51
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Affiliation(s)
- James E. Keener
- Department of Chemistry and Biochemistry, University of Arizona, Tucson, AZ 85721, USA
| | - Guozhi Zhang
- Department of Chemistry and Biochemistry, University of Arizona, Tucson, AZ 85721, USA
| | - Michael T. Marty
- Department of Chemistry and Biochemistry, University of Arizona, Tucson, AZ 85721, USA
- Bio5 Institute, University of Arizona, Tucson, AZ 85721, USA
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52
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Bagheri Y, Ali AA, You M. Current Methods for Detecting Cell Membrane Transient Interactions. Front Chem 2020; 8:603259. [PMID: 33365301 PMCID: PMC7750205 DOI: 10.3389/fchem.2020.603259] [Citation(s) in RCA: 21] [Impact Index Per Article: 5.3] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/06/2020] [Accepted: 10/16/2020] [Indexed: 12/28/2022] Open
Abstract
Short-lived cell membrane complexes play a key role in regulating cell signaling and communication. Many of these complexes are formed based on low-affinity and transient interactions among various lipids and proteins. New techniques have emerged to study these previously overlooked membrane transient interactions. Exciting functions of these transient interactions have been discovered in cellular events such as immune signaling, host-pathogen interactions, and diseases such as cancer. In this review, we have summarized current experimental methods that allow us to detect and analyze short-lived cell membrane protein-protein, lipid-protein, and lipid-lipid interactions. These methods can provide useful information about the strengths, kinetics, and/or spatial patterns of membrane transient interactions. However, each method also has its own limitations. We hope this review can be used as a guideline to help the audience to choose proper approaches for studying membrane transient interactions in different membrane trafficking and cell signaling events.
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Affiliation(s)
| | | | - Mingxu You
- Department of Chemistry, University of Massachusetts, Amherst, MA, United States
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53
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Jiao F, Cannon KS, Lin YC, Gladfelter AS, Scheuring S. The hierarchical assembly of septins revealed by high-speed AFM. Nat Commun 2020; 11:5062. [PMID: 33033254 PMCID: PMC7545167 DOI: 10.1038/s41467-020-18778-x] [Citation(s) in RCA: 18] [Impact Index Per Article: 4.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/24/2020] [Accepted: 09/15/2020] [Indexed: 12/15/2022] Open
Abstract
Septins are GTP-binding proteins involved in diverse cellular processes including division and membrane remodeling. Septins form linear, palindromic heteromeric complexes that can assemble in filaments and higher-order structures. Structural studies revealed various septin architectures, but questions concerning assembly-dynamics and -pathways persist. Here we used high-speed atomic force microscopy (HS-AFM) and kinetic modeling which allowed us to determine that septin filament assembly was a diffusion-driven process, while formation of higher-order structures was complex and involved self-templating. Slightly acidic pH and increased monovalent ion concentrations favor filament-assembly, -alignment and -pairing. Filament-alignment and -pairing further favored diffusion-driven assembly. Pairing is mediated by the septin N-termini face, and may occur symmetrically or staggered, likely important for the formation of higher-order structures of different shapes. Multilayered structures are templated by the morphology of the underlying layers. The septin C-termini face, namely the C-terminal extension of Cdc12, may be involved in membrane binding.
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Affiliation(s)
- Fang Jiao
- Department of Anesthesiology, Weill Cornell Medicine, New York, NY, 10065, USA
- Department of Physiology and Biophysics, Weill Cornell Medicine, New York, NY, 10065, USA
| | - Kevin S Cannon
- Department of Biology, University of North Carolina and Chapel Hill, Chapel Hill, NC, 27599, USA
| | - Yi-Chih Lin
- Department of Anesthesiology, Weill Cornell Medicine, New York, NY, 10065, USA
- Department of Physiology and Biophysics, Weill Cornell Medicine, New York, NY, 10065, USA
| | - Amy S Gladfelter
- Department of Biology, University of North Carolina and Chapel Hill, Chapel Hill, NC, 27599, USA
- Marine Biological Laboratory, Woods Hole, MA, 02543, USA
| | - Simon Scheuring
- Department of Anesthesiology, Weill Cornell Medicine, New York, NY, 10065, USA.
- Department of Physiology and Biophysics, Weill Cornell Medicine, New York, NY, 10065, USA.
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54
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de Vlugt JE, Xiao P, Munro R, Charchoglyan A, Brewer D, Al-Abdul-Wahid MS, Brown LS, Ladizhansky V. Identifying lipids tightly bound to an integral membrane protein. BIOCHIMICA ET BIOPHYSICA ACTA-BIOMEMBRANES 2020; 1862:183345. [DOI: 10.1016/j.bbamem.2020.183345] [Citation(s) in RCA: 8] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Track Full Text] [Subscribe] [Scholar Register] [Received: 02/16/2020] [Revised: 04/11/2020] [Accepted: 05/05/2020] [Indexed: 01/02/2023]
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55
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Park J, Fu Z, Frangaj A, Liu J, Mosyak L, Shen T, Slavkovich VN, Ray KM, Taura J, Cao B, Geng Y, Zuo H, Kou Y, Grassucci R, Chen S, Liu Z, Lin X, Williams JP, Rice WJ, Eng ET, Huang RK, Soni RK, Kloss B, Yu Z, Javitch JA, Hendrickson WA, Slesinger PA, Quick M, Graziano J, Yu H, Fiehn O, Clarke OB, Frank J, Fan QR. Structure of human GABA B receptor in an inactive state. Nature 2020; 584:304-309. [PMID: 32581365 PMCID: PMC7725281 DOI: 10.1038/s41586-020-2452-0] [Citation(s) in RCA: 56] [Impact Index Per Article: 14.0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/08/2019] [Accepted: 05/01/2020] [Indexed: 01/21/2023]
Abstract
The human GABAB receptor-a member of the class C family of G-protein-coupled receptors (GPCRs)-mediates inhibitory neurotransmission and has been implicated in epilepsy, pain and addiction1. A unique GPCR that is known to require heterodimerization for function2-6, the GABAB receptor has two subunits, GABAB1 and GABAB2, that are structurally homologous but perform distinct and complementary functions. GABAB1 recognizes orthosteric ligands7,8, while GABAB2 couples with G proteins9-14. Each subunit is characterized by an extracellular Venus flytrap (VFT) module, a descending peptide linker, a seven-helix transmembrane domain and a cytoplasmic tail15. Although the VFT heterodimer structure has been resolved16, the structure of the full-length receptor and its transmembrane signalling mechanism remain unknown. Here we present a near full-length structure of the GABAB receptor, captured in an inactive state by cryo-electron microscopy. Our structure reveals several ligands that preassociate with the receptor, including two large endogenous phospholipids that are embedded within the transmembrane domains to maintain receptor integrity and modulate receptor function. We also identify a previously unknown heterodimer interface between transmembrane helices 3 and 5 of both subunits, which serves as a signature of the inactive conformation. A unique 'intersubunit latch' within this transmembrane interface maintains the inactive state, and its disruption leads to constitutive receptor activity.
