1
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Nordahl L, Akkuratov EE, Heimgärtner J, Schach K, Meineke B, Elsässer S, Wennmalm S, Brismar H. Detection and quantification of Na,K-ATPase dimers in the plasma membrane of living cells by FRET-FCS. Biochim Biophys Acta Gen Subj 2024; 1868:130619. [PMID: 38643888 DOI: 10.1016/j.bbagen.2024.130619] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/04/2024] [Revised: 03/25/2024] [Accepted: 04/12/2024] [Indexed: 04/23/2024]
Abstract
The sodium potassium pump, Na,K-ATPase (NKA), is an integral plasma membrane protein, expressed in all eukaryotic cells. It is responsible for maintaining the transmembrane Na+ gradient and is the major determinant of the membrane potential. Self-interaction and oligomerization of NKA in cell membranes has been proposed and discussed but is still an open question. Here, we have used a combination of FRET and Fluorescence Correlation Spectroscopy, FRET-FCS, to analyze NKA in the plasma membrane of living cells. Click chemistry was used to conjugate the fluorescent labels Alexa 488 and Alexa 647 to non-canonical amino acids introduced in the NKA α1 and β1 subunits. We demonstrate that FRET-FCS can detect an order of magnitude lower concentration of green-red labeled protein pairs in a single-labeled red and green background than what is possible with cross-correlation (FCCS). We show that a significant fraction of NKA is expressed as a dimer in the plasma membrane. We also introduce a method to estimate not only the number of single and double labeled NKA, but the number of unlabeled, endogenous NKA and estimate the density of endogenous NKA at the plasma membrane to 1400 ± 800 enzymes/μm2.
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Affiliation(s)
- Linnea Nordahl
- Science for Life Laboratory, Department of Applied Physics, KTH Royal Institute of Technology, Box 1031, 171 21 Solna, Sweden
| | - Evgeny E Akkuratov
- University of Oxford, MRC Weatherall Institute of Molecular Medicine, Radcliffe Department of Medicine, Oxford OX3 9DS, UK
| | - Johannes Heimgärtner
- Science for Life Laboratory, Department of Medical Biochemistry and Biophysics, Karolinska Institutet, Box 1031, 171 21 Solna, Sweden
| | - Katja Schach
- Graduate Training Centre of Neuroscience, University of Tübingen, Otfried-Müller-Str. 27, 72076 Tübingen, Germany
| | - Birthe Meineke
- Science for Life Laboratory, Department of Medical Biochemistry and Biophysics, Karolinska Institutet, Box 1031, 171 21 Solna, Sweden
| | - Simon Elsässer
- Science for Life Laboratory, Department of Medical Biochemistry and Biophysics, Karolinska Institutet, Box 1031, 171 21 Solna, Sweden
| | - Stefan Wennmalm
- Science for Life Laboratory, Department of Applied Physics, KTH Royal Institute of Technology, Box 1031, 171 21 Solna, Sweden.
| | - Hjalmar Brismar
- Science for Life Laboratory, Department of Applied Physics, KTH Royal Institute of Technology, Box 1031, 171 21 Solna, Sweden; Science for Life Laboratory, Department of Women's and Children's Health, Karolinska Institutet, Box 1031, 171 21 Solna, Sweden
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2
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Jahn R, Cafiso DC, Tamm LK. Mechanisms of SNARE proteins in membrane fusion. Nat Rev Mol Cell Biol 2024; 25:101-118. [PMID: 37848589 DOI: 10.1038/s41580-023-00668-x] [Citation(s) in RCA: 4] [Impact Index Per Article: 4.0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Accepted: 09/06/2023] [Indexed: 10/19/2023]
Abstract
Soluble N-ethylmaleimide-sensitive factor attachment protein receptors (SNAREs) are a family of small conserved eukaryotic proteins that mediate membrane fusion between organelles and with the plasma membrane. SNAREs are directly or indirectly anchored to membranes. Prior to fusion, complementary SNAREs assemble between membranes with the aid of accessory proteins that provide a scaffold to initiate SNARE zippering, pulling the membranes together and mediating fusion. Recent advances have enabled the construction of detailed models describing bilayer transitions and energy barriers along the fusion pathway and have elucidated the structures of SNAREs complexed in various states with regulatory proteins. In this Review, we discuss how these advances are yielding an increasingly detailed picture of the SNARE-mediated fusion pathway, leading from first contact between the membranes via metastable non-bilayer intermediates towards the opening and expansion of a fusion pore. We describe how SNARE proteins assemble into complexes, how this assembly is regulated by accessory proteins and how SNARE complexes overcome the free energy barriers that prevent spontaneous membrane fusion.
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Affiliation(s)
- Reinhard Jahn
- Laboratory of Neurobiology, Max-Planck Institute for Multidisciplinary Sciences, Göttingen, Germany.
| | - David C Cafiso
- Department of Chemistry, University of Virginia, Charlottesville, VA, USA
| | - Lukas K Tamm
- Department of Molecular Physiology and Biological Physics, University of Virginia, Charlottesville, VA, USA
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3
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Stefani I, Iwaszkiewicz J, Fasshauer D. Exploring the conformational changes of the Munc18-1/syntaxin 1a complex. Protein Sci 2023; 33:e4870. [PMID: 38109275 PMCID: PMC10895456 DOI: 10.1002/pro.4870] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/13/2023] [Revised: 11/30/2023] [Accepted: 12/07/2023] [Indexed: 12/20/2023]
Abstract
Neurotransmitters are released from synaptic vesicles, the membrane of which fuses with the plasma membrane upon calcium influx. This membrane fusion reaction is driven by the formation of a tight complex comprising the plasma membrane N-ethylmaleimide-sensitive factor attachment receptor (SNARE) proteins syntaxin-1a and SNAP-25 with the vesicle SNARE protein synaptobrevin. The neuronal protein Munc18-1 forms a stable complex with syntaxin-1a. Biochemically, syntaxin-1a cannot escape the tight grip of Munc18-1, so formation of the SNARE complex is inhibited. However, Munc18-1 is essential for the release of neurotransmitters in vivo. It has therefore been assumed that Munc18-1 makes the bound syntaxin-1a available for SNARE complex formation. Exactly how this occurs is still unclear, but it is assumed that structural rearrangements occur. Here, we used a series of mutations to specifically weaken the complex at different positions in order to induce these rearrangements biochemically. Our approach was guided through sequence and structural analysis and supported by molecular dynamics simulations. Subsequently, we created a homology model showing the complex in an altered conformation. This conformation presumably represents a more open arrangement of syntaxin-1a that permits the formation of a SNARE complex to be initiated while still bound to Munc18-1. In the future, research should investigate how this central reaction for neuronal communication is controlled by other proteins.
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Affiliation(s)
- Ioanna Stefani
- Department of Computational BiologyUniversity of LausanneLausanneSwitzerland
| | | | - Dirk Fasshauer
- Department of Computational BiologyUniversity of LausanneLausanneSwitzerland
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4
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Holden MR, Krzesinski BJ, Weismiller HA, Shady JR, Margittai M. MAP2 caps tau fibrils and inhibits aggregation. J Biol Chem 2023; 299:104891. [PMID: 37286038 PMCID: PMC10404690 DOI: 10.1016/j.jbc.2023.104891] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/05/2023] [Revised: 05/13/2023] [Accepted: 05/30/2023] [Indexed: 06/09/2023] Open
Abstract
Fibrils of the microtubule-associated protein tau are intimately linked to the pathology of Alzheimer's disease (AD) and related neurodegenerative disorders. A current paradigm for pathology spreading in the human brain is that short tau fibrils transfer between neurons and then recruit naive tau monomers onto their tips, perpetuating the fibrillar conformation with high fidelity and speed. Although it is known that the propagation could be modulated in a cell-specific manner and thereby contribute to phenotypic diversity, there is still limited understanding of how select molecules are involved in this process. MAP2 is a neuronal protein that shares significant sequence homology with the repeat-bearing amyloid core region of tau. There is discrepancy about MAP2's involvement in pathology and its relationship with tau fibrillization. Here, we employed the entire repeat regions of 3R and 4R MAP2, to investigate their modulatory role in tau fibrillization. We find that both proteins block the spontaneous and seeded aggregation of 4R tau, with 4R MAP2 being slightly more potent. The inhibition of tau seeding is observed in vitro, in HEK293 cells, and in AD brain extracts, underscoring its broader scope. MAP2 monomers specifically bind to the end of tau fibrils, preventing recruitment of further tau and MAP2 monomers onto the fibril tip. The findings uncover a new function for MAP2 as a tau fibril cap that could play a significant role in modulating tau propagation in disease and may hold promise as a potential intrinsic protein inhibitor.
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Affiliation(s)
- Michael R Holden
- Department of Chemistry and Biochemistry, University of Denver, Denver, Colorado, USA
| | - Brad J Krzesinski
- Department of Chemistry and Biochemistry, University of Denver, Denver, Colorado, USA
| | - Hilary A Weismiller
- Department of Chemistry and Biochemistry, University of Denver, Denver, Colorado, USA
| | - Justin R Shady
- Department of Chemistry and Biochemistry, University of Denver, Denver, Colorado, USA
| | - Martin Margittai
- Department of Chemistry and Biochemistry, University of Denver, Denver, Colorado, USA.
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5
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Agam G, Gebhardt C, Popara M, Mächtel R, Folz J, Ambrose B, Chamachi N, Chung SY, Craggs TD, de Boer M, Grohmann D, Ha T, Hartmann A, Hendrix J, Hirschfeld V, Hübner CG, Hugel T, Kammerer D, Kang HS, Kapanidis AN, Krainer G, Kramm K, Lemke EA, Lerner E, Margeat E, Martens K, Michaelis J, Mitra J, Moya Muñoz GG, Quast RB, Robb NC, Sattler M, Schlierf M, Schneider J, Schröder T, Sefer A, Tan PS, Thurn J, Tinnefeld P, van Noort J, Weiss S, Wendler N, Zijlstra N, Barth A, Seidel CAM, Lamb DC, Cordes T. Reliability and accuracy of single-molecule FRET studies for characterization of structural dynamics and distances in proteins. Nat Methods 2023; 20:523-535. [PMID: 36973549 PMCID: PMC10089922 DOI: 10.1038/s41592-023-01807-0] [Citation(s) in RCA: 19] [Impact Index Per Article: 19.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/02/2022] [Accepted: 01/31/2023] [Indexed: 03/29/2023]
Abstract
Single-molecule Förster-resonance energy transfer (smFRET) experiments allow the study of biomolecular structure and dynamics in vitro and in vivo. We performed an international blind study involving 19 laboratories to assess the uncertainty of FRET experiments for proteins with respect to the measured FRET efficiency histograms, determination of distances, and the detection and quantification of structural dynamics. Using two protein systems with distinct conformational changes and dynamics, we obtained an uncertainty of the FRET efficiency ≤0.06, corresponding to an interdye distance precision of ≤2 Å and accuracy of ≤5 Å. We further discuss the limits for detecting fluctuations in this distance range and how to identify dye perturbations. Our work demonstrates the ability of smFRET experiments to simultaneously measure distances and avoid the averaging of conformational dynamics for realistic protein systems, highlighting its importance in the expanding toolbox of integrative structural biology.
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Affiliation(s)
- Ganesh Agam
- Department of Chemistry, Ludwig-Maximilians University München, München, Germany
| | - Christian Gebhardt
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians University München, Planegg-Martinsried, Germany
| | - Milana Popara
- Molecular Physical Chemistry, Heinrich-Heine University Düsseldorf, Düsseldorf, Germany
| | - Rebecca Mächtel
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians University München, Planegg-Martinsried, Germany
| | - Julian Folz
- Molecular Physical Chemistry, Heinrich-Heine University Düsseldorf, Düsseldorf, Germany
| | | | - Neharika Chamachi
- B CUBE - Center for Molecular Bioengineering, Technische Universität Dresden, Dresden, Germany
| | - Sang Yoon Chung
- Department of Chemistry and Biochemistry, University of California, Los Angeles, CA, USA
| | | | - Marijn de Boer
- Molecular Microscopy Research Group, Zernike Institute for Advanced Materials, University of Groningen, AG Groningen, the Netherlands
| | - Dina Grohmann
- Department of Biochemistry, Genetics and Microbiology, Institute of Microbiology, Single-Molecule Biochemistry Laboratory, University of Regensburg, Regensburg, Germany
| | - Taekjip Ha
- Department of Biophysics and Biophysical Chemistry, Johns Hopkins University School of Medicine and Howard Hughes Medical Institute, Baltimore, MD, USA
| | - Andreas Hartmann
- B CUBE - Center for Molecular Bioengineering, Technische Universität Dresden, Dresden, Germany
| | - Jelle Hendrix
- Dynamic Bioimaging Laboratory, Advanced Optical Microscopy Center and Biomedical Research Institute, Hasselt University, Agoralaan C (BIOMED), Hasselt, Belgium
- Department of Chemistry, KU Leuven, Leuven, Belgium
| | | | | | - Thorsten Hugel
- Institute of Physical Chemistry, University of Freiburg, Freiburg, Germany
- Signalling Research Centers BIOSS and CIBSS, University of Freiburg, Freiburg, Germany
| | - Dominik Kammerer
- Department of Physics, Clarendon Laboratory, University of Oxford, Oxford, UK
- Kavli Institute of Nanoscience Discovery, University of Oxford, Oxford, UK
| | - Hyun-Seo Kang
- Bayerisches NMR Zentrum, Department of Bioscience, School of Natural Sciences, Technical University of München, Garching, Germany
| | - Achillefs N Kapanidis
- Department of Physics, Clarendon Laboratory, University of Oxford, Oxford, UK
- Kavli Institute of Nanoscience Discovery, University of Oxford, Oxford, UK
| | - Georg Krainer
- B CUBE - Center for Molecular Bioengineering, Technische Universität Dresden, Dresden, Germany
- Yusuf Hamied Department of Chemistry, University of Cambridge, Cambridge, UK
| | - Kevin Kramm
- Department of Biochemistry, Genetics and Microbiology, Institute of Microbiology, Single-Molecule Biochemistry Laboratory, University of Regensburg, Regensburg, Germany
| | - Edward A Lemke
- Biocenter, Johannes Gutenberg University Mainz, Mainz, Germany
- Institute of Molecular Biology, Mainz, Germany
- Structural and Computational Biology Unit, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Eitan Lerner
- Department of Biological Chemistry, The Alexander Silberman Institute of Life Sciences, and The Center for Nanoscience and Nanotechnology, Faculty of Mathematics and Science, The Edmond J. Safra Campus, The Hebrew University of Jerusalem, Jerusalem, Israel
| | - Emmanuel Margeat
- Centre de Biologie Structurale (CBS), University of Montpellier, CNRS, INSERM, Montpellier, France
| | - Kirsten Martens
- Biological and Soft Matter Physics, Huygens-Kamerlingh Onnes Laboratory, Leiden University, Leiden, the Netherlands
| | | | - Jaba Mitra
- Department of Biophysics and Biophysical Chemistry, Johns Hopkins University School of Medicine and Howard Hughes Medical Institute, Baltimore, MD, USA
- Materials Science and Engineering, University of Illinois Urbana-Champaign, Urbana, IL, USA
| | - Gabriel G Moya Muñoz
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians University München, Planegg-Martinsried, Germany
| | - Robert B Quast
- Centre de Biologie Structurale (CBS), University of Montpellier, CNRS, INSERM, Montpellier, France
| | - Nicole C Robb
- Department of Physics, Clarendon Laboratory, University of Oxford, Oxford, UK
- Kavli Institute of Nanoscience Discovery, University of Oxford, Oxford, UK
- Warwick Medical School, The University of Warwick, Coventry, UK
| | - Michael Sattler
- Bayerisches NMR Zentrum, Department of Bioscience, School of Natural Sciences, Technical University of München, Garching, Germany
- Institute of Structural Biology, Molecular Targets and Therapeutics Center, Helmholtz Center Munich, Munich, Germany
| | - Michael Schlierf
- B CUBE - Center for Molecular Bioengineering, Technische Universität Dresden, Dresden, Germany
- Cluster of Excellence Physics of Life, Technische Universität Dresden, Dresden, Germany
| | - Jonathan Schneider
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians University München, Planegg-Martinsried, Germany
| | - Tim Schröder
- Department of Chemistry, Ludwig-Maximilians University München, München, Germany
| | - Anna Sefer
- Institute for Biophysics, Ulm University, Ulm, Germany
| | - Piau Siong Tan
- Biocenter, Johannes Gutenberg University Mainz, Mainz, Germany
- Institute of Molecular Biology, Mainz, Germany
| | - Johann Thurn
- Institute of Physical Chemistry, University of Freiburg, Freiburg, Germany
- Institute of Technical Physics, German Aerospace Center (DLR), Stuttgart, Germany
| | - Philip Tinnefeld
- Department of Chemistry, Ludwig-Maximilians University München, München, Germany
| | - John van Noort
- Biological and Soft Matter Physics, Huygens-Kamerlingh Onnes Laboratory, Leiden University, Leiden, the Netherlands
| | - Shimon Weiss
- Department of Chemistry and Biochemistry, University of California, Los Angeles, CA, USA
- California NanoSystems Institute, University of California, Los Angeles, CA, USA
| | - Nicolas Wendler
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians University München, Planegg-Martinsried, Germany
| | - Niels Zijlstra
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians University München, Planegg-Martinsried, Germany
| | - Anders Barth
- Molecular Physical Chemistry, Heinrich-Heine University Düsseldorf, Düsseldorf, Germany.
