1
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Pati AK, Kilic Z, Martin MI, Terry DS, Borgia A, Bar S, Jockusch S, Kiselev R, Altman RB, Blanchard SC. Recovering true FRET efficiencies from smFRET investigations requires triplet state mitigation. Nat Methods 2024:10.1038/s41592-024-02293-8. [PMID: 38877317 DOI: 10.1038/s41592-024-02293-8] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/29/2023] [Accepted: 04/25/2024] [Indexed: 06/16/2024]
Abstract
Single-molecule fluorescence resonance energy transfer (smFRET) methods employed to quantify time-dependent compositional and conformational changes within biomolecules require elevated illumination intensities to recover robust photon emission streams from individual fluorophores. Here we show that outside the weak-excitation limit, and in regimes where fluorophores must undergo many rapid cycles of excitation and relaxation, non-fluorescing, excitation-induced triplet states with lifetimes orders of magnitude longer lived than photon-emitting singlet states degrade photon emission streams from both donor and acceptor fluorophores resulting in illumination-intensity-dependent changes in FRET efficiency. These changes are not commonly taken into consideration; therefore, robust strategies to suppress excited state accumulations are required to recover accurate and precise FRET efficiency, and thus distance, estimates. We propose both robust triplet state suppression and data correction strategies that enable the recovery of FRET efficiencies more closely approximating true values, thereby extending the spatial and temporal resolution of smFRET.
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Affiliation(s)
- Avik K Pati
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
- Department of Chemistry, Birla Institute of Technology and Science, Pilani, Rajasthan, India
| | - Zeliha Kilic
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
| | - Maxwell I Martin
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
- Department of Chemical Biology & Therapeutics, St. Jude Children's Research Hospital, Memphis, TN, USA
| | - Daniel S Terry
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
| | - Alessandro Borgia
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
| | - Sukanta Bar
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
- Department of Chemical Biology & Therapeutics, St. Jude Children's Research Hospital, Memphis, TN, USA
| | - Steffen Jockusch
- Center for Photochemical Sciences and Department of Chemistry, Bowling Green State University, Bowling Green, OH, USA
| | - Roman Kiselev
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
| | - Roger B Altman
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
- Department of Chemical Biology & Therapeutics, St. Jude Children's Research Hospital, Memphis, TN, USA
| | - Scott C Blanchard
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA.
- Department of Chemical Biology & Therapeutics, St. Jude Children's Research Hospital, Memphis, TN, USA.
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2
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Mooren OL, Stuchell-Brereton MD, McConnell P, Yan C, Wilkerson EM, Goldfarb D, Cooper JA, Sept D, Soranno A. Biophysical Mechanism of Allosteric Regulation of Actin Capping Protein. J Mol Biol 2023; 435:168342. [PMID: 37924863 PMCID: PMC10872493 DOI: 10.1016/j.jmb.2023.168342] [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: 08/13/2023] [Revised: 10/28/2023] [Accepted: 10/30/2023] [Indexed: 11/06/2023]
Abstract
Actin capping protein (CP) can be regulated by steric and allosteric mechanisms. The molecular mechanism of the allosteric regulation at a biophysical level includes linkage between the binding sites for three ligands: F-actin, Capping-Protein-Interacting (CPI) motifs, and V-1/myotrophin, based on biochemical functional studies and solvent accessibility experiments. Here, we investigated the mechanism of allosteric regulation at the atomic level using single-molecule Förster resonance energy transfer (FRET) and molecular dynamics (MD) to assess the conformational and structural dynamics of CP in response to linked-binding site ligands. In the absence of ligand, both single-molecule FRET and MD revealed two distinct conformations of CP in solution; previous crystallographic studies revealed only one. Interaction with CPI-motif peptides induced conformations within CP that bring the cap and stalk closer, while interaction with V-1 moves them away from one another. Comparing CPI-motif peptides from different proteins, we identified variations in CP conformations and dynamics that are specific to each CPI motif. MD simulations for CP alone and in complex with a CPI motif and V-1 reveal atomistic details of the conformational changes. Analysis of the interaction of CP with wild-type (wt) and chimeric CPI-motif peptides using single-molecule FRET, isothermal calorimetry (ITC) and MD simulation indicated that conformational and affinity differences are intrinsic to the C-terminal portion of the CPI motif. We conclude that allosteric regulation of CP involves changes in conformation that disseminate across the protein to link distinct binding-site functions. Our results provide novel insights into the biophysical mechanism of the allosteric regulation of CP.
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Affiliation(s)
- Olivia L Mooren
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, United States
| | - Melissa D Stuchell-Brereton
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, United States; Center for Biomolecular Condensates, Washington University in St Louis, St. Louis, MO, United States
| | - Patrick McConnell
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, United States
| | - Chenbo Yan
- Department of Biophysics, University of Michigan, Ann Arbor, MI, United States
| | - Emily M Wilkerson
- Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, MO, United States; Institute for Informatics, Washington University School of Medicine, St. Louis, MO, United States
| | - Dennis Goldfarb
- Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, MO, United States; Institute for Informatics, Washington University School of Medicine, St. Louis, MO, United States
| | - John A Cooper
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, United States.
| | - David Sept
- Department of Biophysics, University of Michigan, Ann Arbor, MI, United States; Department of Biomedical Engineering, University of Michigan, Ann Arbor, MI, United States.
| | - Andrea Soranno
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, United States; Center for Biomolecular Condensates, Washington University in St Louis, St. Louis, MO, United States.