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Affiliation(s)
- Jinseo Park
- Department of Pharmacology, Columbia University, New York, NY, USA
| | - Ziao Fu
- Department of Biochemistry and Molecular Biophysics, Columbia University, New York, NY, USA
| | - Aurel Frangaj
- Department of Pharmacology, Columbia University, New York, NY, USA
| | - Jonathan Liu
- Department of Pharmacology, Columbia University, New York, NY, USA
| | - Lidia Mosyak
- Department of Pharmacology, Columbia University, New York, NY, USA
| | - Tong Shen
- NIH West Coast Metabolomics Center, University of California Davis, Davis, CA, USA
| | - Vesna N Slavkovich
- Department of Environmental Health Sciences, Columbia University, New York, NY, USA
| | - Kimberly M Ray
- Department of Pharmacology, Columbia University, New York, NY, USA
| | - Jaume Taura
- Nash Family Department of Neuroscience, Friedman Brain Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA
| | - Baohua Cao
- Department of Pharmacology, Columbia University, New York, NY, USA
| | - Yong Geng
- Department of Pharmacology, Columbia University, New York, NY, USA
- Key Laboratory of Receptor Research, Center for Structure and Function of Drug Targets, Shanghai Institute of Materia Medica, Chinese Academy of Sciences, Shanghai, China
| | - Hao Zuo
- Department of Pharmacology, Columbia University, New York, NY, USA
| | - Yongjun Kou
- Key Laboratory of Receptor Research, Center for Structure and Function of Drug Targets, Shanghai Institute of Materia Medica, Chinese Academy of Sciences, Shanghai, China
| | - Robert Grassucci
- Department of Biochemistry and Molecular Biophysics, Columbia University, New York, NY, USA
| | - Shaoxia Chen
- MRC Laboratory of Molecular Biology, Cambridge, UK
| | - Zheng Liu
- Department of Biochemistry and Molecular Biophysics, Columbia University, New York, NY, USA
| | - Xin Lin
- Department of Psychiatry, Columbia University, New York, NY, USA
- Division of Molecular Therapeutics, New York State Psychiatric Institute, New York, NY, USA
| | - Justin P Williams
- Department of Physiology and Cellular Biophysics, Columbia University, New York, NY, USA
| | - William J Rice
- National Resource for Automated Molecular Microscopy, Simons Electron Microscopy Center, New York Structural Biology Center, New York, NY, USA
| | - Edward T Eng
- National Resource for Automated Molecular Microscopy, Simons Electron Microscopy Center, New York Structural Biology Center, New York, NY, USA
| | - Rick K Huang
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA, USA
| | - Rajesh K Soni
- Proteomics Shared Resource, Herbert Irving Comprehensive Cancer Center, Columbia University, New York, NY, USA
| | - Brian Kloss
- Center on Membrane Protein Production and Analysis, New York Structural Biology Center, New York, NY, USA
| | - Zhiheng Yu
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA, USA
| | - Jonathan A Javitch
- Department of Pharmacology, Columbia University, New York, NY, USA
- Department of Psychiatry, Columbia University, New York, NY, USA
- Division of Molecular Therapeutics, New York State Psychiatric Institute, New York, NY, USA
- Department of Physiology and Cellular Biophysics, Columbia University, New York, NY, USA
| | - Wayne A Hendrickson
- Department of Biochemistry and Molecular Biophysics, Columbia University, New York, NY, USA
- Department of Physiology and Cellular Biophysics, Columbia University, New York, NY, USA
- Center on Membrane Protein Production and Analysis, New York Structural Biology Center, New York, NY, USA
| | - Paul A Slesinger
- Nash Family Department of Neuroscience, Friedman Brain Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA
| | - Matthias Quick
- Department of Psychiatry, Columbia University, New York, NY, USA
- Division of Molecular Therapeutics, New York State Psychiatric Institute, New York, NY, USA
| | - Joseph Graziano
- Department of Environmental Health Sciences, Columbia University, New York, NY, USA
| | - Hongtao Yu
- Howard Hughes Medical Institute, Department of Pharmacology, University of Texas Southwestern Medical Center, Dallas, TX, USA
| | - Oliver Fiehn
- NIH West Coast Metabolomics Center, University of California Davis, Davis, CA, USA
| | - Oliver B Clarke
- Department of Physiology and Cellular Biophysics, Columbia University, New York, NY, USA.
- Department of Anesthesiology and the Irving Institute for Clinical and Translational Research, Columbia University, New York, NY, USA.
| | - Joachim Frank
- Department of Biochemistry and Molecular Biophysics, Columbia University, New York, NY, USA.
- Department of Biological Sciences, Columbia University, New York, NY, USA.
| | - Qing R Fan
- Department of Pharmacology, Columbia University, New York, NY, USA.
- Department of Pathology and Cell Biology, Columbia University, New York, NY, USA.
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56
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Cleary SP, Prell JS. Distinct classes of multi-subunit heterogeneity: analysis using Fourier Transform methods and native mass spectrometry. Analyst 2020; 145:4688-4697. [PMID: 32459233 PMCID: PMC8483610 DOI: 10.1039/d0an00726a] [Citation(s) in RCA: 9] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/31/2023]
Abstract
Native electrospray mass spectrometry is a powerful method for determining the native stoichiometry of many polydisperse multi-subunit biological complexes, including multi-subunit protein complexes and lipid-bound transmembrane proteins. However, when polydispersity results from incorporation of multiple copies of two or more different subunits, it can be difficult to analyze subunit stoichiometry using conventional mass spectrometry analysis methods, especially when m/z distributions for different charge states overlap in the mass spectrum. It was recently demonstrated by Marty and co-workers (K. K. Hoi, et al., Anal. Chem., 2016, 88, 6199-6204) that Fourier Transform (FT)-based methods can determine the bulk average lipid composition of protein-lipid Nanodiscs assembled with two different lipids, but a detailed statistical description of the composition of more general polydisperse two-subunit populations is still difficult to achieve. This results from the vast number of ways in which the two types of subunit can be distributed within the analyte ensemble. Here, we present a theoretical description of three common classes of heterogeneity for mixed-subunit analytes and demonstrate how to differentiate and analyze them using mass spectrometry and FT methods. First, we first describe FT-based analysis of mass spectra corresponding to simple superpositions, convolutions, and multinomial distributions for two or more different subunit types using model data sets. We then apply these principles with real samples, including mixtures of single-lipid Nanodiscs in the same solution (superposition), mixed-lipid Nanodiscs and copolymers (convolutions), and isotope distribution for ubiquitin (multinomial distribution). This classification scheme and the FT method used to study these analyte classes should be broadly useful in mass spectrometry as well as other techniques where overlapping, periodic signals arising from analyte mixtures are common.
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Affiliation(s)
- Sean P Cleary
- Department of Chemistry and Biochemistry, 1253 University of Oregon, Eugene, OR 97403-1253, USA.
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57
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Kim J, Mandal R, Hilty C. Characterization of Membrane Protein‐Lipid Interactions in Unfolded OmpX with Enhanced Time Resolution by Hyperpolarized NMR. Chembiochem 2020; 21:2861-2867. [DOI: 10.1002/cbic.202000271] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/04/2020] [Indexed: 12/17/2022]
Affiliation(s)
- Jihyun Kim
- Department of Chemistry Texas A&M University College Station TX 77843-3255 USA
| | - Ratnamala Mandal
- Department of Chemistry Texas A&M University College Station TX 77843-3255 USA
| | - Christian Hilty
- Department of Chemistry Texas A&M University College Station TX 77843-3255 USA
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58
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Puglisi R, Boeri Erba E, Pastore A. A Guide to Native Mass Spectrometry to determine complex interactomes of molecular machines. FEBS J 2020; 287:2428-2439. [PMID: 32142206 PMCID: PMC8647915 DOI: 10.1111/febs.15281] [Citation(s) in RCA: 9] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/08/2019] [Revised: 02/01/2020] [Accepted: 03/04/2020] [Indexed: 01/17/2023]
Abstract
Native mass spectrometry is an emerging technique in biology that gives the possibility to study noncovalently bound complexes with high sensitivity and accuracy. It thus allows the characterization of macromolecular assemblies, assessing their mass and stoichiometries and mapping the interacting surfaces. In this review, we discuss the application of native mass spectrometry to dynamic molecular machines based on multiple weak interactions. In the study of these machines, it is crucial to understand which and under which conditions various complexes form at any time point. We focus on the specific example of the iron-sulfur cluster biogenesis machine because this is an archetype of a dynamic machine that requires very specific and demanding experimental conditions, such as anaerobicity and the need of retaining the fold of marginally folded proteins. We describe the advantages, challenges and current limitations of the technique by providing examples from our own experience and suggesting possible future solutions.
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Affiliation(s)
- Rita Puglisi
- UK Dementia Research Institute at the Wohl Institute of King’s College LondonUK
| | | | - Annalisa Pastore
- UK Dementia Research Institute at the Wohl Institute of King’s College LondonUK
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59
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Exploring the structure and dynamics of macromolecular complexes by native mass spectrometry. J Proteomics 2020; 222:103799. [DOI: 10.1016/j.jprot.2020.103799] [Citation(s) in RCA: 26] [Impact Index Per Article: 6.5] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/28/2019] [Revised: 03/23/2020] [Accepted: 04/25/2020] [Indexed: 12/15/2022]
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60
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Sobti M, Walshe JL, Wu D, Ishmukhametov R, Zeng YC, Robinson CV, Berry RM, Stewart AG. Cryo-EM structures provide insight into how E. coli F 1F o ATP synthase accommodates symmetry mismatch. Nat Commun 2020; 11:2615. [PMID: 32457314 PMCID: PMC7251095 DOI: 10.1038/s41467-020-16387-2] [Citation(s) in RCA: 63] [Impact Index Per Article: 15.8] [Reference Citation Analysis] [Abstract] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/29/2019] [Accepted: 04/30/2020] [Indexed: 12/19/2022] Open
Abstract
F1Fo ATP synthase functions as a biological rotary generator that makes a major contribution to cellular energy production. It comprises two molecular motors coupled together by a central and a peripheral stalk. Proton flow through the Fo motor generates rotation of the central stalk, inducing conformational changes in the F1 motor that catalyzes ATP production. Here we present nine cryo-EM structures of E. coli ATP synthase to 3.1-3.4 Å resolution, in four discrete rotational sub-states, which provide a comprehensive structural model for this widely studied bacterial molecular machine. We observe torsional flexing of the entire complex and a rotational sub-step of Fo associated with long-range conformational changes that indicates how this flexibility accommodates the mismatch between the 3- and 10-fold symmetries of the F1 and Fo motors. We also identify density likely corresponding to lipid molecules that may contribute to the rotor/stator interaction within the Fo motor.