- Department of Bionanoscience, Kavli Institute of Nanoscience, Delft University of Technology, Delft, the Netherlands.
| | - Claus A M Seidel
- Molecular Physical Chemistry, Heinrich-Heine University Düsseldorf, Düsseldorf, Germany.
| | - Don C Lamb
- Department of Chemistry, Ludwig-Maximilians University München, München, Germany.
| | - Thorben Cordes
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians University München, Planegg-Martinsried, Germany.
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6
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Illuminating Intrinsically Disordered Proteins with Integrative Structural Biology. Biomolecules 2023; 13:biom13010124. [PMID: 36671509 PMCID: PMC9856150 DOI: 10.3390/biom13010124] [Citation(s) in RCA: 3] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/01/2022] [Revised: 01/01/2023] [Accepted: 01/04/2023] [Indexed: 01/11/2023] Open
Abstract
Intense study of intrinsically disordered proteins (IDPs) did not begin in earnest until the late 1990s when a few groups, working independently, convinced the community that these 'weird' proteins could have important functions. Over the past two decades, it has become clear that IDPs play critical roles in a multitude of biological phenomena with prominent examples including coordination in signaling hubs, enabling gene regulation, and regulating ion channels, just to name a few. One contributing factor that delayed appreciation of IDP functional significance is the experimental difficulty in characterizing their dynamic conformations. The combined application of multiple methods, termed integrative structural biology, has emerged as an essential approach to understanding IDP phenomena. Here, we review some of the recent applications of the integrative structural biology philosophy to study IDPs.
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7
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Maus H, Hinze G, Hammerschmidt SJ, Basché T, Schirmeister T. A competition smFRET assay to study ligand-induced conformational changes of the dengue virus protease. Protein Sci 2023; 32:e4526. [PMID: 36461913 PMCID: PMC9793963 DOI: 10.1002/pro.4526] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/23/2022] [Revised: 11/07/2022] [Accepted: 11/29/2022] [Indexed: 12/05/2022]
Abstract
Ligand binding to proteins often is accompanied by conformational transitions. Here, we describe a competition assay based on single molecule Förster resonance energy transfer (smFRET) to investigate the ligand-induced conformational changes of the dengue virus (DENV) NS2B-NS3 protease, which can adopt at least two different conformations. First, a competitive ligand was used to stabilize the closed conformation of the protease. Subsequent addition of the allosteric inhibitor reduced the fraction of the closed conformation and simultaneously increased the fraction of the open conformation, demonstrating that the allosteric inhibitor stabilizes the open conformation. In addition, the proportions of open and closed conformations at different concentrations of the allosteric inhibitor were used to determine its binding affinity to the protease. The KD value observed is in accordance with the IC50 determined in the fluorometric assay. Our novel approach appears to be a valuable tool to study conformational transitions of other proteases and enzymes.
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Affiliation(s)
- Hannah Maus
- Institute for Pharmaceutical and Biomedical Sciences, Johannes Gutenberg‐UniversityMainzGermany
| | - Gerald Hinze
- Department of ChemistryJohannes Gutenberg‐UniversityMainzGermany
| | | | - Thomas Basché
- Department of ChemistryJohannes Gutenberg‐UniversityMainzGermany
| | - Tanja Schirmeister
- Institute for Pharmaceutical and Biomedical Sciences, Johannes Gutenberg‐UniversityMainzGermany
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8
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Jas GS, Childs EW, Middaugh CR, Kuczera K. Observing reorientation dynamics with Time-Resolved fluorescence and molecular dynamics in varying periodic boundary conditions. J Biomol Struct Dyn 2022; 40:10614-10628. [PMID: 34308794 DOI: 10.1080/07391102.2021.1947894] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 10/20/2022]
Abstract
This work presents a combined study of time-resolved fluorescence spectroscopy and all-atom molecular dynamics simulation to investigate periodic boundary conditions' and water models' influence on the orientation dynamics and translational and rotational diffusion of peptides in solution. We have characterized the effects of solvent box size and water model choice on the dynamics of two peptide systems, NATA and WK5. Computationally, translational, and rotational diffusion and internal fluctuations are investigated through all-atom molecular dynamics simulation with two water models and different box sizes. These results are compared with time-resolved fluorescence anisotropy decay (FAD) measurements. The associated time constant and orientation dynamics from FAD measurement along the 1Lb axis provided baseline data to validate molecular dynamics simulation. The modeling results show that diffusion rates vary roughly in inverse proportion to water model viscosity, as one would expect. Corrections for finite box size are significant for translational diffusion and insignificant for rotational diffusion. This study also finds that internal dynamics described by autocorrelation functions and kinetic network models are relatively insensitive to both box size and water model properties. Our observation suggests that different peptide properties respond differently to a change in simulation conditions.Communicated by Ramaswamy H. Sarma.
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Affiliation(s)
- Gouri S Jas
- Department of Pharmaceutical Chemistry, The University of Kansas, Lawrence, KS, USA
| | - Ed W Childs
- Department of Surgery, Morehouse School of Medicine, Atlanta, GA, USA
| | - C Russell Middaugh
- Department of Pharmaceutical Chemistry, The University of Kansas, Lawrence, KS, USA
| | - Krzysztof Kuczera
- Department of Chemistry and Department of Molecular Biosciences, The University of Kansas, Lawrence, KS, USA
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9
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Synaptic Secretion and Beyond: Targeting Synapse and Neurotransmitters to Treat Neurodegenerative Diseases. OXIDATIVE MEDICINE AND CELLULAR LONGEVITY 2022; 2022:9176923. [PMID: 35923862 PMCID: PMC9343216 DOI: 10.1155/2022/9176923] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Subscribe] [Scholar Register] [Received: 02/10/2022] [Revised: 05/16/2022] [Accepted: 06/04/2022] [Indexed: 11/17/2022]
Abstract
The nervous system is important, because it regulates the physiological function of the body. Neurons are the most basic structural and functional unit of the nervous system. The synapse is an asymmetric structure that is important for neuronal function. The chemical transmission mode of the synapse is realized through neurotransmitters and electrical processes. Based on vesicle transport, the abnormal information transmission process in the synapse can lead to a series of neurorelated diseases. Numerous proteins and complexes that regulate the process of vesicle transport, such as SNARE proteins, Munc18-1, and Synaptotagmin-1, have been identified. Their regulation of synaptic vesicle secretion is complicated and delicate, and their defects can lead to a series of neurodegenerative diseases. This review will discuss the structure and functions of vesicle-based synapses and their roles in neurons. Furthermore, we will analyze neurotransmitter and synaptic functions in neurodegenerative diseases and discuss the potential of using related drugs in their treatment.
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10
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Opanasyuk O, Barth A, Peulen TO, Felekyan S, Kalinin S, Sanabria H, Seidel CA. Unraveling multi-state molecular dynamics in single-molecule FRET experiments. II. Quantitative analysis of multi-state kinetic networks. J Chem Phys 2022; 157:031501. [DOI: 10.1063/5.0095754] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/14/2022] Open
Abstract
Single-molecule Förster Resonance Energy Transfer (smFRET) experiments are ideally suited to resolve the structural dynamics of biomolecules. A significant challenge to date is capturing and quantifying the exchange between multiple conformational states, mainly when these dynamics occur on the sub-millisecond timescale. Many methods for quantitative analysis are challenged if more than two states are involved, and the appropriate choice of the number of states in the kinetic network is difficult. An additional complication arises if dynamically active molecules coexist with pseudo-static molecules in similar conformational states with undistinguishable FRET efficiencies. To address these problems, we developed a quantitative integrative analysis framework that combines the information from FRET-lines that relate average fluorescence lifetimes and intensities in two-dimensional burst frequency histograms, fluorescence decays obtained by time-correlated single photon counting, photon distribution analysis of the intensities and fluorescence correlation spectroscopy. Individually, these methodologies provide ambiguous results for the characterization of dynamics in complex kinetic networks. However, the global analysis approach enables accurate determination of the number of states, their kinetic connectivity, the transition rate constants, and species fractions. To challenge the potential of smFRET experiments studying multi-state kinetic networks, we apply our integrative framework using a set of synthetic data for three-state systems with different kinetic connectivity and exchange rates. Our methodology paves the way towards an integrated analysis of multiparameter smFRET experiments that spans all dimensions of the experimental data. Finally, we propose a workflow for the analysis and show examples that demonstrate the usefulness of this toolkit for dynamic structural biology.
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Affiliation(s)
| | | | | | - Suren Felekyan
- PC-II, Heinrich Heine University Düsseldorf Department of Chemistry, Germany
| | - Stanislav Kalinin
- Institut für Physikalische Chemie, Heinrich-Heine-Universität Düsseldorf, Germany
| | | | - Claus A.M. Seidel
- Institut fuer Physikalische Chemie, Heinrich-Heine-Universität Düsseldorf, Germany
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11
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Gething C, Ferrar J, Misra B, Howells G, Andrzejewski AL, Bowen ME, Choi UB. Conformational change of Syntaxin-3b in regulating SNARE complex assembly in the ribbon synapses. Sci Rep 2022; 12:9261. [PMID: 35661757 PMCID: PMC9166750 DOI: 10.1038/s41598-022-09654-3] [Citation(s) in RCA: 2] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/07/2021] [Accepted: 03/10/2022] [Indexed: 11/09/2022] Open
Abstract
Neurotransmitter release of synaptic vesicles relies on the assembly of the soluble N-ethylmaleimide-sensitive factor attachment protein receptor (SNARE) complex, consisting of syntaxin and SNAP-25 on the plasma membrane and synaptobrevin on the synaptic vesicle. The formation of the SNARE complex progressively zippers towards the membranes, which drives membrane fusion between the plasma membrane and the synaptic vesicle. However, the underlying molecular mechanism of SNARE complex regulation is unclear. In this study, we investigated the syntaxin-3b isoform found in the retinal ribbon synapses using single-molecule fluorescence resonance energy transfer (smFRET) to monitor the conformational changes of syntaxin-3b that modulate the SNARE complex formation. We found that syntaxin-3b is predominantly in a self-inhibiting closed conformation, inefficiently forming the ternary SNARE complex. Conversely, a phosphomimetic mutation (T14E) at the N-terminal region of syntaxin-3b promoted the open conformation, similar to the constitutively open form of syntaxin LE mutant. When syntaxin-3b is bound to Munc18-1, SNARE complex formation is almost completely blocked. Surprisingly, the T14E mutation of syntaxin-3b partially abolishes Munc18-1 regulation, acting as a conformational switch to trigger SNARE complex assembly. Thus, we suggest a model where the conformational change of syntaxin-3b induced by phosphorylation initiates the release of neurotransmitters in the ribbon synapses.
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Affiliation(s)
- Claire Gething
- Department of Biochemistry, West Virginia University, Morgantown, WV, 26506, USA
| | - Joshua Ferrar
- Department of Biochemistry, West Virginia University, Morgantown, WV, 26506, USA
| | - Bishal Misra
- Department of Biochemistry, West Virginia University, Morgantown, WV, 26506, USA
| | - Giovanni Howells
- Department of Biochemistry, West Virginia University, Morgantown, WV, 26506, USA
| | | | - Mark E Bowen
- Department of Physiology and Biophysics, Stony Brook University, Stony Brook, NY, 11794, USA.,Quantum-Si, Inc, Guilford, CT, 06437, USA
| | - Ucheor B Choi
- Department of Biochemistry, West Virginia University, Morgantown, WV, 26506, USA. .,Quantum-Si, Inc, Guilford, CT, 06437, USA.
