protein assemblies embedded in the membrane under investigation.
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Data availability
Raw data presented in this paper are either deposited in https://doi.org/10.6084/m9.figshare.11376015 or available on request.
Code availability
No code has been written for this manuscript. The software needed is stated and referenced in the text.
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Acknowledgements
The authors thank all C.V.R. group members for helpful discussions, L. Zhou for her help in photographing the capillaries, S. Chen for her help in photographing the experimental setup, R. Dhaliwal and E. Johnson from the Oxford Dunn School EM facility for scanning EM imaging, T. Zeev-Ben-Mordehai for her technical assistance with MPEEVs and H. D. Bernstein for providing the plasmid pJH114 for expressing the Bam complex. D.S.C., H.T. and C.V.R. are grateful for the support of an ERC grant (695511- ENABLE), and J.G., D.W. and C.V.R. for support by a Wellcome Trust Investigator Award (104633/Z/14/Z). J.R.B. and C.V.R. are supported by a Medical Research Council grant (MR/N020413/1). T.A.M.B. is supported by the Wellcome Trust and the Royal Society (grant no. 202231/Z/16/Z). A.v.K., T.A.M.B. and C.V.R. thank the Vallee Foundation for support. S.L.R. and S.J.M. are supported by a Wellcome Trust (Senior Investigator Award 100280). J.G. is a Junior Research Fellow at Queen’s College, University of Oxford. L.A.B. was supported by a Human Frontier Science Program Long Term Fellowship and a Canadian Institutes for Health Research Postdoctoral Fellowship. K.G. was supported by a Wellcome Trust Senior Research Fellowship (090895/Z/09/Z) and a core award (090532/Z/09/Z).
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D.S.C. performed the MS experiments. H.T. performed the MS experiments for S-layer proteins and membranes. J.G. optimized the Orbitrap UHMR platform for the transmission and detection of membrane proteins and to enable high-energy regimes. J.R.B. purified and analyzed the recombinant Bam complex. D.W. established and applied the lipidomics platform. S.L.R. and S.J.M. purified Fap proteins and membranes, and modelled the Fap complexes. A.v.K. and T.A.M.B. purified S-layer proteins and membranes, and performed cryo-EM of cell stalks. L.A.B. and K.G. performed the cryo-EM for inner membrane tubes. D.S.C. and C.V.R. supervised the research and wrote the manuscript with contributions from all authors.
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Chorev, D. S. et al. Science 362, 829–834 (2018): https://doi.org/10.1126/science.aau0976
Integrated supplementary information
Supplementary Fig. 1 Large protein complexes containing gB eject in high-energy conditions.
(a) SDS-gel of MPEEV containing the protein gB. (b) Native gel of the vesicles shows only one protein complex at ~480 kDa. (c) The vesicles are first analysed at 400 V (source fragmentation 300 V, HCD 100 V). As the energy is increased, more proteins are released from the vesicle, at 500 V (additional 100 V in the HCD cell), monomers of gB start appearing, whereas at 600 V (additional 200 V in the HCD cell), a 465 kD complex appears. 2 biological repeats and 4 overall injections were performed.
Supplementary Fig. 2 BN and SDS–PAGE of samples.
(a) BN-PAGE of E. coli membranes, without major outer membrane complexes, overexpressing the fap operon with and without DDM indicates the existence of two 180–200 kDa membrane protein complexes containing Fap proteins in the presence of DDM. (b) SDS–PAGE of C. crescentus membrane stalks containing the S-layer or extracted S-layer protein RsaA shows degradation products following low pH extraction.
Supplementary Fig. 3 Low m/z region of protein from the fap operon purified in detergent shows different FapD species dissociating from the FapF3–FapD complex.
High energy disruption of the FapF3–FapD complexes at 400 V induces their dissociation and reveals the co-existence of three different FapD species—a correctly processed FapD at 23,313 Da, an incompletely processed FapD’ at 24,596 Da and a self-processed FapD’’ with a mass of 22,488 Da. Spectrum is a representative from 2 biological repeats.
Supplementary Fig. 4 Mass spectrum of low-pH-extracted RsaA.
Mass spectrum of solution extracted RsaA shows the presence of RsaA monomer and dimer with only a very low population of trimer. Inset (i) shows the organization of the S-layer associated with LPS and tethered to the outer membrane. The spectrum was acquired at a capillary voltage of 1.4 kV, capillary temperature of 50 °C, with source fragmentation set to 100 V, desolvation voltage set to 0 V and HCD energy at 0 V.
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Supplementary Figs. 1–4 and Supplementary Table 1.
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Chorev, D.S., Tang, H., Rouse, S.L. et al. The use of sonicated lipid vesicles for mass spectrometry of membrane protein complexes. Nat Protoc 15, 1690–1706 (2020). https://doi.org/10.1038/s41596-020-0303-y
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DOI: https://doi.org/10.1038/s41596-020-0303-y
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