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Affiliation(s)
- Meghna Sobti
- Molecular, Structural and Computational Biology Division, The Victor Chang Cardiac Research Institute, Darlinghurst, NSW, 2010, Australia.,Faculty of Medicine, St Vincent's Clinical School, UNSW Sydney, Kensington, NSW, 2052, Australia
| | - James L Walshe
- Molecular, Structural and Computational Biology Division, The Victor Chang Cardiac Research Institute, Darlinghurst, NSW, 2010, Australia
| | - Di Wu
- Department of Chemistry, University of Oxford, Oxford, OX1 3QZ, United Kingdom
| | - Robert Ishmukhametov
- Clarendon Laboratory, Department of Physics, University of Oxford, Oxford, OX1 3PU, United Kingdom
| | - Yi C Zeng
- Molecular, Structural and Computational Biology Division, The Victor Chang Cardiac Research Institute, Darlinghurst, NSW, 2010, Australia
| | - Carol V Robinson
- Department of Chemistry, University of Oxford, Oxford, OX1 3QZ, United Kingdom
| | - Richard M Berry
- Clarendon Laboratory, Department of Physics, University of Oxford, Oxford, OX1 3PU, United Kingdom
| | - Alastair G Stewart
- Molecular, Structural and Computational Biology Division, The Victor Chang Cardiac Research Institute, Darlinghurst, NSW, 2010, Australia. .,Faculty of Medicine, St Vincent's Clinical School, UNSW Sydney, Kensington, NSW, 2052, Australia.
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61
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Chorev DS, Tang H, Rouse SL, Bolla JR, von Kügelgen A, Baker LA, Wu D, Gault J, Grünewald K, Bharat TAM, Matthews SJ, Robinson CV. The use of sonicated lipid vesicles for mass spectrometry of membrane protein complexes. Nat Protoc 2020; 15:1690-1706. [PMID: 32238951 PMCID: PMC7305028 DOI: 10.1038/s41596-020-0303-y] [Citation(s) in RCA: 25] [Impact Index Per Article: 6.3] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/02/2019] [Accepted: 01/23/2020] [Indexed: 12/28/2022]
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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Affiliation(s)
- Dror S Chorev
- Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford, UK
| | - Haiping Tang
- Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford, UK
| | - Sarah L Rouse
- Department of Life Sciences, Imperial College London, London, UK
| | - Jani Reddy Bolla
- Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford, UK
| | - Andriko von Kügelgen
- Sir William Dunn School of Pathology, University of Oxford, Oxford, UK
- Central Oxford Structural Microscopy Imaging Centre, Oxford, UK
| | - Lindsay A Baker
- Division of Structural Biology, University of Oxford, Oxford, UK
| | - Di Wu
- Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford, UK
| | - Joseph Gault
- Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford, UK
| | - Kay Grünewald
- Division of Structural Biology, University of Oxford, Oxford, UK
- Heinrich Pette Institute, Leibniz-Institut für Experimentelle Virologie, Centre for Structural Systems Biology, c/o DESY, Hamburg, Germany
| | - Tanmay A M Bharat
- Sir William Dunn School of Pathology, University of Oxford, Oxford, UK
- Central Oxford Structural Microscopy Imaging Centre, Oxford, UK
| | | | - Carol V Robinson
- Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford, UK.
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62
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Corey RA, Stansfeld PJ, Sansom MS. The energetics of protein-lipid interactions as viewed by molecular simulations. Biochem Soc Trans 2020; 48:25-37. [PMID: 31872229 PMCID: PMC7054751 DOI: 10.1042/bst20190149] [Citation(s) in RCA: 37] [Impact Index Per Article: 9.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/27/2019] [Revised: 12/06/2019] [Accepted: 12/10/2019] [Indexed: 12/20/2022]
Abstract
Membranes are formed from a bilayer containing diverse lipid species with which membrane proteins interact. Integral, membrane proteins are embedded in this bilayer, where they interact with lipids from their surroundings, whilst peripheral membrane proteins bind to lipids at the surface of membranes. Lipid interactions can influence the function of membrane proteins, either directly or allosterically. Both experimental (structural) and computational approaches can reveal lipid binding sites on membrane proteins. It is, therefore, important to understand the free energies of these interactions. This affords a more complete view of the engagement of a particular protein with the biological membrane surrounding it. Here, we describe many computational approaches currently in use for this purpose, including recent advances using both free energy and unbiased simulation methods. In particular, we focus on interactions of integral membrane proteins with cholesterol, and with anionic lipids such as phosphatidylinositol 4,5-bis-phosphate and cardiolipin. Peripheral membrane proteins are exemplified via interactions of PH domains with phosphoinositide-containing membranes. We summarise the current state of the field and provide an outlook on likely future directions of investigation.
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Affiliation(s)
- Robin A. Corey
- Department of Biochemistry, University of Oxford, South Parks Road, Oxford OX1 3QU, U.K
| | - Phillip J. Stansfeld
- Department of Biochemistry, University of Oxford, South Parks Road, Oxford OX1 3QU, U.K
- School of Life Sciences and Department of Chemistry, University of Warwick, Coventry CV4 7AL, U.K
| | - Mark S.P. Sansom
- Department of Biochemistry, University of Oxford, South Parks Road, Oxford OX1 3QU, U.K
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63
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Li B, He X, Lei SS, Zhou FC, Zhang NY, Chen YH, Wang YZ, Su J, Yu JJ, Li LZ, Zheng X, Luo R, Kołodyńska D, Xiong S, Lv GY, Chen SH. Hypertensive Rats Treated Chronically With N ω-Nitro-L-Arginine Methyl Ester (L-NAME) Induced Disorder of Hepatic Fatty Acid Metabolism and Intestinal Pathophysiology. Front Pharmacol 2020; 10:1677. [PMID: 32076406 PMCID: PMC7006817 DOI: 10.3389/fphar.2019.01677] [Citation(s) in RCA: 23] [Impact Index Per Article: 5.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/29/2019] [Accepted: 12/23/2019] [Indexed: 01/14/2023] Open
Abstract
Nω-nitro-L-arginine methyl ester (L-NAME), an inhibitor of nitric oxide (NO) biosynthesis, results in hypertension and liver injury. This study aimed at investigating the changes of liver lipometabonomics and exploring the underlying mechanisms of liver injury in the L-NAME-treated rats. The male Sprague-Dawley (SD) rats were treated with L-NAME (40 mg/kg, p.o.) for 8 weeks. After that, the liver, aorta, fecal, and serum were collected for analysis. The results showed that L-NAME induced hypertension and disordered the endothelial nitric oxide synthase (eNOS)-NO pathway in the treated rats. L-NAME could also increase the levels of serum total cholesterol (TC), triglyceride (TG), alanine transaminase (ALT), and aspartate transaminase (AST). The multidimensional mass spectrometry-based shotgun lipidomics (MDMS-SL) analysis showed that L-NAME could induce significant changes of the total hepatic lipids and most hepatic triglycerides, as well as fatty acid (FA). A positive correlation was found between the blood pressure and TAG. Immunofluorescence and Western-Blot experiments indicated that the L-NAME treatment significantly influenced some FA β-oxidation, desaturation, and synthesis-related proteins. The increase of intestinal inflammation, decrease of microcirculation and tight junction proteins, as well as alterations of microbial communities were observed in the L-NAME induced hypertensive rats, as well as alterations of microbial communities were notable correlation to TAG and FA species. This study demonstrated that the L-NAME-induced hypertensive rats exhibiting liver injury were the joint action of hepatic abnormal fatty acid metabolism and microcirculation disorder. Furthermore, the gut microflora, as well as the changes of FA β-oxidation (ACOX, CPT1α), desaturation (SCD-1), and synthesis (FAS) may be the potential mechanisms for abnormal fatty acid metabolism.