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12
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Barth A, Opanasyuk O, Peulen TO, Felekyan S, Kalinin S, Sanabria H, Seidel CAM. Unraveling multi-state molecular dynamics in single-molecule FRET experiments. I. Theory of FRET-lines. J Chem Phys 2022; 156:141501. [PMID: 35428384 PMCID: PMC9014241 DOI: 10.1063/5.0089134] [Citation(s) in RCA: 14] [Impact Index Per Article: 7.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/31/2023] Open
Abstract
Conformational dynamics of biomolecules are of fundamental importance for their function. Single-molecule studies of Förster Resonance Energy Transfer (smFRET) between a tethered donor and acceptor dye pair are a powerful tool to investigate the structure and dynamics of labeled molecules. However, capturing and quantifying conformational dynamics in intensity-based smFRET experiments remains challenging when the dynamics occur on the sub-millisecond timescale. The method of multiparameter fluorescence detection addresses this challenge by simultaneously registering fluorescence intensities and lifetimes of the donor and acceptor. Together, two FRET observables, the donor fluorescence lifetime τD and the intensity-based FRET efficiency E, inform on the width of the FRET efficiency distribution as a characteristic fingerprint for conformational dynamics. We present a general framework for analyzing dynamics that relates average fluorescence lifetimes and intensities in two-dimensional burst frequency histograms. We present parametric relations of these observables for interpreting the location of FRET populations in E–τD diagrams, called FRET-lines. To facilitate the analysis of complex exchange equilibria, FRET-lines serve as reference curves for a graphical interpretation of experimental data to (i) identify conformational states, (ii) resolve their dynamic connectivity, (iii) compare different kinetic models, and (iv) infer polymer properties of unfolded or intrinsically disordered proteins. For a simplified graphical analysis of complex kinetic networks, we derive a moment-based representation of the experimental data that decouples the motion of the fluorescence labels from the conformational dynamics of the biomolecule. Importantly, FRET-lines facilitate exploring complex dynamic models via easily computed experimental observables. We provide extensive computational tools to facilitate applying FRET-lines.
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Affiliation(s)
- Anders Barth
- Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, Düsseldorf, Germany
| | - Oleg Opanasyuk
- Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, Düsseldorf, Germany
| | - Thomas-Otavio Peulen
- Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, Düsseldorf, Germany
| | - Suren Felekyan
- Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, Düsseldorf, Germany
| | - Stanislav Kalinin
- Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, Düsseldorf, Germany
| | - Hugo Sanabria
- Department of Physics and Astronomy, Clemson University, Clemson, South Carolina 29631, USA
| | - Claus A. M. Seidel
- Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, Düsseldorf, Germany
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13
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Liu Y, Pan T, Wang K, Wang Y, Yan S, Wang L, Zhang S, Du X, Jia W, Zhang P, Chen H, Huang S. Allosteric Switching of Calmodulin in a
Mycobacterium smegmatis
porin A (MspA) Nanopore‐Trap. Angew Chem Int Ed Engl 2021. [DOI: 10.1002/ange.202110545] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.7] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/11/2023]
Affiliation(s)
- Yao Liu
- State Key Laboratory of Analytical Chemistry for Life Sciences School of Chemistry and Chemical Engineering Nanjing University 210023 Nanjing China
- Chemistry and Biomedicine Innovation Center (ChemBIC) Nanjing University 210023 Nanjing China
| | - Tiezheng Pan
- School of Life Sciences Northwestern Polytechnical University 710072 Xi'an China
| | - Kefan Wang
- State Key Laboratory of Analytical Chemistry for Life Sciences School of Chemistry and Chemical Engineering Nanjing University 210023 Nanjing China
- Chemistry and Biomedicine Innovation Center (ChemBIC) Nanjing University 210023 Nanjing China
| | - Yuqin Wang
- State Key Laboratory of Analytical Chemistry for Life Sciences School of Chemistry and Chemical Engineering Nanjing University 210023 Nanjing China
- Chemistry and Biomedicine Innovation Center (ChemBIC) Nanjing University 210023 Nanjing China
| | - Shuanghong Yan
- State Key Laboratory of Analytical Chemistry for Life Sciences School of Chemistry and Chemical Engineering Nanjing University 210023 Nanjing China
- Chemistry and Biomedicine Innovation Center (ChemBIC) Nanjing University 210023 Nanjing China
| | - Liying Wang
- State Key Laboratory of Analytical Chemistry for Life Sciences School of Chemistry and Chemical Engineering Nanjing University 210023 Nanjing China
- Chemistry and Biomedicine Innovation Center (ChemBIC) Nanjing University 210023 Nanjing China
| | - Shanyu Zhang
- State Key Laboratory of Analytical Chemistry for Life Sciences School of Chemistry and Chemical Engineering Nanjing University 210023 Nanjing China
- Chemistry and Biomedicine Innovation Center (ChemBIC) Nanjing University 210023 Nanjing China
| | - Xiaoyu Du
- State Key Laboratory of Analytical Chemistry for Life Sciences School of Chemistry and Chemical Engineering Nanjing University 210023 Nanjing China
- Chemistry and Biomedicine Innovation Center (ChemBIC) Nanjing University 210023 Nanjing China
| | - Wendong Jia
- State Key Laboratory of Analytical Chemistry for Life Sciences School of Chemistry and Chemical Engineering Nanjing University 210023 Nanjing China
- Chemistry and Biomedicine Innovation Center (ChemBIC) Nanjing University 210023 Nanjing China
| | - Panke Zhang
- State Key Laboratory of Analytical Chemistry for Life Sciences School of Chemistry and Chemical Engineering Nanjing University 210023 Nanjing China
- Chemistry and Biomedicine Innovation Center (ChemBIC) Nanjing University 210023 Nanjing China
| | - Hong‐Yuan Chen
- State Key Laboratory of Analytical Chemistry for Life Sciences School of Chemistry and Chemical Engineering Nanjing University 210023 Nanjing China
- Chemistry and Biomedicine Innovation Center (ChemBIC) Nanjing University 210023 Nanjing China
| | - Shuo Huang
- State Key Laboratory of Analytical Chemistry for Life Sciences School of Chemistry and Chemical Engineering Nanjing University 210023 Nanjing China
- Chemistry and Biomedicine Innovation Center (ChemBIC) Nanjing University 210023 Nanjing China
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14
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Liu Y, Pan T, Wang K, Wang Y, Yan S, Wang L, Zhang S, Du X, Jia W, Zhang P, Chen HY, Huang S. Allosteric Switching of Calmodulin in a Mycobacterium smegmatis porin A (MspA) Nanopore-Trap. Angew Chem Int Ed Engl 2021; 60:23863-23870. [PMID: 34449124 DOI: 10.1002/anie.202110545] [Citation(s) in RCA: 8] [Impact Index Per Article: 2.7] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/06/2021] [Revised: 08/21/2021] [Indexed: 01/23/2023]
Abstract
Recent developments concerning large protein nanopores suggest a new approach to structure profiling of native folded proteins. In this work, the large vestibule of Mycobacterium smegmatis porin A (MspA) and calmodulin (CaM), a Ca2+ -binding protein, were used in the direct observation of the protein structure. Three conformers, including the Ca2+ -free, Ca2+ -bound, and target peptide-bound states of CaM, were unambiguously distinguished. A disease related mutant, CaM D129G was also discriminated by MspA, revealing how a single amino acid replacement can interfere with the Ca2+ -binding capacity of the whole protein. The binding capacity and aggregation effect of CaM induced by different ions (Mg2+ /Sr2+ /Ba2+ /Ca2+ /Pb2+ /Tb3+ ) were also investigated and the stability of MspA in extreme conditions was evaluated. This work demonstrates the most systematic single-molecule investigation of different allosteric conformers of CaM, acknowledging the high sensing resolution offered by the MspA nanopore trap.
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Affiliation(s)
- Yao Liu
- State Key Laboratory of Analytical Chemistry for Life Sciences, School of Chemistry and Chemical Engineering, Nanjing University, 210023, Nanjing, China.,Chemistry and Biomedicine Innovation Center (ChemBIC), Nanjing University, 210023, Nanjing, China
| | - Tiezheng Pan
- School of Life Sciences, Northwestern Polytechnical University, 710072, Xi'an, China
| | - Kefan Wang
- State Key Laboratory of Analytical Chemistry for Life Sciences, School of Chemistry and Chemical Engineering, Nanjing University, 210023, Nanjing, China.,Chemistry and Biomedicine Innovation Center (ChemBIC), Nanjing University, 210023, Nanjing, China
| | - Yuqin Wang
- State Key Laboratory of Analytical Chemistry for Life Sciences, School of Chemistry and Chemical Engineering, Nanjing University, 210023, Nanjing, China.,Chemistry and Biomedicine Innovation Center (ChemBIC), Nanjing University, 210023, Nanjing, China
| | - Shuanghong Yan
- State Key Laboratory of Analytical Chemistry for Life Sciences, School of Chemistry and Chemical Engineering, Nanjing University, 210023, Nanjing, China.,Chemistry and Biomedicine Innovation Center (ChemBIC), Nanjing University, 210023, Nanjing, China
| | - Liying Wang
- State Key Laboratory of Analytical Chemistry for Life Sciences, School of Chemistry and Chemical Engineering, Nanjing University, 210023, Nanjing, China.,Chemistry and Biomedicine Innovation Center (ChemBIC), Nanjing University, 210023, Nanjing, China
| | - Shanyu Zhang
- State Key Laboratory of Analytical Chemistry for Life Sciences, School of Chemistry and Chemical Engineering, Nanjing University, 210023, Nanjing, China.,Chemistry and Biomedicine Innovation Center (ChemBIC), Nanjing University, 210023, Nanjing, China
| | - Xiaoyu Du
- State Key Laboratory of Analytical Chemistry for Life Sciences, School of Chemistry and Chemical Engineering, Nanjing University, 210023, Nanjing, China.,Chemistry and Biomedicine Innovation Center (ChemBIC), Nanjing University, 210023, Nanjing, China
| | - Wendong Jia
- State Key Laboratory of Analytical Chemistry for Life Sciences, School of Chemistry and Chemical Engineering, Nanjing University, 210023, Nanjing, China.,Chemistry and Biomedicine Innovation Center (ChemBIC), Nanjing University, 210023, Nanjing, China
| | - Panke Zhang
- State Key Laboratory of Analytical Chemistry for Life Sciences, School of Chemistry and Chemical Engineering, Nanjing University, 210023, Nanjing, China.,Chemistry and Biomedicine Innovation Center (ChemBIC), Nanjing University, 210023, Nanjing, China
| | - Hong-Yuan Chen
- State Key Laboratory of Analytical Chemistry for Life Sciences, School of Chemistry and Chemical Engineering, Nanjing University, 210023, Nanjing, China.,Chemistry and Biomedicine Innovation Center (ChemBIC), Nanjing University, 210023, Nanjing, China
| | - Shuo Huang
- State Key Laboratory of Analytical Chemistry for Life Sciences, School of Chemistry and Chemical Engineering, Nanjing University, 210023, Nanjing, China.,Chemistry and Biomedicine Innovation Center (ChemBIC), Nanjing University, 210023, Nanjing, China
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15
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Lento C, Wilson DJ. Subsecond Time-Resolved Mass Spectrometry in Dynamic Structural Biology. Chem Rev 2021; 122:7624-7646. [PMID: 34324314 DOI: 10.1021/acs.chemrev.1c00222] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/04/2023]
Abstract
Life at the molecular level is a dynamic world, where the key players-proteins, oligonucleotides, lipids, and carbohydrates-are in a perpetual state of structural flux, shifting rapidly between local minima on their conformational free energy landscapes. The techniques of classical structural biology, X-ray crystallography, structural NMR, and cryo-electron microscopy (cryo-EM), while capable of extraordinary structural resolution, are innately ill-suited to characterize biomolecules in their dynamically active states. Subsecond time-resolved mass spectrometry (MS) provides a unique window into the dynamic world of biological macromolecules, offering the capacity to directly monitor biochemical processes and conformational shifts with a structural dimension provided by the electrospray charge-state distribution, ion mobility, covalent labeling, or hydrogen-deuterium exchange. Over the past two decades, this suite of techniques has provided important insights into the inherently dynamic processes that drive function and pathogenesis in biological macromolecules, including (mis)folding, complexation, aggregation, ligand binding, and enzyme catalysis, among others. This Review provides a comprehensive account of subsecond time-resolved MS and the advances it has enabled in dynamic structural biology, with an emphasis on insights into the dynamic drivers of protein function.
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Affiliation(s)
- Cristina Lento
- Department of Chemistry, York University, Toronto, Ontario M3J 1P3, Canada
| | - Derek J Wilson
- Department of Chemistry, York University, Toronto, Ontario M3J 1P3, Canada
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16
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Hosoya Y, Ohkanda J. Intrinsically Disordered Proteins as Regulators of Transient Biological Processes and as Untapped Drug Targets. Molecules 2021; 26:2118. [PMID: 33917117 PMCID: PMC8067799 DOI: 10.3390/molecules26082118] [Citation(s) in RCA: 12] [Impact Index Per Article: 4.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/27/2021] [Revised: 03/29/2021] [Accepted: 03/30/2021] [Indexed: 12/31/2022] Open
Abstract
Intrinsically disordered proteins (IDPs) are critical players in the dynamic control of diverse cellular processes, and provide potential new drug targets because their dysregulation is closely related to many diseases. This review focuses on several medicinal studies that have identified low-molecular-weight inhibitors of IDPs. In addition, clinically relevant liquid-liquid phase separations-which critically involve both intermolecular interactions between IDPs and their posttranslational modification-are analyzed to understand the potential of IDPs as new drug targets.