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Affiliation(s)
- Bo Li
- Collaborative Innovation Center of Yangtze River Delta Region Green Pharmaceuticals, Zhejiang University of Technology, Hangzhou, China
| | - Xinglishang He
- Collaborative Innovation Center of Yangtze River Delta Region Green Pharmaceuticals, Zhejiang University of Technology, Hangzhou, China
| | - Shan-Shan Lei
- Collaborative Innovation Center of Yangtze River Delta Region Green Pharmaceuticals, Zhejiang University of Technology, Hangzhou, China
| | - Fu-Chen Zhou
- Collaborative Innovation Center of Yangtze River Delta Region Green Pharmaceuticals, Zhejiang University of Technology, Hangzhou, China
| | - Ning-Yu Zhang
- Collaborative Innovation Center of Yangtze River Delta Region Green Pharmaceuticals, Zhejiang University of Technology, Hangzhou, China
| | - Ye-Hui Chen
- Collaborative Innovation Center of Yangtze River Delta Region Green Pharmaceuticals, Zhejiang University of Technology, Hangzhou, China
| | - Yu-Zhi Wang
- Collaborative Innovation Center of Yangtze River Delta Region Green Pharmaceuticals, Zhejiang University of Technology, Hangzhou, China
| | - Jie Su
- College of Pharmaceutical Science, Zhejiang Chinese Medical University, Hangzhou, China
| | - Jing-Jing Yu
- College of Pharmaceutical Science, Zhejiang Chinese Medical University, Hangzhou, China
| | - Lin-Zi Li
- Collaborative Innovation Center of Yangtze River Delta Region Green Pharmaceuticals, Zhejiang University of Technology, Hangzhou, China
| | - Xiang Zheng
- Collaborative Innovation Center of Yangtze River Delta Region Green Pharmaceuticals, Zhejiang University of Technology, Hangzhou, China
| | - Rong Luo
- Collaborative Innovation Center of Yangtze River Delta Region Green Pharmaceuticals, Zhejiang University of Technology, Hangzhou, China
| | - Dorota Kołodyńska
- Department of Inorganic Chemistry, Faculty of Chemistry, Institute of Chemical Sciences, Maria Curie-Skłodowska University, Lublin, Poland
| | - Shan Xiong
- Institute of Materia Medica, Shandong First Medical University & Shandong Academy of Medical Sciences, Jinan, China
| | - Gui-Yuan Lv
- College of Pharmaceutical Science, Zhejiang Chinese Medical University, Hangzhou, China
| | - Su-Hong Chen
- Collaborative Innovation Center of Yangtze River Delta Region Green Pharmaceuticals, Zhejiang University of Technology, Hangzhou, China
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64
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Urner LH, Liko I, Yen HY, Hoi KK, Bolla JR, Gault J, Almeida FG, Schweder MP, Shutin D, Ehrmann S, Haag R, Robinson CV, Pagel K. Modular detergents tailor the purification and structural analysis of membrane proteins including G-protein coupled receptors. Nat Commun 2020; 11:564. [PMID: 31992701 PMCID: PMC6987200 DOI: 10.1038/s41467-020-14424-8] [Citation(s) in RCA: 67] [Impact Index Per Article: 16.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/18/2019] [Accepted: 12/19/2019] [Indexed: 01/29/2023] Open
Abstract
Detergents enable the purification of membrane proteins and are indispensable reagents in structural biology. Even though a large variety of detergents have been developed in the last century, the challenge remains to identify guidelines that allow fine-tuning of detergents for individual applications in membrane protein research. Addressing this challenge, here we introduce the family of oligoglycerol detergents (OGDs). Native mass spectrometry (MS) reveals that the modular OGD architecture offers the ability to control protein purification and to preserve interactions with native membrane lipids during purification. In addition to a broad range of bacterial membrane proteins, OGDs also enable the purification and analysis of a functional G-protein coupled receptor (GPCR). Moreover, given the modular design of these detergents, we anticipate fine-tuning of their properties for specific applications in structural biology. Seen from a broader perspective, this represents a significant advance for the investigation of membrane proteins and their interactions with lipids. Detergents are indispensable reagents in membrane protein structural biology. Here, L. H. Urner and co-workers introduce oligoglycerol detergents (OGDs) and use native mass spectrometry to show how interactions of membrane proteins with native membrane lipids can be preserved during purification.
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Affiliation(s)
- Leonhard H Urner
- Institute of Chemistry and Biochemistry, Freie Universität Berlin, 14195, Berlin, Germany.,Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford, OX1 3QZ, UK
| | - Idlir Liko
- Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford, OX1 3QZ, UK.,OMass Therapeutics, The Schrödinger Building, Heatley Road, The Oxford Science Park, Oxford, OX4 4GE, UK
| | - Hsin-Yung Yen
- Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford, OX1 3QZ, UK.,OMass Therapeutics, The Schrödinger Building, Heatley Road, The Oxford Science Park, Oxford, OX4 4GE, UK
| | - Kin-Kuan Hoi
- Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford, OX1 3QZ, UK
| | - Jani Reddy Bolla
- Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford, OX1 3QZ, UK
| | - Joseph Gault
- Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford, OX1 3QZ, UK
| | - Fernando Gonçalves Almeida
- OMass Therapeutics, The Schrödinger Building, Heatley Road, The Oxford Science Park, Oxford, OX4 4GE, UK
| | - Marc-Philip Schweder
- Institute of Chemistry and Biochemistry, Freie Universität Berlin, 14195, Berlin, Germany
| | - Denis Shutin
- Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford, OX1 3QZ, UK
| | - Svenja Ehrmann
- Institute of Chemistry and Biochemistry, Freie Universität Berlin, 14195, Berlin, Germany
| | - Rainer Haag
- Institute of Chemistry and Biochemistry, Freie Universität Berlin, 14195, Berlin, Germany.
| | - Carol V Robinson
- Physical and Theoretical Chemistry Laboratory, University of Oxford, Oxford, OX1 3QZ, UK.
| | - Kevin Pagel
- Institute of Chemistry and Biochemistry, Freie Universität Berlin, 14195, Berlin, Germany. .,Department of Molecular Physics, Fritz Haber Institute of the Max Planck Society, 14195, Berlin, Germany.
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65
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Styrene maleic-acid lipid particles (SMALPs) into detergent or amphipols: An exchange protocol for membrane protein characterisation. BIOCHIMICA ET BIOPHYSICA ACTA-BIOMEMBRANES 2020; 1862:183192. [PMID: 31945320 PMCID: PMC7086155 DOI: 10.1016/j.bbamem.2020.183192] [Citation(s) in RCA: 23] [Impact Index Per Article: 5.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Figures] [Subscribe] [Scholar Register] [Received: 10/01/2019] [Revised: 12/06/2019] [Accepted: 01/10/2020] [Indexed: 12/24/2022]
Abstract
Membrane proteins are traditionally extracted and purified in detergent for biochemical and structural characterisation. This process is often costly and laborious, and the stripping away of potentially stabilising lipids from the membrane protein of interest can have detrimental effects on protein integrity. Recently, styrene-maleic acid (SMA) co-polymers have offered a solution to this problem by extracting membrane proteins directly from their native membrane, while retaining their naturally associated lipids in the form of stable SMA lipid particles (SMALPs). However, the inherent nature and heterogeneity of the polymer renders their use challenging for some downstream applications – particularly mass spectrometry (MS). While advances in cryo-electron microscopy (cryo-EM) have enhanced our understanding of membrane protein:lipid interactions in both SMALPs and detergent, the resolution obtained with this technique is often insufficient to accurately identify closely associated lipids within the transmembrane annulus. Native-MS has the power to fill this knowledge gap, but the SMA polymer itself remains largely incompatible with this technique. To increase sample homogeneity and allow characterisation of membrane protein:lipid complexes by native-MS, we have developed a novel SMA-exchange method; whereby the membrane protein of interest is first solubilised and purified in SMA, then transferred into amphipols or detergents. This allows the membrane protein and endogenously associated lipids extracted by SMA co-polymer to be identified and examined by MS, thereby complementing results obtained by cryo-EM and creating a better understanding of how the lipid bilayer directly affects membrane protein structure and function. First reported exchange protocol for transferring membrane proteins solubilised in SMALPs, into detergent or amphipols. Purification of protein:lipid complexes without detergent for mass spectrometry and subsequent lipid identification. Cost effective membrane protein purification requiring only minimal amounts of detergents in the exchange process.
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66
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Martens C, Shekhar M, Lau AM, Tajkhorshid E, Politis A. Integrating hydrogen-deuterium exchange mass spectrometry with molecular dynamics simulations to probe lipid-modulated conformational changes in membrane proteins. Nat Protoc 2019; 14:3183-3204. [PMID: 31605097 PMCID: PMC7058097 DOI: 10.1038/s41596-019-0219-6] [Citation(s) in RCA: 30] [Impact Index Per Article: 6.0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/03/2019] [Accepted: 06/20/2019] [Indexed: 12/20/2022]
Abstract
Biological membranes define the boundaries of cells and are composed primarily of phospholipids and membrane proteins. It has become increasingly evident that direct interactions of membrane proteins with their surrounding lipids play key roles in regulating both protein conformations and function. However, the exact nature and structural consequences of these interactions remain difficult to track at the molecular level. Here, we present a protocol that specifically addresses this challenge. First, hydrogen-deuterium exchange mass spectrometry (HDX-MS) of membrane proteins incorporated into nanodiscs of controlled lipid composition is used to obtain information on the lipid species that are involved in modulating the conformational changes in the membrane protein. Then molecular dynamics (MD) simulations in lipid bilayers are used to pinpoint likely lipid-protein interactions, which can be tested experimentally using HDX-MS. By bringing together the MD predictions with the conformational readouts from HDX-MS, we have uncovered key lipid-protein interactions implicated in stabilizing important functional conformations. This protocol can be applied to virtually any integral membrane protein amenable to classic biophysical studies and for which a near-atomic-resolution structure or homology model is available. This protocol takes ~4 d to complete, excluding the time for data analysis and MD simulations, which depends on the size of the protein under investigation.
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Affiliation(s)
- Chloe Martens
- Department of Chemistry, King's College London, London, UK
- Department of Chemistry, Center for Structural Biology and Bioinformatics, Université Libre de Bruxelles, Brussels, Belgium
| | - Mrinal Shekhar
- Center for Biophysics and Quantitative Biology, Department of Biochemistry, NIH Center for Macromolecular Modeling and Bioinformatics, Beckman Institute for Advanced Science and Technology, University of Illinois at Urbana-Champaign, Urbana, IL, USA
| | - Andy M Lau
- Department of Chemistry, King's College London, London, UK
| | - Emad Tajkhorshid
- Center for Biophysics and Quantitative Biology, Department of Biochemistry, NIH Center for Macromolecular Modeling and Bioinformatics, Beckman Institute for Advanced Science and Technology, University of Illinois at Urbana-Champaign, Urbana, IL, USA.
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67
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Conrard L, Tyteca D. Regulation of Membrane Calcium Transport Proteins by the Surrounding Lipid Environment. Biomolecules 2019; 9:E513. [PMID: 31547139 PMCID: PMC6843150 DOI: 10.3390/biom9100513] [Citation(s) in RCA: 24] [Impact Index Per Article: 4.8] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/08/2019] [Revised: 09/09/2019] [Accepted: 09/10/2019] [Indexed: 12/11/2022] Open
Abstract
Calcium ions (Ca2+) are major messengers in cell signaling, impacting nearly every aspect of cellular life. Those signals are generated within a wide spatial and temporal range through a large variety of Ca2+ channels, pumps, and exchangers. More and more evidences suggest that Ca2+ exchanges are regulated by their surrounding lipid environment. In this review, we point out the technical challenges that are currently being overcome and those that still need to be defeated to analyze the Ca2+ transport protein-lipid interactions. We then provide evidences for the modulation of Ca2+ transport proteins by lipids, including cholesterol, acidic phospholipids, sphingolipids, and their metabolites. We also integrate documented mechanisms involved in the regulation of Ca2+ transport proteins by the lipid environment. Those include: (i) Direct interaction inside the protein with non-annular lipids; (ii) close interaction with the first shell of annular lipids; (iii) regulation of membrane biophysical properties (e.g., membrane lipid packing, thickness, and curvature) directly around the protein through annular lipids; and (iv) gathering and downstream signaling of several proteins inside lipid domains. We finally discuss recent reports supporting the related alteration of Ca2+ and lipids in different pathophysiological events and the possibility to target lipids in Ca2+-related diseases.
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Affiliation(s)
- Louise Conrard
- CELL Unit, de Duve Institute and Université catholique de Louvain, UCL B1.75.05, avenue Hippocrate, 75, B-1200 Brussels, Belgium
| | - Donatienne Tyteca
- CELL Unit, de Duve Institute and Université catholique de Louvain, UCL B1.75.05, avenue Hippocrate, 75, B-1200 Brussels, Belgium.
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68
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Corey RA, Vickery ON, Sansom MSP, Stansfeld PJ. Insights into Membrane Protein-Lipid Interactions from Free Energy Calculations. J Chem Theory Comput 2019; 15:5727-5736. [PMID: 31476127 PMCID: PMC6785801 DOI: 10.1021/acs.jctc.9b00548] [Citation(s) in RCA: 49] [Impact Index Per Article: 9.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/19/2022]
Abstract
![]()
Integral membrane proteins are regulated
by specific interactions
with lipids from the surrounding bilayer. The structures of protein–lipid
complexes can be determined through a combination of experimental
and computational approaches, but the energetic basis of these interactions
is difficult to resolve. Molecular dynamics simulations provide the
primary computational technique to estimate the free energies of these
interactions. We demonstrate that the energetics of protein–lipid
interactions may be reliably and reproducibly calculated using three
simulation-based approaches: potential of mean force calculations,
alchemical free energy perturbation, and well-tempered metadynamics.
We employ these techniques within the framework of a coarse-grained
force field and apply them to both bacterial and mammalian membrane
protein–lipid systems. We demonstrate good agreement between
the different techniques, providing a robust framework for their automated
implementation within a pipeline for annotation of newly determined
membrane protein structures.
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Affiliation(s)
- Robin A Corey
- Department of Biochemistry , University of Oxford , South Parks Road , Oxford OX1 3QU , U.K
| | - Owen N Vickery
- Department of Biochemistry , University of Oxford , South Parks Road , Oxford OX1 3QU , U.K
| | - Mark S P Sansom
- Department of Biochemistry , University of Oxford , South Parks Road , Oxford OX1 3QU , U.K
| | - Phillip J Stansfeld
- Department of Biochemistry , University of Oxford , South Parks Road , Oxford OX1 3QU , U.K
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69
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Frick M, Schmidt C. Mass spectrometry—A versatile tool for characterising the lipid environment of membrane protein assemblies. Chem Phys Lipids 2019; 221:145-157. [DOI: 10.1016/j.chemphyslip.2019.04.001] [Citation(s) in RCA: 20] [Impact Index Per Article: 4.0] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/31/2019] [Revised: 03/28/2019] [Accepted: 04/01/2019] [Indexed: 01/02/2023]
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70
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Marinko J, Huang H, Penn WD, Capra JA, Schlebach JP, Sanders CR. Folding and Misfolding of Human Membrane Proteins in Health and Disease: From Single Molecules to Cellular Proteostasis. Chem Rev 2019; 119:5537-5606. [PMID: 30608666 PMCID: PMC6506414 DOI: 10.1021/acs.chemrev.8b00532] [Citation(s) in RCA: 162] [Impact Index Per Article: 32.4] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/25/2018] [Indexed: 12/13/2022]
Abstract
Advances over the past 25 years have revealed much about how the structural properties of membranes and associated proteins are linked to the thermodynamics and kinetics of membrane protein (MP) folding. At the same time biochemical progress has outlined how cellular proteostasis networks mediate MP folding and manage misfolding in the cell. When combined with results from genomic sequencing, these studies have established paradigms for how MP folding and misfolding are linked to the molecular etiologies of a variety of diseases. This emerging framework has paved the way for the development of a new class of small molecule "pharmacological chaperones" that bind to and stabilize misfolded MP variants, some of which are now in clinical use. In this review, we comprehensively outline current perspectives on the folding and misfolding of integral MPs as well as the mechanisms of cellular MP quality control. Based on these perspectives, we highlight new opportunities for innovations that bridge our molecular understanding of the energetics of MP folding with the nuanced complexity of biological systems. Given the many linkages between MP misfolding and human disease, we also examine some of the exciting opportunities to leverage these advances to address emerging challenges in the development of therapeutics and precision medicine.