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Affiliation(s)
| | - Junko Ohkanda
- Academic Assembly, Institute of Agriculture, Shinshu University, 8304 Minami-Minowa, Kami-Ina, Nagano 399-4598, Japan;
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17
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Lerner E, Barth A, Hendrix J, Ambrose B, Birkedal V, Blanchard SC, Börner R, Sung Chung H, Cordes T, Craggs TD, Deniz AA, Diao J, Fei J, Gonzalez RL, Gopich IV, Ha T, Hanke CA, Haran G, Hatzakis NS, Hohng S, Hong SC, Hugel T, Ingargiola A, Joo C, Kapanidis AN, Kim HD, Laurence T, Lee NK, Lee TH, Lemke EA, Margeat E, Michaelis J, Michalet X, Myong S, Nettels D, Peulen TO, Ploetz E, Razvag Y, Robb NC, Schuler B, Soleimaninejad H, Tang C, Vafabakhsh R, Lamb DC, Seidel CAM, Weiss S. FRET-based dynamic structural biology: Challenges, perspectives and an appeal for open-science practices. eLife 2021; 10:e60416. [PMID: 33779550 PMCID: PMC8007216 DOI: 10.7554/elife.60416] [Citation(s) in RCA: 119] [Impact Index Per Article: 39.7] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/29/2020] [Accepted: 02/09/2021] [Indexed: 12/18/2022] Open
Abstract
Single-molecule FRET (smFRET) has become a mainstream technique for studying biomolecular structural dynamics. The rapid and wide adoption of smFRET experiments by an ever-increasing number of groups has generated significant progress in sample preparation, measurement procedures, data analysis, algorithms and documentation. Several labs that employ smFRET approaches have joined forces to inform the smFRET community about streamlining how to perform experiments and analyze results for obtaining quantitative information on biomolecular structure and dynamics. The recent efforts include blind tests to assess the accuracy and the precision of smFRET experiments among different labs using various procedures. These multi-lab studies have led to the development of smFRET procedures and documentation, which are important when submitting entries into the archiving system for integrative structure models, PDB-Dev. This position paper describes the current 'state of the art' from different perspectives, points to unresolved methodological issues for quantitative structural studies, provides a set of 'soft recommendations' about which an emerging consensus exists, and lists openly available resources for newcomers and seasoned practitioners. To make further progress, we strongly encourage 'open science' practices.
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Affiliation(s)
- Eitan Lerner
- Department of Biological Chemistry, The Alexander Silberman Institute of Life Sciences, and The Center for Nanoscience and Nanotechnology, Faculty of Mathematics & Science, The Edmond J. Safra Campus, The Hebrew University of JerusalemJerusalemIsrael
| | - Anders Barth
- Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-UniversitätDüsseldorfGermany
| | - Jelle Hendrix
- Dynamic Bioimaging Lab, Advanced Optical Microscopy Centre and Biomedical Research Institute (BIOMED), Hasselt UniversityDiepenbeekBelgium
| | - Benjamin Ambrose
- Department of Chemistry, University of SheffieldSheffieldUnited Kingdom
| | - Victoria Birkedal
- Department of Chemistry and iNANO center, Aarhus UniversityAarhusDenmark
| | - Scott C Blanchard
- Department of Structural Biology, St. Jude Children's Research HospitalMemphisUnited States
| | - Richard Börner
- Laserinstitut HS Mittweida, University of Applied Science MittweidaMittweidaGermany
| | - Hoi Sung Chung
- Laboratory of Chemical Physics, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of HealthBethesdaUnited States
| | - Thorben Cordes
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians-Universität MünchenPlanegg-MartinsriedGermany
| | - Timothy D Craggs
- Department of Chemistry, University of SheffieldSheffieldUnited Kingdom
| | - Ashok A Deniz
- Department of Integrative Structural and Computational Biology, The Scripps Research InstituteLa JollaUnited States
| | - Jiajie Diao
- Department of Cancer Biology, University of Cincinnati School of MedicineCincinnatiUnited States
| | - Jingyi Fei
- Department of Biochemistry and Molecular Biology and The Institute for Biophysical Dynamics, University of ChicagoChicagoUnited States
| | - Ruben L Gonzalez
- Department of Chemistry, Columbia UniversityNew YorkUnited States
| | - Irina V Gopich
- Laboratory of Chemical Physics, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of HealthBethesdaUnited States
| | - Taekjip Ha
- Department of Biophysics and Biophysical Chemistry, Department of Biomedical Engineering, Johns Hopkins University School of Medicine, Howard Hughes Medical InstituteBaltimoreUnited States
| | - Christian A Hanke
- Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-UniversitätDüsseldorfGermany
| | - Gilad Haran
- Department of Chemical and Biological Physics, Weizmann Institute of ScienceRehovotIsrael
| | - Nikos S Hatzakis
- Department of Chemistry & Nanoscience Centre, University of CopenhagenCopenhagenDenmark
- Denmark Novo Nordisk Foundation Centre for Protein Research, Faculty of Health and Medical Sciences, University of CopenhagenCopenhagenDenmark
| | - Sungchul Hohng
- Department of Physics and Astronomy, and Institute of Applied Physics, Seoul National UniversitySeoulRepublic of Korea
| | - Seok-Cheol Hong
- Center for Molecular Spectroscopy and Dynamics, Institute for Basic Science and Department of Physics, Korea UniversitySeoulRepublic of Korea
| | - Thorsten Hugel
- Institute of Physical Chemistry and Signalling Research Centres BIOSS and CIBSS, University of FreiburgFreiburgGermany
| | - Antonino Ingargiola
- Department of Chemistry and Biochemistry, and Department of Physiology, University of California, Los AngelesLos AngelesUnited States
| | - Chirlmin Joo
- Department of BioNanoScience, Kavli Institute of Nanoscience, Delft University of TechnologyDelftNetherlands
| | - Achillefs N Kapanidis
- Biological Physics Research Group, Clarendon Laboratory, Department of Physics, University of OxfordOxfordUnited Kingdom
| | - Harold D Kim
- School of Physics, Georgia Institute of TechnologyAtlantaUnited States
| | - Ted Laurence
- Physical and Life Sciences Directorate, Lawrence Livermore National LaboratoryLivermoreUnited States
| | - Nam Ki Lee
- School of Chemistry, Seoul National UniversitySeoulRepublic of Korea
| | - Tae-Hee Lee
- Department of Chemistry, Pennsylvania State UniversityUniversity ParkUnited States
| | - Edward A Lemke
- Departments of Biology and Chemistry, Johannes Gutenberg UniversityMainzGermany
- Institute of Molecular Biology (IMB)MainzGermany
| | - Emmanuel Margeat
- Centre de Biologie Structurale (CBS), CNRS, INSERM, Universitié de MontpellierMontpellierFrance
| | | | - Xavier Michalet
- Department of Chemistry and Biochemistry, and Department of Physiology, University of California, Los AngelesLos AngelesUnited States
| | - Sua Myong
- Department of Biophysics, Johns Hopkins UniversityBaltimoreUnited States
| | - Daniel Nettels
- Department of Biochemistry and Department of Physics, University of ZurichZurichSwitzerland
| | - Thomas-Otavio Peulen
- Department of Bioengineering and Therapeutic Sciences, University of California, San FranciscoSan FranciscoUnited States
| | - Evelyn Ploetz
- Physical Chemistry, Department of Chemistry, Center for Nanoscience (CeNS), Center for Integrated Protein Science Munich (CIPSM) and Nanosystems Initiative Munich (NIM), Ludwig-Maximilians-UniversitätMünchenGermany
| | - Yair Razvag
- Department of Biological Chemistry, The Alexander Silberman Institute of Life Sciences, and The Center for Nanoscience and Nanotechnology, Faculty of Mathematics & Science, The Edmond J. Safra Campus, The Hebrew University of JerusalemJerusalemIsrael
| | - Nicole C Robb
- Warwick Medical School, University of WarwickCoventryUnited Kingdom
| | - Benjamin Schuler
- Department of Biochemistry and Department of Physics, University of ZurichZurichSwitzerland
| | - Hamid Soleimaninejad
- Biological Optical Microscopy Platform (BOMP), University of MelbourneParkvilleAustralia
| | - Chun Tang
- College of Chemistry and Molecular Engineering, PKU-Tsinghua Center for Life Sciences, Beijing National Laboratory for Molecular Sciences, Peking UniversityBeijingChina
| | - Reza Vafabakhsh
- Department of Molecular Biosciences, Northwestern UniversityEvanstonUnited States
| | - Don C Lamb
- Physical Chemistry, Department of Chemistry, Center for Nanoscience (CeNS), Center for Integrated Protein Science Munich (CIPSM) and Nanosystems Initiative Munich (NIM), Ludwig-Maximilians-UniversitätMünchenGermany
| | - Claus AM Seidel
- Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-UniversitätDüsseldorfGermany
| | - Shimon Weiss
- Department of Chemistry and Biochemistry, and Department of Physiology, University of California, Los AngelesLos AngelesUnited States
- Department of Physiology, CaliforniaNanoSystems Institute, University of California, Los AngelesLos AngelesUnited States
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18
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Dimura M, Peulen TO, Sanabria H, Rodnin D, Hemmen K, Hanke CA, Seidel CAM, Gohlke H. Automated and optimally FRET-assisted structural modeling. Nat Commun 2020; 11:5394. [PMID: 33106483 PMCID: PMC7589535 DOI: 10.1038/s41467-020-19023-1] [Citation(s) in RCA: 18] [Impact Index Per Article: 4.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/01/2020] [Accepted: 09/28/2020] [Indexed: 12/27/2022] Open
Abstract
FRET experiments can provide state-specific structural information of complex dynamic biomolecular assemblies. However, to overcome the sparsity of FRET experiments, they need to be combined with computer simulations. We introduce a program suite with (i) an automated design tool for FRET experiments, which determines how many and which FRET pairs should be used to minimize the uncertainty and maximize the accuracy of an integrative structure, (ii) an efficient approach for FRET-assisted coarse-grained structural modeling, and all-atom molecular dynamics simulations-based refinement, and (iii) a quantitative quality estimate for judging the accuracy of FRET-derived structures as opposed to precision. We benchmark our tools against simulated and experimental data of proteins with multiple conformational states and demonstrate an accuracy of ~3 Å RMSDCα against X-ray structures for sets of 15 to 23 FRET pairs. Free and open-source software for the introduced workflow is available at https://github.com/Fluorescence-Tools. A web server for FRET-assisted structural modeling of proteins is available at http://nmsim.de. To overcome the limitation of FRET data being too sparse to cover all structural details, FRET experiments need to be carefully designed and complemented with simulations. Here the authors present a toolkit for automated design of FRET experiments, which determines how many and which FRET pairs should be used to maximize the accuracy, and for FRET-assisted structural modeling and refinement at the atomistic level.
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Affiliation(s)
- Mykola Dimura
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany.,Institute for Pharmaceutical and Medicinal Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany
| | - Thomas-Otavio Peulen
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany
| | - Hugo Sanabria
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany.,Department of Physics and Astronomy, Clemson University, Clemson, SC, USA
| | - Dmitro Rodnin
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany
| | - Katherina Hemmen
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany
| | - Christian A Hanke
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany
| | - Claus A M Seidel
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany.
| | - Holger Gohlke
- Institute for Pharmaceutical and Medicinal Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany. .,John von Neumann Institute for Computing (NIC), Jülich Supercomputing Centre (JSC), and Institute of Biological Information Processing (IBI-7: Structural Biochemistry), Forschungszentrum Jülich GmbH, 52425, Jülich, Germany.
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19
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Pan M, Zhang Y, Yan G, Chen TY. Dissection of Interaction Kinetics through Single-Molecule Interaction Simulation. Anal Chem 2020; 92:11582-11589. [PMID: 32786469 DOI: 10.1021/acs.analchem.0c01014] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/12/2022]
Abstract
The ability to extract kinetic interaction parameters from single-molecule fluorescence resonance energy transfer trajectories without the need for solving complex single-molecule differential equations has the potential to address some of the critical biophysical questions. Here, we provide a noise-free single-molecule interaction simulation (SMIS) tool to give the expected dwell-time distributions and relative populations of each FRET level based on the assigned kinetic model and to dissect kinetic interaction parameters from single-molecule FRET trajectories. The method provides the expected dwell-time distributions, average transition rates, and relative populations of each FRET level based on the assigned kinetic model. By comparing with ground truth data and experimental data, we demonstrated that SMIS is useful to quantify the interaction kinetic rate constants without using the traditional single-molecule analytical solution approach.
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Affiliation(s)
- Manhua Pan
- Department of Chemistry, University of Houston, Houston, Texas 77204, United States
| | - Yuteng Zhang
- Department of Chemistry, University of Houston, Houston, Texas 77204, United States
| | - Guangjie Yan
- Department of Chemistry, University of Houston, Houston, Texas 77204, United States
| | - Tai-Yen Chen
- Department of Chemistry, University of Houston, Houston, Texas 77204, United States
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20
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Munc18-1 induces conformational changes of syntaxin-1 in multiple intermediates for SNARE assembly. Sci Rep 2020; 10:11623. [PMID: 32669573 PMCID: PMC7363831 DOI: 10.1038/s41598-020-68476-3] [Citation(s) in RCA: 10] [Impact Index Per Article: 2.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/16/2020] [Accepted: 06/25/2020] [Indexed: 01/16/2023] Open
Abstract
In neuronal exocytosis, SNARE assembly into a stable four-helix bundle drives membrane fusion. Previous studies have revealed that the SM protein Munc18-1 plays a critical role for precise SNARE assembly with the help of Munc13-1, but the underlying mechanism remains unclear. Here, we used single-molecule FRET assays with a nanodisc membrane reconstitution system to investigate the conformational dynamics of SNARE/Munc18-1 complexes in multiple intermediate steps towards the SNARE complex. We found that single Munc18-1 proteins induce the closed conformation of syntaxin-1 not only in the free syntaxin-1 but also in the t-SNARE (syntaxin-1/SNAP-25) complex. These results implicate that Munc18-1 may act as a gatekeeper for both binary and ternary SNARE complex formation by locking the syntaxin-1 in a cleft of Munc18-1. Furthermore, the kinetic analysis of the opening/closing transition reveals that the closed syntaxin-1 in the syntaxin-1/SNAP-25/Munc18-1 complex is less stable than that in the closed syntaxin-1/Munc18-1 complex, which is manifested by the infrequent closing transition, indicating that the conformational equilibrium of the ternary complex is biased toward the open conformation of syntaxin-1 compared with the binary complex.
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21
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Shinobu A, Kobayashi C, Matsunaga Y, Sugita Y. Building a macro-mixing dual-basin Gō model using the Multistate Bennett Acceptance Ratio. Biophys Physicobiol 2019; 16:310-321. [PMID: 31984186 PMCID: PMC6975896 DOI: 10.2142/biophysico.16.0_310] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.6] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/01/2019] [Accepted: 08/02/2019] [Indexed: 12/01/2022] Open
Abstract
The dual-basin Gō-model is a structural-based coarsegrained model for simulating a conformational transition between two known structures of a protein. Two parameters are required to produce a dual-basin potential mixed using two single-basin potentials, although the determination of mixing parameters is usually not straightforward. Here, we have developed an efficient scheme to determine the mixing parameters using the Multistate Bennett Acceptance Ratio (MBAR) method after short simulations with a set of parameters. In the scheme, MBAR allows us to predict observables at various unsimulated conditions, which are useful to improve the mixing parameters in the next round of iterative simulations. The number of iterations that are necessary for obtaining the converged mixing parameters are significantly reduced in the scheme. We applied the scheme to two proteins, the glutamine binding protein and the ribose binding protein, for showing the effectiveness in the parameter determination. After obtaining the converged parameters, both proteins show frequent conformational transitions between open and closed states, providing the theoretical basis to investigate structure-dynamics-function relationships of the proteins.