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Affiliation(s)
- Justin
T. Marinko
- Department
of Biochemistry, Vanderbilt University, Nashville, Tennessee 37240, United States
- Center
for Structural Biology, Vanderbilt University, Nashville, Tennessee 37240, United States
| | - Hui Huang
- Department
of Biochemistry, Vanderbilt University, Nashville, Tennessee 37240, United States
- Center
for Structural Biology, Vanderbilt University, Nashville, Tennessee 37240, United States
| | - Wesley D. Penn
- Department
of Chemistry, Indiana University, Bloomington, Indiana 47405, United States
| | - John A. Capra
- Center
for Structural Biology, Vanderbilt University, Nashville, Tennessee 37240, United States
- Department
of Biological Sciences, Vanderbilt University, Nashville, Tennessee 37245, United States
| | - Jonathan P. Schlebach
- Department
of Chemistry, Indiana University, Bloomington, Indiana 47405, United States
| | - Charles R. Sanders
- Department
of Biochemistry, Vanderbilt University, Nashville, Tennessee 37240, United States
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71
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Lipid Modulation of Membrane Protein Function. Cell Chem Biol 2019; 25:803-804. [PMID: 30028973 DOI: 10.1016/j.chembiol.2018.07.003] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/23/2022]
Abstract
In the last decade, native mass spectrometry has emerged as a powerful tool for studying the interactions of lipids and other small molecules with integral membrane proteins and their complexes. In this issue of Cell Chemical Biology, Pyle et al. (2018) establishes the role of phosphatidyl-inositol in dimerization and activity of a eukaryotic purine transporter (UapA).
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72
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Bolla JR, Agasid MT, Mehmood S, Robinson CV. Membrane Protein-Lipid Interactions Probed Using Mass Spectrometry. Annu Rev Biochem 2019; 88:85-111. [PMID: 30901263 DOI: 10.1146/annurev-biochem-013118-111508] [Citation(s) in RCA: 125] [Impact Index Per Article: 25.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/09/2022]
Abstract
Membrane proteins that exist in lipid bilayers are not isolated molecular entities. The lipid molecules that surround them play crucial roles in maintaining their full structural and functional integrity. Research directed at investigating these critical lipid-protein interactions is developing rapidly. Advancements in both instrumentation and software, as well as in key biophysical and biochemical techniques, are accelerating the field. In this review, we provide a brief outline of structural techniques used to probe protein-lipid interactions and focus on the molecular aspects of these interactions obtained from native mass spectrometry (native MS). We highlight examples in which lipids have been shown to modulate membrane protein structure and show how native MS has emerged as a complementary technique to X-ray crystallography and cryo-electron microscopy. We conclude with a short perspective on future developments that aim to better understand protein-lipid interactions in the native environment.
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Affiliation(s)
- Jani Reddy Bolla
- Department of Chemistry, University of Oxford, Oxford OX1 3QZ, United Kingdom;
| | - Mark T Agasid
- Department of Chemistry, University of Oxford, Oxford OX1 3QZ, United Kingdom;
| | - Shahid Mehmood
- Department of Chemistry, University of Oxford, Oxford OX1 3QZ, United Kingdom;
| | - Carol V Robinson
- Department of Chemistry, University of Oxford, Oxford OX1 3QZ, United Kingdom;
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73
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Hedger G, Koldsø H, Chavent M, Siebold C, Rohatgi R, Sansom MSP. Cholesterol Interaction Sites on the Transmembrane Domain of the Hedgehog Signal Transducer and Class F G Protein-Coupled Receptor Smoothened. Structure 2019; 27:549-559.e2. [PMID: 30595453 PMCID: PMC6408332 DOI: 10.1016/j.str.2018.11.003] [Citation(s) in RCA: 63] [Impact Index Per Article: 12.6] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/03/2018] [Revised: 09/28/2018] [Accepted: 11/02/2018] [Indexed: 12/12/2022]
Abstract
Transduction of Hedgehog signals across the plasma membrane is facilitated by the class F G-protein-coupled-receptor (GPCR) Smoothened (SMO). Recent studies suggest that SMO is modulated via interactions of its transmembrane (TM) domain with cholesterol. We apply molecular dynamics simulations of SMO embedded in cholesterol containing lipid bilayers, revealing a direct interaction of cholesterol with the TM domain at regions distinct from those observed in class A GPCRs. In particular the extracellular tips of helices TM2 and TM3 form a well-defined cholesterol interaction site. Potential of mean force calculations yield a free energy landscape for cholesterol binding. Alongside analysis of equilibrium cholesterol occupancy, this reveals the existence of a dynamic "greasy patch" interaction with the TM domain of SMO, which may be compared with previously identified lipid interaction sites on other membrane proteins. These predictions provide molecular-level insights into cholesterol interactions with a class F GPCR, suggesting potential druggable sites.
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Affiliation(s)
- George Hedger
- Department of Biochemistry, University of Oxford, South Parks Road, Oxford OX1 3QU, UK
| | - Heidi Koldsø
- Department of Biochemistry, University of Oxford, South Parks Road, Oxford OX1 3QU, UK
| | - Matthieu Chavent
- Institut de Pharmacologie et de Biologie Structurale IPBS, Université de Toulouse, CNRS, UPS, Toulouse, France
| | - Christian Siebold
- Division of Structural Biology, Wellcome Centre for Human Genetics, University of Oxford, Oxford OX3 7BN, UK
| | - Rajat Rohatgi
- Departments of Biochemistry and Medicine, Stanford University School of Medicine, Stanford, CA 94305, USA
| | - Mark S P Sansom
- Department of Biochemistry, University of Oxford, South Parks Road, Oxford OX1 3QU, UK.
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74
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Nguyen GH, Tran TN, Podgorski MN, Bell SG, Supuran CT, Donald WA. Nanoscale Ion Emitters in Native Mass Spectrometry for Measuring Ligand-Protein Binding Affinities. ACS CENTRAL SCIENCE 2019; 5:308-318. [PMID: 30834319 PMCID: PMC6396573 DOI: 10.1021/acscentsci.8b00787] [Citation(s) in RCA: 72] [Impact Index Per Article: 14.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Received: 10/29/2018] [Indexed: 05/20/2023]
Abstract
Electrospray ionization (ESI) mass spectrometry (MS) is a crucial method for rapidly determining the interactions between small molecules and proteins with ultrahigh sensitivity. However, nonvolatile molecules and salts that are often necessary to stabilize the native structures of protein-ligand complexes can readily adduct to protein ions, broaden spectral peaks, and lower signal-to-noise ratios in native MS. ESI emitters with narrow tip diameters (∼250 nm) were used to significantly reduce the extent of adduction of salt and nonvolatile molecules to protein complexes to more accurately measure ligand-protein binding constants than by use of conventional larger-bore emitters under these conditions. As a result of decreased salt adduction, peaks corresponding to protein-ligand complexes that differ in relative molecular weight by as low as 0.06% can be readily resolved. For low-molecular-weight anion ligands formed from sodium salts, anion-bound and unbound protein ions that differ in relative mass by 0.2% were completely baseline resolved using nanoscale emitters, which was not possible under these conditions using conventional emitters. Owing to the improved spectral resolution obtained using narrow-bore emitters and an analytically derived equation, K d values were simultaneously obtained for at least six ligands to a single druggable protein target from one spectrum for the first time. This research suggests that ligand-protein binding constants can be directly and accurately measured from solutions with high concentrations of nonvolatile buffers and salts by native MS.
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Affiliation(s)
- Giang
T. H. Nguyen
- School
of Chemistry, University of New South Wales, Dalton Building, Sydney, New South Wales 2052, Australia
| | - Thinh N. Tran
- School
of Electrical Engineering and Telecommunications, University of New South Wales, Sydney, New South Wales 2052, Australia
| | - Matthew N. Podgorski
- Department
of Chemistry, University of Adelaide, Adelaide, South Australia 5005, Australia
| | - Stephen G. Bell
- Department
of Chemistry, University of Adelaide, Adelaide, South Australia 5005, Australia
| | - Claudiu T. Supuran
- Department
of Neuroscience, Psychology, Drug Research and Child’s Health,
Section of Pharmaceutical and Nutraceutical Sciences, University of Florence, Via Ugo Schiff 6, 50019 Sesto Fiorentino, Italy
| | - William A. Donald
- School
of Chemistry, University of New South Wales, Dalton Building, Sydney, New South Wales 2052, Australia
- Phone: +61 (2) 9385 8827. E-mail:
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75
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Teo ACK, Lee SC, Pollock NL, Stroud Z, Hall S, Thakker A, Pitt AR, Dafforn TR, Spickett CM, Roper DI. Analysis of SMALP co-extracted phospholipids shows distinct membrane environments for three classes of bacterial membrane protein. Sci Rep 2019; 9:1813. [PMID: 30755655 PMCID: PMC6372662 DOI: 10.1038/s41598-018-37962-0] [Citation(s) in RCA: 52] [Impact Index Per Article: 10.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/06/2018] [Accepted: 12/05/2018] [Indexed: 11/24/2022] Open
Abstract
Biological characterisation of membrane proteins lags behind that of soluble proteins. This reflects issues with the traditional use of detergents for extraction, as the surrounding lipids are generally lost, with adverse structural and functional consequences. In contrast, styrene maleic acid (SMA) copolymers offer a detergent-free method for biological membrane solubilisation to produce SMA-lipid particles (SMALPs) containing membrane proteins together with their surrounding lipid environment. We report the development of a reverse-phase LC-MS/MS method for bacterial phospholipids and the first comparison of the profiles of SMALP co-extracted phospholipids from three exemplar bacterial membrane proteins with different topographies: FtsA (associated membrane protein), ZipA (single transmembrane helix), and PgpB (integral membrane protein). The data showed that while SMA treatment per se did not preferentially extract specific phospholipids from the membrane, SMALP-extracted ZipA showed an enrichment in phosphatidylethanolamines and depletion in cardiolipins compared to the bulk membrane lipid. Comparison of the phospholipid profiles of the 3 SMALP-extracted proteins revealed distinct lipid compositions for each protein: ZipA and PgpB were similar, but in FtsA samples longer chain phosphatidylglycerols and phosphatidylethanolamines were more abundant. This method offers novel information on the phospholipid interactions of these membrane proteins.