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Affiliation(s)
- Ai Shinobu
- Computational Biophysics Research Team, RIKEN Center for Computational Science, Kobe, Hyogo 650-0047, Japan
| | - Chigusa Kobayashi
- Computational Biophysics Research Team, RIKEN Center for Computational Science, Kobe, Hyogo 650-0047, Japan
| | - Yasuhiro Matsunaga
- Graduate School of Science and Engineering, Saitama University, Saitama 338-8570, Japan
| | - Yuji Sugita
- Computational Biophysics Research Team, RIKEN Center for Computational Science, Kobe, Hyogo 650-0047, Japan
- Theoretical Molecular Science Laboratory, RIKEN Cluster for Pioneering Research, Wako, Saitama 351-0198, Japan
- Laboratory for Biomolecular Function Simulation, RIKEN Center for Biosystems Dynamics Research, Kobe, Hyogo 650-0047, Japan
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22
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Xiao M, Lai W, Man T, Chang B, Li L, Chandrasekaran AR, Pei H. Rationally Engineered Nucleic Acid Architectures for Biosensing Applications. Chem Rev 2019; 119:11631-11717. [DOI: 10.1021/acs.chemrev.9b00121] [Citation(s) in RCA: 147] [Impact Index Per Article: 29.4] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/16/2022]
Affiliation(s)
- Mingshu Xiao
- Shanghai Key Laboratory of Green Chemistry and Chemical Processes, School of Chemistry and Molecular Engineering, East China Normal University, 500 Dongchuan Road, Shanghai 200241, P. R. China
| | - Wei Lai
- Shanghai Key Laboratory of Green Chemistry and Chemical Processes, School of Chemistry and Molecular Engineering, East China Normal University, 500 Dongchuan Road, Shanghai 200241, P. R. China
| | - Tiantian Man
- Shanghai Key Laboratory of Green Chemistry and Chemical Processes, School of Chemistry and Molecular Engineering, East China Normal University, 500 Dongchuan Road, Shanghai 200241, P. R. China
| | - Binbin Chang
- Shanghai Key Laboratory of Green Chemistry and Chemical Processes, School of Chemistry and Molecular Engineering, East China Normal University, 500 Dongchuan Road, Shanghai 200241, P. R. China
| | - Li Li
- Shanghai Key Laboratory of Green Chemistry and Chemical Processes, School of Chemistry and Molecular Engineering, East China Normal University, 500 Dongchuan Road, Shanghai 200241, P. R. China
| | - Arun Richard Chandrasekaran
- The RNA Institute, University at Albany, State University of New York, Albany, New York 12222, United States
| | - Hao Pei
- Shanghai Key Laboratory of Green Chemistry and Chemical Processes, School of Chemistry and Molecular Engineering, East China Normal University, 500 Dongchuan Road, Shanghai 200241, P. R. China
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23
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Hays JM, Cafiso DS, Kasson PM. Hybrid Refinement of Heterogeneous Conformational Ensembles Using Spectroscopic Data. J Phys Chem Lett 2019; 10:3410-3414. [PMID: 31181934 PMCID: PMC6605767 DOI: 10.1021/acs.jpclett.9b01407] [Citation(s) in RCA: 4] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 05/28/2023]
Abstract
Multistructured biomolecular systems play crucial roles in a wide variety of cellular processes but have resisted traditional methods of structure determination, which often resolve only a few low-energy states. High-resolution structure determination using experimental methods that yield distributional data remains extremely difficult, especially when the underlying conformational ensembles are quite heterogeneous. We have therefore developed a method to integrate sparse, multimultimodal spectroscopic data to obtain high-resolution estimates of conformational ensembles. We have tested our method by incorporating double electron-electron resonance data on the soluble N-ethylmaleimide-sensitive factor attachment receptor (SNARE) protein syntaxin-1a into biased molecular dynamics simulations. We find that our method substantially outperforms existing state-of-the-art methods in capturing syntaxin's open-closed conformational equilibrium and further yields new conformational states that are consistent with experimental data and may help in understanding syntaxin's function. Our improved methods for refining heterogeneous conformational ensembles from spectroscopic data will greatly accelerate the structural understanding of such systems.
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Affiliation(s)
- Jennifer M. Hays
- Department of Biomedical Engineering, University of Virginia, Charlottesville, VA, 22903
- Department of Molecular Physiology and Biophysics, University of Virginia, Charlottesville, VA, 22903
| | - David S. Cafiso
- Department of Molecular Physiology and Biophysics, University of Virginia, Charlottesville, VA, 22903
- Department of Chemistry, University of Virginia, Charlottesville, VA, 22903
| | - Peter M. Kasson
- Department of Biomedical Engineering, University of Virginia, Charlottesville, VA, 22903
- Department of Molecular Physiology and Biophysics, University of Virginia, Charlottesville, VA, 22903
- Science for Life Laboratory, Department of Cell and Molecular Biology, Uppsala University, 75124 Uppsala,
Sweden
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24
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Correlated protein conformational states and membrane dynamics during attack by pore-forming toxins. Proc Natl Acad Sci U S A 2019; 116:12839-12844. [PMID: 31189600 DOI: 10.1073/pnas.1821897116] [Citation(s) in RCA: 26] [Impact Index Per Article: 5.2] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/20/2022] Open
Abstract
Pore-forming toxins (PFTs) are a class of proteins implicated in a wide range of virulent bacterial infections and diseases. These toxins bind to target membranes and subsequently oligomerize to form functional pores that eventually lead to cell lysis. While the protein undergoes large conformational changes on the bilayer, the connection between intermediate oligomeric states and lipid reorganization during pore formation is largely unexplored. Cholesterol-dependent cytolysins (CDCs) are a subclass of PFTs widely implicated in food poisoning and other related infections. Using a prototypical CDC, listeriolysin O (LLO), we provide a microscopic connection between pore formation, lipid dynamics, and leakage kinetics by using a combination of Förster resonance energy transfer (FRET) and fluorescence correlation spectroscopy (FCS) measurements on single giant unilamellar vesicles (GUVs). Upon exposure to LLO, two distinct populations of GUVs with widely different leakage kinetics emerge. We attribute these differences to the existence of oligomeric intermediates, sampling various membrane-bound conformational states of the protein, and their intimate coupling to lipid rearrangement and dynamics. Molecular dynamics simulations capture the influence of various membrane-bound conformational states on the lipid and cholesterol dynamics, providing molecular interpretations to the FRET and FCS experiments. Our study establishes a microscopic connection between membrane binding and conformational changes and their influence on lipid reorganization during PFT-mediated cell lysis. Additionally, our study provides insights into membrane-mediated protein interactions widely implicated in cell signaling, fusion, folding, and other biomolecular processes.
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25
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Tunable microsecond dynamics of an allosteric switch regulate the activity of a AAA+ disaggregation machine. Nat Commun 2019; 10:1438. [PMID: 30926805 PMCID: PMC6440998 DOI: 10.1038/s41467-019-09474-6] [Citation(s) in RCA: 30] [Impact Index Per Article: 6.0] [Reference Citation Analysis] [Abstract] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/17/2018] [Accepted: 03/14/2019] [Indexed: 12/13/2022] Open
Abstract
Large protein machines are tightly regulated through allosteric communication channels. Here we demonstrate the involvement of ultrafast conformational dynamics in allosteric regulation of ClpB, a hexameric AAA+ machine that rescues aggregated proteins. Each subunit of ClpB contains a unique coiled-coil structure, the middle domain (M domain), proposed as a control element that binds the co-chaperone DnaK. Using single-molecule FRET spectroscopy, we probe the M domain during the chaperone cycle and find it to jump on the microsecond time scale between two states, whose structures are determined. The M-domain jumps are much faster than the overall activity of ClpB, making it an effectively continuous, tunable switch. Indeed, a series of allosteric interactions are found to modulate the dynamics, including binding of nucleotides, DnaK and protein substrates. This mode of dynamic control enables fast cellular adaptation and may be a general mechanism for the regulation of cellular machineries. Large protein machines are tightly regulated through allosteric communication channels. Here authors use single-molecule FRET and demonstrate the involvement of ultrafast conformational dynamics in the allosteric regulation of ClpB, a hexameric AAA+ machine that rescues aggregated proteins.
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26
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Jiang X, Zhang Z, Cheng K, Wu Q, Jiang L, Pielak GJ, Liu M, Li C. Membrane-mediated disorder-to-order transition of SNAP25 flexible linker facilitates its interaction with syntaxin-1 and SNARE-complex assembly. FASEB J 2019; 33:7985-7994. [PMID: 30916996 DOI: 10.1096/fj.201802796r] [Citation(s) in RCA: 6] [Impact Index Per Article: 1.2] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/15/2023]
Abstract
The soluble N-ethylmaleimide-sensitive factor attachment protein receptor (SNARE) complex comprises synaptosome-associated protein of 25 kDa (SNAP25), syntaxin-1a (syx-1), and synaptobrevin 2, which is essential for many physiologic processes requiring membrane fusion. Several studies imply that the loop region of SNAP25 plays important roles in SNARE-complex assembly. However, why and how the flexible loop facilitates the complex assembly remains poorly understood because it is purposely deleted in almost all structural studies. By using NMR spectroscopy and circular dichroism spectropolarimetry, we characterized SNAP25 structure and interactions with other SNAREs in aqueous buffer and in the membrane. We found that the N-terminal of the SNAP25 loop region binds with membrane, and this interaction induced a disorder-to-order conformational change of the loop, resulting in enhanced interaction between the C-terminal of the SNAP25 loop and syx-1. We further proved that SNARE-complex assembly efficiency decreased when we disrupted the electrostatic interaction between C-terminal of the SNAP25 loop and syx-1, suggesting that the SNAP25 loop region facilitates SNARE-complex assembly through promoting prefusion SNARE binary complex formation. Our work elucidates the role of the flexible loop and the membrane environment in SNARE-complex assembly at the residue level, which helps to understand membrane fusion, a fundamental transport and communication process in cells.-Jiang, X., Zhang, Z., Cheng, K., Wu, Q., Jiang, L., Pielak, G. J., Liu, M., Li, C. Membrane-mediated disorder-to-order transition of SNAP25 flexible linker facilitates its interaction with syntaxin-1 and SNARE-complex assembly.
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Affiliation(s)
- Xin Jiang
- Key Laboratory of Magnetic Resonance in Biological Systems, State Key Laboratory of Magnetic Resonance and Atomic and Molecular Physics, National Center for Magnetic Resonance in Wuhan, Wuhan National Laboratory for Optoelectronics, Wuhan Institute of Physics and Mathematics, Chinese Academy of Sciences, Wuhan, China.,Graduate University of Chinese Academy of Sciences, Beijing, China
| | - Zeting Zhang
- Key Laboratory of Magnetic Resonance in Biological Systems, State Key Laboratory of Magnetic Resonance and Atomic and Molecular Physics, National Center for Magnetic Resonance in Wuhan, Wuhan National Laboratory for Optoelectronics, Wuhan Institute of Physics and Mathematics, Chinese Academy of Sciences, Wuhan, China
| | - Kai Cheng
- Key Laboratory of Magnetic Resonance in Biological Systems, State Key Laboratory of Magnetic Resonance and Atomic and Molecular Physics, National Center for Magnetic Resonance in Wuhan, Wuhan National Laboratory for Optoelectronics, Wuhan Institute of Physics and Mathematics, Chinese Academy of Sciences, Wuhan, China
| | - Qiong Wu
- Key Laboratory of Magnetic Resonance in Biological Systems, State Key Laboratory of Magnetic Resonance and Atomic and Molecular Physics, National Center for Magnetic Resonance in Wuhan, Wuhan National Laboratory for Optoelectronics, Wuhan Institute of Physics and Mathematics, Chinese Academy of Sciences, Wuhan, China
| | - Ling Jiang
- Key Laboratory of Magnetic Resonance in Biological Systems, State Key Laboratory of Magnetic Resonance and Atomic and Molecular Physics, National Center for Magnetic Resonance in Wuhan, Wuhan National Laboratory for Optoelectronics, Wuhan Institute of Physics and Mathematics, Chinese Academy of Sciences, Wuhan, China.,Graduate University of Chinese Academy of Sciences, Beijing, China
| | - Gary J Pielak
- Department of Chemistry and Department of Biochemistry and Biophysics, University of North Carolina-Chapel Hill, Chapel Hill, North Carolina, USA.,Lineberger Comprehensive Cancer Center, University of North Carolina-Chapel Hill, Chapel Hill, North Carolina, USA.,Integrative Program for Biological and Genome Sciences, University of North Carolina-Chapel Hill, Chapel Hill, North Carolina, USA
| | - Maili Liu
- Key Laboratory of Magnetic Resonance in Biological Systems, State Key Laboratory of Magnetic Resonance and Atomic and Molecular Physics, National Center for Magnetic Resonance in Wuhan, Wuhan National Laboratory for Optoelectronics, Wuhan Institute of Physics and Mathematics, Chinese Academy of Sciences, Wuhan, China.,Graduate University of Chinese Academy of Sciences, Beijing, China
| | - Conggang Li
- Key Laboratory of Magnetic Resonance in Biological Systems, State Key Laboratory of Magnetic Resonance and Atomic and Molecular Physics, National Center for Magnetic Resonance in Wuhan, Wuhan National Laboratory for Optoelectronics, Wuhan Institute of Physics and Mathematics, Chinese Academy of Sciences, Wuhan, China.,Graduate University of Chinese Academy of Sciences, Beijing, China
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27
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Dingjan I, Linders PTA, Verboogen DRJ, Revelo NH, Ter Beest M, van den Bogaart G. Endosomal and Phagosomal SNAREs. Physiol Rev 2018; 98:1465-1492. [PMID: 29790818 DOI: 10.1152/physrev.00037.2017] [Citation(s) in RCA: 51] [Impact Index Per Article: 8.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/26/2022] Open
Abstract
The soluble N-ethylmaleimide-sensitive factor attachment protein receptor (SNARE) protein family is of vital importance for organelle communication. The complexing of cognate SNARE members present in both the donor and target organellar membranes drives the membrane fusion required for intracellular transport. In the endocytic route, SNARE proteins mediate trafficking between endosomes and phagosomes with other endosomes, lysosomes, the Golgi apparatus, the plasma membrane, and the endoplasmic reticulum. The goal of this review is to provide an overview of the SNAREs involved in endosomal and phagosomal trafficking. Of the 38 SNAREs present in humans, 30 have been identified at endosomes and/or phagosomes. Many of these SNAREs are targeted by viruses and intracellular pathogens, which thereby reroute intracellular transport for gaining access to nutrients, preventing their degradation, and avoiding their detection by the immune system. A fascinating picture is emerging of a complex transport network with multiple SNAREs being involved in consecutive trafficking routes.
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Affiliation(s)
- Ilse Dingjan
- Department of Tumor Immunology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center , Nijmegen , The Netherlands ; and Department of Molecular Immunology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen , Groningen , The Netherlands
| | - Peter T A Linders
- Department of Tumor Immunology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center , Nijmegen , The Netherlands ; and Department of Molecular Immunology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen , Groningen , The Netherlands
| | - Danielle R J Verboogen
- Department of Tumor Immunology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center , Nijmegen , The Netherlands ; and Department of Molecular Immunology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen , Groningen , The Netherlands
| | - Natalia H Revelo
- Department of Tumor Immunology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center , Nijmegen , The Netherlands ; and Department of Molecular Immunology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen , Groningen , The Netherlands
| | - Martin Ter Beest
- Department of Tumor Immunology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center , Nijmegen , The Netherlands ; and Department of Molecular Immunology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen , Groningen , The Netherlands
| | - Geert van den Bogaart
- Department of Tumor Immunology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center , Nijmegen , The Netherlands ; and Department of Molecular Immunology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen , Groningen , The Netherlands
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28
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Abstract
Web spiders synthesize silk fibers of unique strength and extensibility through the controlled self-assembly of protein building blocks, so-called spidroins. The spidroin C-terminal domain is highly conserved and connects two polypeptide chains through formation of an all-helical, intertwined dimer. Here we use contact-induced fluorescence self-quenching and resonance energy transfer in combination with far-UV circular dichroism spectroscopy as three orthogonal structural probes to dissect the mechanism of folding and dimerization of a spidroin C-terminal domain from the major ampullate gland of the nursery web spider Euprosthenops australis. We show that helices forming the dimer core assemble very rapidly and fold on association. Subsequently, peripheral helices fold and dock slowly onto the preformed core. Lability of outer helices facilitates formation of a highly expanded, partially folded dimer. The high end-to-end distance of chain termini in the partially folded dimer suggests an extensibility module that contributes to elasticity of spider silk.
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29
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Quantifying Intramolecular Protein Conformational Dynamics Under Lipid Interaction Using smFRET and FCCS. Methods Mol Biol 2018. [PMID: 30317517 DOI: 10.1007/978-1-4939-8760-3_23] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register]
Abstract
Fӧrster-type resonance energy transfer (FRET) with fluorescence cross-correlation spectroscopy (FCCS) is a powerful combination for observing intramolecular conformational dynamics on the micro- to millisecond timescale. Owing to its sensitivity to various physical parameters, FRET-FCCS has also been used to detect the reagent effects on proteins dynamics. However, FRET-FCCS alone cannot acquire the exact measurements of rate constants. Moreover, this technique is highly model dependent and can be unreliable when determining too many parameters at once. On the contrary, single-molecular FRET (smFRET) can measure the conformational states and their populations directly, although it is extremely challenging for probing fast dynamics under 1 ms. In this chapter, we describe how to realize sub-millisecond conformational dynamics measurements of a SNARE protein Ykt6 under lipid environments by smFRET and FRET-FCCS. This protocol includes sample preparation, microscope designs, data acquisition, and analysis methodology.
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30
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Yin P, Gandasi NR, Arora S, Omar-Hmeadi M, Saras J, Barg S. Syntaxin clusters at secretory granules in a munc18-bound conformation. Mol Biol Cell 2018; 29:2700-2708. [PMID: 30156474 PMCID: PMC6249827 DOI: 10.1091/mbc.e17-09-0541] [Citation(s) in RCA: 10] [Impact Index Per Article: 1.7] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/26/2022] Open
Abstract
Syntaxin (stx)-1 is an integral plasma membrane protein that is crucial for two distinct steps of regulated exocytosis, docking of secretory granules at the plasma membrane and membrane fusion. During docking, stx1 clusters at the granule docking site, together with the S/M protein munc18. Here we determined features of stx1 that contribute to its clustering at granules. In live insulin-secreting cells, stx1 and stx3 (but not stx4 or stx11) accumulated at docked granules, and stx1 (but not stx4) rescued docking in cells expressing botulinum neurotoxin-C. Using a series of stx1 deletion mutants and stx1/4 chimeras, we found that all four helical domains (Ha, Hb, Hc, SNARE) and the short N-terminal peptide contribute to recruitment to granules. However, only the Hc domain confers specificity, and it must be derived from stx1 for recruitment to occur. Point mutations in the Hc or the N-terminal peptide designed to interfere with binding to munc18-1 prevent stx1 from clustering at granules, and a mutant munc18 deficient in binding to stx1 does not cluster at granules. We conclude that stx1 is recruited to the docking site in a munc18-1–bound conformation, providing a rationale for the requirement for both proteins for granule docking.
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Affiliation(s)
- Peng Yin
- Institute of Medical Cell Biology, Uppsala University, 75123 Uppsala, Sweden
| | - Nikhil R Gandasi
- Institute of Medical Cell Biology, Uppsala University, 75123 Uppsala, Sweden
| | - Swati Arora
- Institute of Medical Cell Biology, Uppsala University, 75123 Uppsala, Sweden
| | - Muhmmad Omar-Hmeadi
- Institute of Medical Cell Biology, Uppsala University, 75123 Uppsala, Sweden
| | - Jan Saras
- Institute of Medical Cell Biology, Uppsala University, 75123 Uppsala, Sweden
| | - Sebastian Barg
- Institute of Medical Cell Biology, Uppsala University, 75123 Uppsala, Sweden
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31
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Devaux J, Dhifallah S, De Maria M, Stuart-Lopez G, Becq H, Milh M, Molinari F, Aniksztejn L. A possible link betweenKCNQ2- andSTXBP1-related encephalopathies: STXBP1 reduces the inhibitory impact of syntaxin-1A on M current. Epilepsia 2017; 58:2073-2084. [DOI: 10.1111/epi.13927] [Citation(s) in RCA: 6] [Impact Index Per Article: 0.9] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Accepted: 09/26/2017] [Indexed: 01/05/2023]
Affiliation(s)
- Jérôme Devaux
- CNRS, CRN2M-UMR7286; Aix-Marseille University; Marseille France
| | - Sandra Dhifallah
- INSERM UMR_S901; Mediterranean Neurobiology Institute (INMED); Aix-Marseille University; Marseille France
- Institute of Molecular and Cellular Pharmacology (IPMC); CNRS; Nice Sophia-Antipolis University; Valbonne France
| | - Michela De Maria
- INSERM UMR_S901; Mediterranean Neurobiology Institute (INMED); Aix-Marseille University; Marseille France
- Department of Medicine and Health Sciences; University of Molise; Campobasso Italy
| | - Geoffrey Stuart-Lopez
- INSERM UMR_S901; Mediterranean Neurobiology Institute (INMED); Aix-Marseille University; Marseille France
- UMR5203 Institute of Functional Genomic (IGF); CNRS; Montpellier France
| | - Hélène Becq
- INSERM UMR_S901; Mediterranean Neurobiology Institute (INMED); Aix-Marseille University; Marseille France
| | - Mathieu Milh
- Timone Children Hospital, Pediatric Neurology department; APHM; Marseille France
- GMGF, INSERM UMR_S910; Aix-Marseille University; Marseille France
| | - Florence Molinari
- INSERM UMR_S901; Mediterranean Neurobiology Institute (INMED); Aix-Marseille University; Marseille France
| | - Laurent Aniksztejn
- INSERM UMR_S901; Mediterranean Neurobiology Institute (INMED); Aix-Marseille University; Marseille France
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32
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Morey C, Kienle CN, Klöpper TH, Burkhardt P, Fasshauer D. Evidence for a conserved inhibitory binding mode between the membrane fusion assembly factors Munc18 and syntaxin in animals. J Biol Chem 2017; 292:20449-20460. [PMID: 29046354 DOI: 10.1074/jbc.m117.811182] [Citation(s) in RCA: 8] [Impact Index Per Article: 1.1] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/08/2017] [Revised: 10/10/2017] [Indexed: 12/30/2022] Open
Abstract
The membrane fusion necessary for vesicle trafficking is driven by the assembly of heterologous SNARE proteins orchestrated by the binding of Sec1/Munc18 (SM) proteins to specific syntaxin SNARE proteins. However, the precise mode of interaction between SM proteins and SNAREs is debated, as contrasting binding modes have been found for different members of the SM protein family, including the three vertebrate Munc18 isoforms. While different binding modes could be necessary, given their roles in different secretory processes in different tissues, the structural similarity of the three isoforms makes this divergence perplexing. Although the neuronal isoform Munc18a is well-established to bind tightly to both the closed conformation and the N-peptide of syntaxin 1a, thereby inhibiting SNARE complex formation, Munc18b and -c, which have a more widespread distribution, are reported to mainly interact with the N-peptide of their partnering syntaxins and are thought to instead promote SNARE complex formation. We have reinvestigated the interaction between Munc18c and syntaxin 4 (Syx4). Using isothermal titration calorimetry, we found that Munc18c, like Munc18a, binds to both the closed conformation and the N-peptide of Syx4. Furthermore, using a novel kinetic approach, we found that Munc18c, like Munc18a, slows down SNARE complex formation through high-affinity binding to syntaxin. This strongly suggests that secretory Munc18s in general control the accessibility of the bound syntaxin, probably preparing it for SNARE complex assembly.
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Affiliation(s)
- Czuee Morey
- From the Département des neurosciences fondamentales, Université de Lausanne, Rue du Bugnon 9, CH-1005 Lausanne, Switzerland
| | - C Nickias Kienle
- From the Département des neurosciences fondamentales, Université de Lausanne, Rue du Bugnon 9, CH-1005 Lausanne, Switzerland
| | - Tobias H Klöpper
- Whitehat Life Sciences Ltd., 20 Wenlock Road, N1 7GU London, United Kingdom, and
| | - Pawel Burkhardt
- the Marine Biological Association, Citadel Hill Marine Laboratory, Plymouth PL1 2PB, United Kingdom
| | - Dirk Fasshauer
- From the Département des neurosciences fondamentales, Université de Lausanne, Rue du Bugnon 9, CH-1005 Lausanne, Switzerland,
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33
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Peulen TO, Opanasyuk O, Seidel CAM. Combining Graphical and Analytical Methods with Molecular Simulations To Analyze Time-Resolved FRET Measurements of Labeled Macromolecules Accurately. J Phys Chem B 2017; 121:8211-8241. [PMID: 28709377 PMCID: PMC5592652 DOI: 10.1021/acs.jpcb.7b03441] [Citation(s) in RCA: 49] [Impact Index Per Article: 7.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/22/2022]
Abstract
![]()
Förster resonance energy transfer
(FRET) measurements from
a donor, D, to an acceptor, A, fluorophore are frequently used in vitro and in live cells to reveal information on the
structure and dynamics of DA labeled macromolecules. Accurate descriptions
of FRET measurements by molecular models are complicated because the
fluorophores are usually coupled to the macromolecule via flexible
long linkers allowing for diffusional exchange between multiple states
with different fluorescence properties caused by distinct environmental
quenching, dye mobilities, and variable DA distances. It is often
assumed for the analysis of fluorescence intensity decays that DA
distances and D quenching are uncorrelated (homogeneous quenching
by FRET) and that the exchange between distinct fluorophore states
is slow (quasistatic). This allows us to introduce the FRET-induced
donor decay, εD(t), a function solely
depending on the species fraction distribution of the rate constants
of energy transfer by FRET, for a convenient joint analysis of fluorescence
decays of FRET and reference samples by integrated graphical and analytical
procedures. Additionally, we developed a simulation toolkit to model
dye diffusion, fluorescence quenching by the protein surface, and
FRET. A benchmark study with simulated fluorescence decays of 500
protein structures demonstrates that the quasistatic homogeneous model
works very well and recovers for single conformations the average
DA distances with an accuracy of < 2%. For more complex
cases, where proteins adopt multiple conformations with significantly
different dye environments (heterogeneous case), we introduce a general
analysis framework and evaluate its power in resolving heterogeneities
in DA distances. The developed fast simulation methods, relying on
Brownian dynamics of a coarse-grained dye in its sterically accessible
volume, allow us to incorporate structural information in the decay
analysis for heterogeneous cases by relating dye states with protein
conformations to pave the way for fluorescence and FRET-based dynamic
structural biology. Finally, we present theories and simulations to
assess the accuracy and precision of steady-state and time-resolved
FRET measurements in resolving DA distances on the single-molecule
and ensemble level and provide a rigorous framework for estimating
approximation, systematic, and statistical errors.
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Affiliation(s)
- Thomas-Otavio Peulen
- Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität , Universitätsstraße 1, 40225 Düsseldorf, Germany
| | - Oleg Opanasyuk
- Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität , Universitätsstraße 1, 40225 Düsseldorf, Germany
| | - Claus A M Seidel
- Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität , Universitätsstraße 1, 40225 Düsseldorf, Germany
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34
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Abstract
In this study, we overview resonance energy transfer between molecules in the presence of plasmonic structures and derive an explicit Förster-type expression for the rate of plasmon-coupled resonance energy transfer (PC-RET). The proposed theory is general for energy transfer in the presence of materials with any space-dependent, frequency-dependent, or complex dielectric functions. Furthermore, the theory allows us to develop the concept of a generalized spectral overlap (GSO) J̃ (the integral of the molecular absorption coefficient, normalized emission spectrum, and the plasmon coupling factor) for understanding the wavelength dependence of PC-RET and to estimate the rate of PC-RET WET. Indeed, WET = (8.785 × 10-25 mol) ϕDτD-1J̃, where ϕD is donor fluorescence quantum yield and τD is the emission lifetime. Simulations of the GSO for PC-RET show that the most important spectral region for PC-RET is not necessarily near the maximum overlap of donor emission and acceptor absorption. Instead a significant plasmonic contribution can involve a different spectral region from the extinction maximum of the plasmonic structure. This study opens a promising direction for exploring exciton transport in plasmonic nanostructures, with possible applications in spectroscopy, photonics, biosensing, and energy devices.