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Affiliation(s)
- Alvin C K Teo
- School of Life Sciences, Gibbet Hill Road, University of Warwick, Coventry, CV4 7AL, UK
| | - Sarah C Lee
- School of Biosciences, University of Birmingham, Edgbaston, Birmingham, B15 2TT, UK
| | - Naomi L Pollock
- School of Biosciences, University of Birmingham, Edgbaston, Birmingham, B15 2TT, UK
| | - Zoe Stroud
- School of Biosciences, University of Birmingham, Edgbaston, Birmingham, B15 2TT, UK
| | - Stephen Hall
- School of Biosciences, University of Birmingham, Edgbaston, Birmingham, B15 2TT, UK
| | - Alpesh Thakker
- School of Life and Health Sciences, Aston University, Aston Triangle, Birmingham, B4 7ET, UK
| | - Andrew R Pitt
- School of Life and Health Sciences, Aston University, Aston Triangle, Birmingham, B4 7ET, UK
| | - Timothy R Dafforn
- School of Biosciences, University of Birmingham, Edgbaston, Birmingham, B15 2TT, UK
| | - Corinne M Spickett
- School of Life and Health Sciences, Aston University, Aston Triangle, Birmingham, B4 7ET, UK.
| | - David I Roper
- School of Life Sciences, Gibbet Hill Road, University of Warwick, Coventry, CV4 7AL, UK.
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76
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Abstract
Following initial discoveries of noncovalent associations surviving in the gas phase, only a few practitioners pursued this research area. Today scientists around the world are using these approaches to ascertain the heterogeneity and stoichiometry of proteins within complexes. Recent developments further highlight opportunities for studying the effects of protein glycosylation on antibody–antigen interactions and drug binding, as well as site-directed mutagenesis and posttranslational modification on membrane protein interfaces. As a result of many developments over the last two decades, mass spectrometry of protein complexes has exploded and is now undertaken not just in dedicated research laboratories in academia, but also in pharmaceutical and biotechnology companies. It is therefore timely to trace the history of these developments in this personal perspective. In this Inaugural Article, I trace some key steps that have enabled the development of mass spectrometry for the study of intact protein complexes from a variety of cellular environments. Beginning with the preservation of the first soluble complexes from plasma, I describe our early experiments that capitalize on the heterogeneity of subunit composition during assembly and exchange reactions. During these investigations, we observed many assemblies and intermediates with different subunit stoichiometries, and were keen to ascertain whether or not their overall topology was preserved in the mass spectrometer. Adapting ion mobility and soft-landing methodologies, we showed how ring-shaped complexes could survive the phase transition. The next logical progression from soluble complexes was to membrane protein assemblies but this was not straightforward. We encountered many pitfalls along the way, largely due to the use of detergent micelles to protect and stabilize complexes. Further obstacles presented when we attempted to distinguish lipids that copurify from those that are important for function. Developing new experimental protocols, we have subsequently defined lipids that change protein conformation, mediate oligomeric states, and facilitate downstream coupling of G protein-coupled receptors. Very recently, using a radical method—ejecting protein complexes directly from native membranes into mass spectrometers—we provided insights into associations within membranes and mitochondria. Together, these developments suggest the beginnings of mass spectrometry meeting with cell biology.
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77
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Sharp L, Salari R, Brannigan G. Boundary lipids of the nicotinic acetylcholine receptor: Spontaneous partitioning via coarse-grained molecular dynamics simulation. BIOCHIMICA ET BIOPHYSICA ACTA-BIOMEMBRANES 2019; 1861:887-896. [PMID: 30664881 DOI: 10.1016/j.bbamem.2019.01.005] [Citation(s) in RCA: 15] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 07/26/2018] [Revised: 01/10/2019] [Accepted: 01/11/2019] [Indexed: 12/20/2022]
Abstract
Reconstituted nicotinic acetylcholine receptors (nAChRs) exhibit significant gain-of-function upon addition of cholesterol to reconstitution mixtures, and cholesterol affects the organization of nAChRs within domain-forming membranes, but whether nAChR partitions to cholesterol-rich liquid-ordered ("raft" or lo) domains or cholesterol-poor liquid-disordered (ldo) domains is unknown. We use coarse-grained molecular dynamics simulations to observe spontaneous interactions of cholesterol, saturated lipids, and polyunsaturated (PUFA) lipids with nAChRs. In binary Dipalmitoylphosphatidylcholine:Cholesterol (DPPC:CHOL) mixtures, both CHOL and DPPC acyl chains were observed spontaneously entering deep "non-annular" cavities in the nAChR TMD, particularly at the subunit interface and the β subunit center, facilitated by the low amino acid density in the cryo-EM structure of nAChR in a native membrane. Cholesterol was highly enriched in the annulus around the TMD, but this effect extended over (at most) 5-10 Å. In domain-forming ternary mixtures containing PUFAs, the presence of a single receptor did not significantly affect the likelihood of domain formation. nAChR partitioned to any cholesterol-poor ldo domain that was present, regardless of whether the ldo or lo domain lipids had PC or PE headgroups. Enrichment of PUFAs among boundary lipids was positively correlated with their propensity for demixing from cholesterol-rich phases. Long n-3 chains (tested here with Docosahexaenoic Acid, DHA) were highly enriched in annular and non-annular embedded sites, partially displacing cholesterol and completely displacing DPPC, and occupying sites even deeper within the bundle. Shorter n-6 chains were far less effective at displacing cholesterol from non-annular sites.
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Affiliation(s)
- Liam Sharp
- Center for Computational and Integrative Biology, Rutgers University-Camden, Camden, NJ, United States of America
| | - Reza Salari
- Center for Computational and Integrative Biology, Rutgers University-Camden, Camden, NJ, United States of America
| | - Grace Brannigan
- Center for Computational and Integrative Biology, Rutgers University-Camden, Camden, NJ, United States of America; Department of Physics, Rutgers University-Camden, Camden, NJ, United States of America.
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78
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Structural mass spectrometry comes of age: new insight into protein structure, function and interactions. Biochem Soc Trans 2019; 47:317-327. [DOI: 10.1042/bst20180356] [Citation(s) in RCA: 25] [Impact Index Per Article: 5.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/13/2018] [Revised: 11/27/2018] [Accepted: 12/04/2018] [Indexed: 12/15/2022]
Abstract
Abstract
Mass spectrometry (MS) provides an impressive array of information about the structure, function and interactions of proteins. In recent years, many new developments have been in the field of native MS and these exemplify a new coming of age of this field. In this mini review, we connect the latest methodological and instrumental developments in native MS to the new insights these have enabled. We highlight the prominence of an increasingly common strategy of using hybrid approaches, where multiple MS-based techniques are used in combination, and integrative approaches, where MS is used alongside other techniques such as ion-mobility spectrometry. We also review how the emergence of a native top-down approach, which combines native MS with top-down proteomics into a single experiment, is the pièce de résistance of structural mass spectrometry's coming of age. Finally, we outline key developments that have enabled membrane protein native MS to shift from being extremely challenging to routine, and how this technique is uncovering inaccessible details of membrane protein–lipid interactions.