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Affiliation(s)
- Liang-Yan Hsu
- Department of Chemistry, Northwestern University , 2145 Sheridan Road, Evanston, Illinois 60208-3113, United States
| | - Wendu Ding
- Department of Chemistry, Northwestern University , 2145 Sheridan Road, Evanston, Illinois 60208-3113, United States
| | - George C Schatz
- Department of Chemistry, Northwestern University , 2145 Sheridan Road, Evanston, Illinois 60208-3113, United States
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35
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Ma J, Yanez-Orozco IS, Rezaei Adariani S, Dolino D, Jayaraman V, Sanabria H. High Precision FRET at Single-molecule Level for Biomolecule Structure Determination. J Vis Exp 2017. [PMID: 28570518 DOI: 10.3791/55623] [Citation(s) in RCA: 12] [Impact Index Per Article: 1.7] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/16/2023] Open
Abstract
A protocol on how to perform high-precision interdye distance measurements using Förster resonance energy transfer (FRET) at the single-molecule level in multiparameter fluorescence detection (MFD) mode is presented here. MFD maximizes the usage of all "dimensions" of fluorescence to reduce photophysical and experimental artifacts and allows for the measurement of interdye distance with an accuracy up to ~1 Å in rigid biomolecules. This method was used to identify three conformational states of the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor to explain the activation of the receptor upon ligand binding. When comparing the known crystallographic structures with experimental measurements, they agreed within less than 3 Å for more dynamic biomolecules. Gathering a set of distance restraints that covers the entire dimensionality of the biomolecules would make it possible to provide a structural model of dynamic biomolecules.
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Affiliation(s)
- Junyan Ma
- Department of Chemistry, Clemson University
| | | | | | - Drew Dolino
- Department of Biochemistry and Molecular Biology, Center for Membrane Biology, Graduate School for Biomedical Sciences, University of Texas Health Science Center
| | - Vasanthi Jayaraman
- Department of Biochemistry and Molecular Biology, Center for Membrane Biology, Graduate School for Biomedical Sciences, University of Texas Health Science Center
| | - Hugo Sanabria
- Department of Physics and Astronomy, Clemson University;
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36
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Okamoto K, Sako Y. Recent advances in FRET for the study of protein interactions and dynamics. Curr Opin Struct Biol 2017; 46:16-23. [PMID: 29800904 DOI: 10.1016/j.sbi.2017.03.010] [Citation(s) in RCA: 68] [Impact Index Per Article: 9.7] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/07/2016] [Revised: 02/20/2017] [Accepted: 03/20/2017] [Indexed: 11/20/2022]
Abstract
Förster/fluorescence resonance energy transfer (FRET) has been extensively used to detect the binding state or conformation of biomolecules. In the past few decades, various in vitro and in vivo applications of FRET measurement have been developed, including FRET probes, in-cell measurements, single-molecule measurements, and combination with computer simulation. In this review, we describe recent advances in FRET methods for examining biomolecular interactions and dynamics: (i) phasor plot analysis for quantitative analysis of protein interactions, (ii) single-molecule FRET measurement for detecting conformational dynamics in live cells, and (iii) data assimilation using molecular dynamics simulation to evaluate conformation of the whole protein.
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Affiliation(s)
- Kenji Okamoto
- Cellular Informatics Laboratory, RIKEN, 2-1 Hirosawa, Wako, Saitama 351-0198, Japan.
| | - Yasushi Sako
- Cellular Informatics Laboratory, RIKEN, 2-1 Hirosawa, Wako, Saitama 351-0198, Japan
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37
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Dörfler T, Eilert T, Röcker C, Nagy J, Michaelis J. Structural Information from Single-molecule FRET Experiments Using the Fast Nano-positioning System. J Vis Exp 2017:54782. [PMID: 28287526 PMCID: PMC5407667 DOI: 10.3791/54782] [Citation(s) in RCA: 10] [Impact Index Per Article: 1.4] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/24/2022] Open
Abstract
Single-molecule Förster Resonance Energy Transfer (smFRET) can be used to obtain structural information on biomolecular complexes in real-time. Thereby, multiple smFRET measurements are used to localize an unknown dye position inside a protein complex by means of trilateration. In order to obtain quantitative information, the Nano-Positioning System (NPS) uses probabilistic data analysis to combine structural information from X-ray crystallography with single-molecule fluorescence data to calculate not only the most probable position but the complete three-dimensional probability distribution, termed posterior, which indicates the experimental uncertainty. The concept was generalized for the analysis of smFRET networks containing numerous dye molecules. The latest version of NPS, Fast-NPS, features a new algorithm using Bayesian parameter estimation based on Markov Chain Monte Carlo sampling and parallel tempering that allows for the analysis of large smFRET networks in a comparably short time. Moreover, Fast-NPS allows the calculation of the posterior by choosing one of five different models for each dye, that account for the different spatial and orientational behavior exhibited by the dye molecules due to their local environment. Here we present a detailed protocol for obtaining smFRET data and applying the Fast-NPS. We provide detailed instructions for the acquisition of the three input parameters of Fast-NPS: the smFRET values, as well as the quantum yield and anisotropy of the dye molecules. Recently, the NPS has been used to elucidate the architecture of an archaeal open promotor complex. This data is used to demonstrate the influence of the five different dye models on the posterior distribution.
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38
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The use of fluorescence correlation spectroscopy to characterize the molecular mobility of fluorescently labelled G protein-coupled receptors. Biochem Soc Trans 2016; 44:624-9. [PMID: 27068980 PMCID: PMC5264494 DOI: 10.1042/bst20150285] [Citation(s) in RCA: 14] [Impact Index Per Article: 1.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/15/2016] [Indexed: 01/10/2023]
Abstract
The membranes of living cells have been shown to be highly organized into distinct microdomains, which has spatial and temporal consequences for the interaction of membrane bound receptors and their signalling partners as complexes. Fluorescence correlation spectroscopy (FCS) is a technique with single cell sensitivity that sheds light on the molecular dynamics of fluorescently labelled receptors, ligands or signalling complexes within small plasma membrane regions of living cells. This review provides an overview of the use of FCS to probe the real time quantification of the diffusion and concentration of G protein-coupled receptors (GPCRs), primarily to gain insights into ligand–receptor interactions and the molecular composition of signalling complexes. In addition we document the use of photon counting histogram (PCH) analysis to investigate how changes in molecular brightness (ε) can be a sensitive indicator of changes in molecular mass of fluorescently labelled moieties.
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39
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Dimura M, Peulen TO, Hanke CA, Prakash A, Gohlke H, Seidel CA. Quantitative FRET studies and integrative modeling unravel the structure and dynamics of biomolecular systems. Curr Opin Struct Biol 2016; 40:163-185. [PMID: 27939973 DOI: 10.1016/j.sbi.2016.11.012] [Citation(s) in RCA: 113] [Impact Index Per Article: 14.1] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/04/2016] [Revised: 11/11/2016] [Accepted: 11/11/2016] [Indexed: 01/11/2023]
Abstract
Förster Resonance Energy Transfer (FRET) combined with single-molecule spectroscopy probes macromolecular structure and dynamics and identifies coexisting conformational states. We review recent methodological developments in integrative structural modeling by satisfying spatial restraints on networks of FRET pairs (hybrid-FRET). We discuss procedures to incorporate prior structural knowledge and to obtain optimal distance networks. Finally, a workflow for hybrid-FRET is presented that automates integrative structural modeling and experiment planning to put hybrid-FRET on rails. To test this workflow, we simulate realistic single-molecule experiments and resolve three protein conformers, exchanging at 30μs and 10ms, with accuracies of 1-3Å RMSD versus the target structure. Incorporation of data from other spectroscopies and imaging is also discussed.
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Affiliation(s)
- Mykola Dimura
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225 Düsseldorf, Germany; Institute for Pharmaceutical and Medicinal Chemistry, Heinrich Heine University Düsseldorf, 40225 Düsseldorf, Germany
| | - Thomas O Peulen
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225 Düsseldorf, Germany
| | - Christian A Hanke
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225 Düsseldorf, Germany
| | - Aiswaria Prakash
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225 Düsseldorf, Germany
| | - Holger Gohlke
- Institute for Pharmaceutical and Medicinal Chemistry, Heinrich Heine University Düsseldorf, 40225 Düsseldorf, Germany
| | - Claus Am Seidel
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225 Düsseldorf, Germany.
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40
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Delaforge E, Milles S, Huang JR, Bouvier D, Jensen MR, Sattler M, Hart DJ, Blackledge M. Investigating the Role of Large-Scale Domain Dynamics in Protein-Protein Interactions. Front Mol Biosci 2016; 3:54. [PMID: 27679800 PMCID: PMC5020063 DOI: 10.3389/fmolb.2016.00054] [Citation(s) in RCA: 18] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/04/2016] [Accepted: 08/30/2016] [Indexed: 12/21/2022] Open
Abstract
Intrinsically disordered linkers provide multi-domain proteins with degrees of conformational freedom that are often essential for function. These highly dynamic assemblies represent a significant fraction of all proteomes, and deciphering the physical basis of their interactions represents a considerable challenge. Here we describe the difficulties associated with mapping the large-scale domain dynamics and describe two recent examples where solution state methods, in particular NMR spectroscopy, are used to investigate conformational exchange on very different timescales.
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Affiliation(s)
- Elise Delaforge
- Institut de Biologie Structurale, CEA, Centre National de la Recherche Scientifique, University Grenoble Alpes Grenoble, France
| | - Sigrid Milles
- Institut de Biologie Structurale, CEA, Centre National de la Recherche Scientifique, University Grenoble Alpes Grenoble, France
| | - Jie-Rong Huang
- Institut de Biologie Structurale, CEA, Centre National de la Recherche Scientifique, University Grenoble Alpes Grenoble, France
| | - Denis Bouvier
- Institut de Biologie Structurale, CEA, Centre National de la Recherche Scientifique, University Grenoble Alpes Grenoble, France
| | - Malene Ringkjøbing Jensen
- Institut de Biologie Structurale, CEA, Centre National de la Recherche Scientifique, University Grenoble Alpes Grenoble, France
| | - Michael Sattler
- Institute of Structural Biology, Helmholtz Zentrum MünchenNeuherberg, Germany; Center for Integrated Protein Science Munich at Biomolecular NMR, Technische Universität MünchenGarching, Germany
| | - Darren J Hart
- Institut de Biologie Structurale, CEA, Centre National de la Recherche Scientifique, University Grenoble Alpes Grenoble, France
| | - Martin Blackledge
- Institut de Biologie Structurale, CEA, Centre National de la Recherche Scientifique, University Grenoble Alpes Grenoble, France
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41
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Chen J, Pyle JR, Sy Piecco KW, Kolomeisky AB, Landes CF. A Two-Step Method for smFRET Data Analysis. J Phys Chem B 2016; 120:7128-32. [DOI: 10.1021/acs.jpcb.6b05697] [Citation(s) in RCA: 15] [Impact Index Per Article: 1.9] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/31/2023]
Affiliation(s)
- Jixin Chen
- Department
of Chemistry and Biochemistry, Ohio University, Athens, Ohio 45701, United States
| | - Joseph R. Pyle
- Department
of Chemistry and Biochemistry, Ohio University, Athens, Ohio 45701, United States
| | - Kurt Waldo Sy Piecco
- Department
of Chemistry and Biochemistry, Ohio University, Athens, Ohio 45701, United States
| | | | - Christy F. Landes
- Department
of Chemistry, Rice University, Houston, Texas 77251, United States
- Department
of Electrical and Computer Engineering, Rice University, Houston, Texas 77251, United States
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42
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DeVore MS, Braimah A, Benson DR, Johnson CK. Single-Molecule FRET States, Conformational Interchange, and Conformational Selection by Dye Labels in Calmodulin. J Phys Chem B 2016; 120:4357-64. [PMID: 27111039 DOI: 10.1021/acs.jpcb.6b00864] [Citation(s) in RCA: 6] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/28/2022]
Abstract
We investigate the roles of measurement time scale and the nature of the fluorophores in the FRET states measured for calmodulin, a calcium signaling protein known to undergo pronounced conformational changes. The measured FRET distributions depend markedly on the measurement time scale (nanosecond or microsecond). Comparison of FRET distributions measured by donor fluorescence decay with FRET distributions recovered from single-molecule burst measurements binned over time scales of 90 μs to 1 ms reveals conformational averaging over the intervening time regimes. We find further that, particularly in the presence of saturating Ca(2+), the nature of the measured single-molecule FRET distribution depends markedly on the identity of the FRET pair. The results suggest interchange between conformational states on time scales of hundreds of microseconds or less. Interaction with a fluorophore such as the dye Texas Red alters both the nature of the measured FRET distributions and the dynamics of conformational interchange. The results further suggest that the fluorophore may not be merely a benign reporter of protein conformations in FRET studies, but may in fact alter the conformational landscape.
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Affiliation(s)
- Matthew S DeVore
- Department of Chemistry, University of Kansas , Lawrence, Kansas 66045, United States
| | - Adebayo Braimah
- Department of Chemistry, University of Kansas , Lawrence, Kansas 66045, United States
| | - David R Benson
- Department of Chemistry, University of Kansas , Lawrence, Kansas 66045, United States
| | - Carey K Johnson
- Department of Chemistry, University of Kansas , Lawrence, Kansas 66045, United States
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43
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Ungvári T, Gogolák P, Bagdány M, Damjanovich L, Bene L. Perrin and Förster unified: Dual-laser triple-polarization FRET (3polFRET) for interactions at the Förster-distance and beyond. BIOCHIMICA ET BIOPHYSICA ACTA-MOLECULAR CELL RESEARCH 2016; 1863:703-16. [DOI: 10.1016/j.bbamcr.2016.02.002] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.4] [Reference Citation Analysis] [Track Full Text] [Subscribe] [Scholar Register] [Received: 12/18/2015] [Revised: 02/01/2016] [Accepted: 02/03/2016] [Indexed: 11/28/2022]
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44
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Best RB, Hofmann H, Nettels D, Schuler B. Quantitative interpretation of FRET experiments via molecular simulation: force field and validation. Biophys J 2016; 108:2721-31. [PMID: 26039173 DOI: 10.1016/j.bpj.2015.04.038] [Citation(s) in RCA: 51] [Impact Index Per Article: 6.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/03/2015] [Revised: 04/23/2015] [Accepted: 04/27/2015] [Indexed: 12/14/2022] Open
Abstract
Molecular simulation is a valuable and complementary tool that may assist with the interpretation of single-molecule Förster resonance energy transfer (FRET) experiments, if the energy function is of sufficiently high quality. Here we present force-field parameters for one of the most common pairs of chromophores used in experiments, AlexaFluor 488 and 594. From microsecond molecular-dynamics simulations, we are able to recover both experimentally determined equilibrium constants and association/dissociation rates of the chromophores with free tryptophan, as well as the decay of fluorescence anisotropy of a labeled protein. We find that it is particularly important to obtain a correct balance of solute-water interactions in the simulations in order to faithfully capture the experimental anisotropy decays, which provide a sensitive benchmark for fluorophore mobility. Lastly, by a combination of experiment and simulation, we address a potential complication in the interpretation of experiments on polyproline, used as a molecular ruler for FRET experiments, namely the potential association of one of the chromophores with the polyproline helix. Under conditions where simulations accurately capture the fluorescence anisotropy decay, we find at most a modest, transient population of conformations in which the chromophores associate with the polyproline. Explicit calculation of FRET transfer efficiencies for short polyprolines yields results in good agreement with experiment. These results illustrate the potential power of a combination of molecular simulation and experiment in quantifying biomolecular dynamics.
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Affiliation(s)
- Robert B Best
- Laboratory of Chemical Physics, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, Maryland.
| | - Hagen Hofmann
- Department of Biochemistry, University of Zurich, Zurich, Switzerland
| | - Daniel Nettels
- Department of Biochemistry, University of Zurich, Zurich, Switzerland
| | - Benjamin Schuler
- Department of Biochemistry, University of Zurich, Zurich, Switzerland.
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45
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Dawidowski D, Cafiso DS. Munc18-1 and the Syntaxin-1 N Terminus Regulate Open-Closed States in a t-SNARE Complex. Structure 2016; 24:392-400. [PMID: 26876096 DOI: 10.1016/j.str.2016.01.005] [Citation(s) in RCA: 23] [Impact Index Per Article: 2.9] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/15/2015] [Revised: 12/29/2015] [Accepted: 01/08/2016] [Indexed: 10/22/2022]
Abstract
Neuronal exocytosis is mediated by SNARE proteins, which assemble into a highly stable four-helical bundle in a process that is not well understood. Here, electron paramagnetic resonance spectroscopy was used to examine how the t-SNAREs syntaxin and SNAP25 assemble in the presence and absence of the regulatory protein Munc18-1. Syntaxin and SNAP25 form a 2:1 complex, which is structurally heterogeneous and persists in the presence of excess SNAP25. Munc18-1 dissociates this 2:1 complex, but a 1:1 complex is retained where syntaxin is in a closed state. In the absence of an N-terminal fragment of syntaxin, Munc18-1 also stabilizes a 1:1 complex of sytaxin/SNAP25; however, syntaxin now samples an open state. These data demonstrate that the open-closed syntaxin equilibrium is shifted toward the open state when syntaxin and Munc18-1 are associated with SNAP25, and the results indicate that a syntaxin/SNAP25:Munc18-1 complex is a likely starting point for SNARE assembly.
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Affiliation(s)
- Damian Dawidowski
- Department of Chemistry, Center for Membrane Biology at the University of Virginia, McCormick Road, Charlottesville, VA 22904-4319, USA
| | - David S Cafiso
- Department of Chemistry, Center for Membrane Biology at the University of Virginia, McCormick Road, Charlottesville, VA 22904-4319, USA.
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46
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Rahamim G, Chemerovski-Glikman M, Rahimipour S, Amir D, Haas E. Resolution of Two Sub-Populations of Conformers and Their Individual Dynamics by Time Resolved Ensemble Level FRET Measurements. PLoS One 2015; 10:e0143732. [PMID: 26699718 PMCID: PMC4689530 DOI: 10.1371/journal.pone.0143732] [Citation(s) in RCA: 12] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/07/2015] [Accepted: 11/08/2015] [Indexed: 11/19/2022] Open
Abstract
Most active biopolymers are dynamic structures; thus, ensembles of such molecules should be characterized by distributions of intra- or intermolecular distances and their fast fluctuations. A method of choice to determine intramolecular distances is based on Förster resonance energy transfer (FRET) measurements. Major advances in such measurements were achieved by single molecule FRET measurements. Here, we show that by global analysis of the decay of the emission of both the donor and the acceptor it is also possible to resolve two sub-populations in a mixture of two ensembles of biopolymers by time resolved FRET (trFRET) measurements at the ensemble level. We show that two individual intramolecular distance distributions can be determined and characterized in terms of their individual means, full width at half maximum (FWHM), and two corresponding diffusion coefficients which reflect the rates of fast ns fluctuations within each sub-population. An important advantage of the ensemble level trFRET measurements is the ability to use low molecular weight small-sized probes and to determine nanosecond fluctuations of the distance between the probes. The limits of the possible resolution were first tested by simulation and then by preparation of mixtures of two model peptides. The first labeled polypeptide was a relatively rigid Pro7 and the second polypeptide was a flexible molecule consisting of (Gly-Ser)7 repeats. The end to end distance distributions and the diffusion coefficients of each peptide were determined. Global analysis of trFRET measurements of a series of mixtures of polypeptides recovered two end-to-end distance distributions and associated intramolecular diffusion coefficients, which were very close to those determined from each of the pure samples. This study is a proof of concept study demonstrating the power of ensemble level trFRET based methods in resolution of subpopulations in ensembles of flexible macromolecules.
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Affiliation(s)
- Gil Rahamim
- The Goodman Faculty of Life Sciences Bar Ilan University, Ramat Gan Israel 52900
| | | | - Shai Rahimipour
- Department of Chemistry, Bar-Ilan University, Ramat Gan Israel 52900
| | - Dan Amir
- The Goodman Faculty of Life Sciences Bar Ilan University, Ramat Gan Israel 52900
| | - Elisha Haas
- The Goodman Faculty of Life Sciences Bar Ilan University, Ramat Gan Israel 52900
- * E-mail:
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47
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Wennmalm S, Chmyrov V, Widengren J, Tjernberg L. Highly Sensitive FRET-FCS Detects Amyloid β-Peptide Oligomers in Solution at Physiological Concentrations. Anal Chem 2015; 87:11700-5. [PMID: 26489794 DOI: 10.1021/acs.analchem.5b02630] [Citation(s) in RCA: 34] [Impact Index Per Article: 3.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/28/2022]
Abstract
Oligomers formed by the amyloid β-peptide (Aβ) are pathogens in Alzheimer's disease. Increased knowledge on the oligomerization process is crucial for understanding the disease and for finding treatments. Ideally, Aβ oligomerization should be studied in solution and at physiologically relevant concentrations, but most popular techniques of today are not capable of such analyses. We demonstrate here that the combination of Förster Resonance Energy Transfer and Fluorescence Correlation Spectroscopy (FRET-FCS) has a unique ability to detect small subpopulations of FRET-active molecules and oligomers. FRET-FCS could readily detect a FRET-active oligonucleotide present at levels as low as 0.5% compared to FRET-inactive dye molecules. In contrast, three established fluorescence fluctuation techniques (FCS, FCCS, and PCH) required fractions between 7 and 11%. When applied to the analysis of Aβ, FRET-FCS detected oligomers consisting of less than 10 Aβ molecules, which coexisted with the monomers at fractions as low as 2 ± 2%. Thus, we demonstrate for the first time direct detection of small fractions of Aβ oligomers in solution at physiological concentrations. This ability of FRET-FCS could be an indispensable tool for studying biological oligomerization processes, in general, and for finding therapeutically useful oligomerization inhibitors.
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Affiliation(s)
- Stefan Wennmalm
- Royal Institute of Technology-KTH, Department of Applied Physics, Experimental Biomolecular Physics Group, Scilifelab , 171 65 Solna, Sweden
| | - Volodymyr Chmyrov
- Royal Institute of Technology-KTH , Department of Applied Physics, Experimental Biomolecular Physics Group, AlbaNova, 106 91 Stockholm, Sweden
| | - Jerker Widengren
- Royal Institute of Technology-KTH , Department of Applied Physics, Experimental Biomolecular Physics Group, AlbaNova, 106 91 Stockholm, Sweden
| | - Lars Tjernberg
- Karolinska Institutet , Department of NVS, Center for Alzheimer Research, Division for Neurogeriatrics, Novum plan 5, 141 86 Stockholm, Sweden
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48
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Khrenova M, Topol I, Collins J, Nemukhin A. Estimating orientation factors in the FRET theory of fluorescent proteins: the TagRFP-KFP pair and beyond. Biophys J 2015; 108:126-32. [PMID: 25564859 DOI: 10.1016/j.bpj.2014.11.1859] [Citation(s) in RCA: 21] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/07/2014] [Revised: 10/20/2014] [Accepted: 11/06/2014] [Indexed: 11/29/2022] Open
Abstract
The orientation factor κ(2), one of the key parameters defining Förster resonance energy transfer efficiency, is determined by the transition dipole moment orientations of the donor and acceptor species. Using the results of quantum chemical and quantum mechanical/molecular mechanical calculations for the chromophore-containing pockets in selected colored proteins of the green fluorescent protein family, we derived transition dipole moments corresponding to the S0,min → S1 excitation for green fluorescent protein, red fluorescent protein (TagRFP), and kindling fluorescent protein, and the S1,min → S0 emission for TagRFP. These data allowed us to estimate κ(2) values for the TagRFP-linker-kindling fluorescent protein tetrameric complex required for constructing novel sensors.
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Affiliation(s)
- Maria Khrenova
- Department of Chemistry, Lomonosov Moscow State University, Moscow, Russian Federation
| | - Igor Topol
- Advanced Biomedical Computing Center, Information Systems Program, Leidos Biomedical Research, Inc., Frederick National Laboratory for Cancer Research, Frederick, Maryland.
| | - Jack Collins
- Advanced Biomedical Computing Center, Information Systems Program, Leidos Biomedical Research, Inc., Frederick National Laboratory for Cancer Research, Frederick, Maryland
| | - Alexander Nemukhin
- Department of Chemistry, Lomonosov Moscow State University, Moscow, Russian Federation; Emanuel Institute of Biochemical Physics, Moscow, Russian Federation
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49
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Ullrich A, Böhme MA, Schöneberg J, Depner H, Sigrist SJ, Noé F. Dynamical Organization of Syntaxin-1A at the Presynaptic Active Zone. PLoS Comput Biol 2015; 11:e1004407. [PMID: 26367029 PMCID: PMC4569342 DOI: 10.1371/journal.pcbi.1004407] [Citation(s) in RCA: 52] [Impact Index Per Article: 5.8] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/22/2014] [Accepted: 06/15/2015] [Indexed: 01/10/2023] Open
Abstract
Synaptic vesicle fusion is mediated by SNARE proteins forming in between synaptic vesicle (v-SNARE) and plasma membrane (t-SNARE), one of which is Syntaxin-1A. Although exocytosis mainly occurs at active zones, Syntaxin-1A appears to cover the entire neuronal membrane. By using STED super-resolution light microscopy and image analysis of Drosophila neuro-muscular junctions, we show that Syntaxin-1A clusters are more abundant and have an increased size at active zones. A computational particle-based model of syntaxin cluster formation and dynamics is developed. The model is parametrized to reproduce Syntaxin cluster-size distributions found by STED analysis, and successfully reproduces existing FRAP results. The model shows that the neuronal membrane is adjusted in a way to strike a balance between having most syntaxins stored in large clusters, while still keeping a mobile fraction of syntaxins free or in small clusters that can efficiently search the membrane or be traded between clusters. This balance is subtle and can be shifted toward almost no clustering and almost complete clustering by modifying the syntaxin interaction energy on the order of only 1 kBT. This capability appears to be exploited at active zones. The larger active-zone syntaxin clusters are more stable and provide regions of high docking and fusion capability, whereas the smaller clusters outside may serve as flexible reserve pool or sites of spontaneous ectopic release.
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Affiliation(s)
- Alexander Ullrich
- Department of Mathematics, Freie Universität Berlin, Berlin, Germany
| | - Mathias A. Böhme
- Institute for Biology/Genetics, Freie Universität Berlin, Berlin, Germany
- NeuroCure Cluster of Excellence, Charité Berlin, Berlin, Germany
| | | | - Harald Depner
- Institute for Biology/Genetics, Freie Universität Berlin, Berlin, Germany
| | - Stephan J. Sigrist
- Institute for Biology/Genetics, Freie Universität Berlin, Berlin, Germany
- NeuroCure Cluster of Excellence, Charité Berlin, Berlin, Germany
| | - Frank Noé
- Department of Mathematics, Freie Universität Berlin, Berlin, Germany
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Shoura MJ, Ranatunga RJKU, Harris SA, Nielsen SO, Levene SD. Contribution of fluorophore dynamics and solvation to resonant energy transfer in protein-DNA complexes: a molecular-dynamics study. Biophys J 2015; 107:700-710. [PMID: 25099809 DOI: 10.1016/j.bpj.2014.06.023] [Citation(s) in RCA: 13] [Impact Index Per Article: 1.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/20/2013] [Revised: 06/09/2014] [Accepted: 06/10/2014] [Indexed: 01/23/2023] Open
Abstract
In Förster resonance energy transfer (FRET) experiments, extracting accurate structural information about macromolecules depends on knowing the positions and orientations of donor and acceptor fluorophores. Several approaches have been employed to reduce uncertainties in quantitative FRET distance measurements. Fluorophore-position distributions can be estimated by surface accessibility (SA) calculations, which compute the region of space explored by the fluorophore within a static macromolecular structure. However, SA models generally do not take fluorophore shape, dye transition-moment orientation, or dye-specific chemical interactions into account. We present a detailed molecular-dynamics (MD) treatment of fluorophore dynamics for an ATTO donor/acceptor dye pair and specifically consider as case studies dye-labeled protein-DNA intermediates in Cre site-specific recombination. We carried out MD simulations in both an aqueous solution and glycerol/water mixtures to assess the effects of experimental solvent systems on dye dynamics. Our results unequivocally show that MD simulations capture solvent effects and dye-dye interactions that can dramatically affect energy transfer efficiency. We also show that results from SA models and MD simulations strongly diverge in cases where donor and acceptor fluorophores are in close proximity. Although atomistic simulations are computationally more expensive than SA models, explicit MD studies are likely to give more realistic results in both homogeneous and mixed solvents. Our study underscores the model-dependent nature of FRET analyses, but also provides a starting point to develop more realistic in silico approaches for obtaining experimental ensemble and single-molecule FRET data.
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Affiliation(s)
- Massa J Shoura
- Department of Bioengineering, University of Texas at Dallas, Richardson, Texas; Department of Molecular and Cell Biology, University of Texas at Dallas, Richardson, Texas
| | | | - Sarah A Harris
- Department of Physics and Astronomy, University of Leeds, Leeds, UK
| | - Steven O Nielsen
- Department of Chemistry, University of Texas at Dallas, Richardson, Texas
| | - Stephen D Levene
- Department of Bioengineering, University of Texas at Dallas, Richardson, Texas; Department of Molecular and Cell Biology, University of Texas at Dallas, Richardson, Texas; Department of Physics, University of Texas at Dallas, Richardson, Texas.
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