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79
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Keener JE, Zambrano DE, Zhang G, Zak CK, Reid DJ, Deodhar BS, Pemberton JE, Prell JS, Marty MT. Chemical Additives Enable Native Mass Spectrometry Measurement of Membrane Protein Oligomeric State within Intact Nanodiscs. J Am Chem Soc 2019; 141:1054-1061. [PMID: 30586296 DOI: 10.1021/jacs.8b11529] [Citation(s) in RCA: 61] [Impact Index Per Article: 12.2] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/18/2023]
Abstract
Membrane proteins play critical biochemical roles but remain challenging to study. Recently, native or nondenaturing mass spectrometry (MS) has made great strides in characterizing membrane protein interactions. However, conventional native MS relies on detergent micelles, which may disrupt natural interactions. Lipoprotein nanodiscs provide a platform to present membrane proteins for native MS within a lipid bilayer environment, but previous native MS of membrane proteins in nanodiscs has been limited by the intermediate stability of nanodiscs. It is difficult to eject membrane proteins from nanodiscs for native MS but also difficult to retain intact nanodisc complexes with membrane proteins inside. Here, we employed chemical reagents that modulate the charge acquired during electrospray ionization (ESI). By modulating ESI conditions, we could either eject the membrane protein complex with few bound lipids or capture the intact membrane protein nanodisc complex-allowing measurement of the membrane protein oligomeric state within an intact lipid bilayer environment. The dramatic differences in the stability of nanodiscs under different ESI conditions opens new applications for native MS of nanodiscs.
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Affiliation(s)
- James E Keener
- Department of Chemistry and Biochemistry , University of Arizona , Tucson , Arizona 85721 , United States
| | - Dane Evan Zambrano
- Department of Chemistry and Biochemistry , University of Arizona , Tucson , Arizona 85721 , United States
| | - Guozhi Zhang
- Department of Chemistry and Biochemistry , University of Arizona , Tucson , Arizona 85721 , United States
| | - Ciara K Zak
- Department of Chemistry and Biochemistry , University of Arizona , Tucson , Arizona 85721 , United States
| | - Deseree J Reid
- Department of Chemistry and Biochemistry , University of Arizona , Tucson , Arizona 85721 , United States
| | - Bhushan S Deodhar
- Department of Chemistry and Biochemistry , University of Arizona , Tucson , Arizona 85721 , United States
| | - Jeanne E Pemberton
- Department of Chemistry and Biochemistry , University of Arizona , Tucson , Arizona 85721 , United States
| | - James S Prell
- Department of Chemistry and Biochemistry , University of Oregon , Eugene , Oregon 97403 , United States
| | - Michael T Marty
- Department of Chemistry and Biochemistry , University of Arizona , Tucson , Arizona 85721 , United States
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80
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Kaur U, Johnson DT, Chea EE, Deredge DJ, Espino JA, Jones LM. Evolution of Structural Biology through the Lens of Mass Spectrometry. Anal Chem 2019; 91:142-155. [PMID: 30457831 PMCID: PMC6472977 DOI: 10.1021/acs.analchem.8b05014] [Citation(s) in RCA: 39] [Impact Index Per Article: 7.8] [Reference Citation Analysis] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/01/2023]
Affiliation(s)
- Upneet Kaur
- Department of Pharmaceutical Sciences, University of Maryland, Baltimore, Maryland 21201, United States
| | - Danté T. Johnson
- Department of Pharmaceutical Sciences, University of Maryland, Baltimore, Maryland 21201, United States
| | - Emily E. Chea
- Department of Pharmaceutical Sciences, University of Maryland, Baltimore, Maryland 21201, United States
| | - Daniel J. Deredge
- Department of Pharmaceutical Sciences, University of Maryland, Baltimore, Maryland 21201, United States
| | - Jessica A. Espino
- Department of Pharmaceutical Sciences, University of Maryland, Baltimore, Maryland 21201, United States
| | - Lisa M. Jones
- Department of Pharmaceutical Sciences, University of Maryland, Baltimore, Maryland 21201, United States
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81
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Stiving AQ, VanAernum ZL, Busch F, Harvey SR, Sarni SH, Wysocki VH. Surface-Induced Dissociation: An Effective Method for Characterization of Protein Quaternary Structure. Anal Chem 2019; 91:190-209. [PMID: 30412666 PMCID: PMC6571034 DOI: 10.1021/acs.analchem.8b05071] [Citation(s) in RCA: 61] [Impact Index Per Article: 12.2] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/01/2023]
Affiliation(s)
- Alyssa Q. Stiving
- Department of Chemistry and Biochemistry and Resource for Native Mass Spectrometry Guided Structural Biology, The Ohio State University, Columbus, OH 43210
| | - Zachary L. VanAernum
- Department of Chemistry and Biochemistry and Resource for Native Mass Spectrometry Guided Structural Biology, The Ohio State University, Columbus, OH 43210
| | - Florian Busch
- Department of Chemistry and Biochemistry and Resource for Native Mass Spectrometry Guided Structural Biology, The Ohio State University, Columbus, OH 43210
- Campus Chemical Instrument Center, The Ohio State University, Columbus, OH 43210
| | - Sophie R. Harvey
- Department of Chemistry and Biochemistry and Resource for Native Mass Spectrometry Guided Structural Biology, The Ohio State University, Columbus, OH 43210
- Campus Chemical Instrument Center, The Ohio State University, Columbus, OH 43210
| | - Samantha H. Sarni
- Department of Chemistry and Biochemistry and Resource for Native Mass Spectrometry Guided Structural Biology, The Ohio State University, Columbus, OH 43210
- Ohio State Biochemistry Program, The Ohio State University, Columbus, OH 43210
- The Center for RNA Biology, The Ohio State University, Columbus, OH 43210
| | - Vicki H. Wysocki
- Department of Chemistry and Biochemistry and Resource for Native Mass Spectrometry Guided Structural Biology, The Ohio State University, Columbus, OH 43210
- Campus Chemical Instrument Center, The Ohio State University, Columbus, OH 43210
- The Center for RNA Biology, The Ohio State University, Columbus, OH 43210
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82
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Kaur U, Johnson DT, Chea EE, Deredge DJ, Espino JA, Jones LM. Evolution of Structural Biology through the Lens of Mass Spectrometry. Anal Chem 2019; 91:142-155. [PMID: 30457831 DOI: 10.1021/acs.analchem.1028b05014] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.2] [Reference Citation Analysis] [MESH Headings] [Grants] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 05/26/2023]
Affiliation(s)
- Upneet Kaur
- Department of Pharmaceutical Sciences , University of Maryland , Baltimore , Maryland 21201 , United States
| | - Danté T Johnson
- Department of Pharmaceutical Sciences , University of Maryland , Baltimore , Maryland 21201 , United States
| | - Emily E Chea
- Department of Pharmaceutical Sciences , University of Maryland , Baltimore , Maryland 21201 , United States
| | - Daniel J Deredge
- Department of Pharmaceutical Sciences , University of Maryland , Baltimore , Maryland 21201 , United States
| | - Jessica A Espino
- Department of Pharmaceutical Sciences , University of Maryland , Baltimore , Maryland 21201 , United States
| | - Lisa M Jones
- Department of Pharmaceutical Sciences , University of Maryland , Baltimore , Maryland 21201 , United States
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83
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Ion Mobility in Structural Biology. ADVANCES IN ION MOBILITY-MASS SPECTROMETRY: FUNDAMENTALS, INSTRUMENTATION AND APPLICATIONS 2019. [DOI: 10.1016/bs.coac.2018.10.001] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 12/15/2022]
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84
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Membrane protein nanoparticles: the shape of things to come. Biochem Soc Trans 2018; 46:1495-1504. [PMID: 30464048 PMCID: PMC6299238 DOI: 10.1042/bst20180139] [Citation(s) in RCA: 41] [Impact Index Per Article: 6.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/10/2018] [Revised: 09/23/2018] [Accepted: 10/02/2018] [Indexed: 12/11/2022]
Abstract
The use of styrene–maleic acid (SMA) for the purification of a wide range of membrane proteins (MPs) from both prokaryotic and eukaryotic sources has begun to make an impact in the field of MP biology. This method is growing in popularity as a means to purify and thoroughly investigate the structure and function of MPs and biological membranes. The amphiphilic SMA copolymer can effectively extract MPs directly from a native lipid bilayer to form discs ∼10 nm in diameter. The resulting lipid particles, or styrene–maleic acid lipid particles (SMALPs), contain SMA, protein and membrane lipid. MPs purified in SMALPs are able to retain their native structure and, in many cases, functional activity, and growing evidence suggests that MPs purified using SMA have enhanced thermal stability compared with detergent-purified proteins. The SMALP method is versatile and is compatible with a wide range of cell types across taxonomic domains. It can readily be adapted to replace detergent in many protein purification methods, often with only minor changes made to the existing protocol. Moreover, biophysical analysis and structural determination may now be a possibility for many large, unstable MPs. Here, we review recent advances in the area of SMALP purification and how it is affecting the field of MP biology, critically assess recent progress made with this method, address some of the associated technical challenges which may remain unresolved and discuss opportunities for exploiting SMALPs to expand our understanding of structural and functional properties of MPs.
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