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3
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Sohmen B, Beck C, Frank V, Seydel T, Hoffmann I, Hermann B, Nüesch M, Grimaldo M, Schreiber F, Wolf S, Roosen‐Runge F, Hugel T. The Onset of Molecule-Spanning Dynamics in Heat Shock Protein Hsp90. ADVANCED SCIENCE (WEINHEIM, BADEN-WURTTEMBERG, GERMANY) 2023; 10:e2304262. [PMID: 37984887 PMCID: PMC10754087 DOI: 10.1002/advs.202304262] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 06/26/2023] [Revised: 10/06/2023] [Indexed: 11/22/2023]
Abstract
Protein dynamics have been investigated on a wide range of time scales. Nano- and picosecond dynamics have been assigned to local fluctuations, while slower dynamics have been attributed to larger conformational changes. However, it is largely unknown how fast (local) fluctuations can lead to slow global (allosteric) changes. Here, fast molecule-spanning dynamics on the 100 to 200 ns time scale in the heat shock protein 90 (Hsp90) are shown. Global real-space movements are assigned to dynamic modes on this time scale, which is possible by a combination of single-molecule fluorescence, quasi-elastic neutron scattering and all-atom molecular dynamics (MD) simulations. The time scale of these dynamic modes depends on the conformational state of the Hsp90 dimer. In addition, the dynamic modes are affected to various degrees by Sba1, a co-chaperone of Hsp90, depending on the location within Hsp90, which is in very good agreement with MD simulations. Altogether, this data is best described by fast molecule-spanning dynamics, which precede larger conformational changes in Hsp90 and might be the molecular basis for allostery. This integrative approach provides comprehensive insights into molecule-spanning dynamics on the nanosecond time scale for a multi-domain protein.
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Affiliation(s)
- Benedikt Sohmen
- Institute of Physical ChemistryUniversity of FreiburgAlbertstrasse 2179104FreiburgGermany
| | - Christian Beck
- Institute of Applied PhysicsUniversity of TübingenAuf der Morgenstelle 1072076TübingenGermany
- Science DivisionInstitut Max von Laue ‐ Paul Langevin71 avenue des MartyrsGrenoble38042France
| | - Veronika Frank
- Institute of Physical ChemistryUniversity of FreiburgAlbertstrasse 2179104FreiburgGermany
| | - Tilo Seydel
- Science DivisionInstitut Max von Laue ‐ Paul Langevin71 avenue des MartyrsGrenoble38042France
| | - Ingo Hoffmann
- Science DivisionInstitut Max von Laue ‐ Paul Langevin71 avenue des MartyrsGrenoble38042France
| | - Bianca Hermann
- Institute of Physical ChemistryUniversity of FreiburgAlbertstrasse 2179104FreiburgGermany
| | - Mark Nüesch
- Department of BiochemistryUniversity of ZurichWinterthurerstrasse 190CH‐8057ZurichSwitzerland
| | - Marco Grimaldo
- Science DivisionInstitut Max von Laue ‐ Paul Langevin71 avenue des MartyrsGrenoble38042France
| | - Frank Schreiber
- Institute of Applied PhysicsUniversity of TübingenAuf der Morgenstelle 1072076TübingenGermany
| | - Steffen Wolf
- Biomolecular Dynamics, Institute of PhysicsUniversity of FreiburgHermann‐Herder‐Strasse 379104FreiburgGermany
| | - Felix Roosen‐Runge
- Department of Biomedical Sciences and Biofilms‐Research Center for Biointerfaces (BRCB)Malmö University20506MalmöSweden
- Division of Physical ChemistryLund UniversityNaturvetarvägen 1422100LundSweden
| | - Thorsten Hugel
- Institute of Physical ChemistryUniversity of FreiburgAlbertstrasse 2179104FreiburgGermany
- Signalling Research Centers BIOSS and CIBSSUniversity of FreiburgSchänzlestrasse 1879104FreiburgGermany
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4
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Mooren OL, Stuchell-Brereton MD, McConnell P, Yan C, Wilkerson EM, Goldfarb D, Cooper JA, Sept D, Soranno A. Biophysical Mechanism of Allosteric Regulation of Actin Capping Protein. BIORXIV : THE PREPRINT SERVER FOR BIOLOGY 2023:2023.08.16.553570. [PMID: 37645735 PMCID: PMC10462145 DOI: 10.1101/2023.08.16.553570] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 08/31/2023]
Abstract
Actin capping protein (CP) can be regulated by steric and allosteric mechanisms. The molecular mechanism of the allosteric regulation at a biophysical level includes linkage between the binding sites for three ligands: F-actin, Capping-Protein-Interacting (CPI) motifs, and V-1/myotrophin, based on biochemical functional studies and solvent accessibility experiments. Here, we investigated the mechanism of allosteric regulation at the atomic level using single-molecule Förster resonance energy transfer (FRET) and molecular dynamics (MD) to assess the conformational and structural dynamics of CP in response to linked-binding site ligands. In the absence of ligand, both single-molecule FRET and MD revealed two distinct conformations of CP in solution; previous crystallographic studies revealed only one. CPI-motif peptide association induced conformational changes within CP that propagate in one direction, while V-1 association induced conformational changes in the opposite direction. Comparing CPI-motif peptides from different proteins, we identified variations in CP conformations and dynamics that are specific to each CPI motif. MD simulations for CP alone and in complex with a CPI motif and V-1 reveal atomistic details of the conformational changes. Analysis of the interaction of CP with wildtype (wt) and chimeric CPI-motif peptides using single-molecule FRET, isothermal calorimetry (ITC) and MD simulation indicated that conformational and affinity differences are intrinsic to the C-terminal portion of the CPI-motif. We conclude that allosteric regulation of CP involves changes in conformation that disseminate across the protein to link distinct binding-site functions. Our results provide novel insights into the biophysical mechanism of the allosteric regulation of CP.
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5
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Harris PD, Lerner E. Identification and quantification of within-burst dynamics in singly labeled single-molecule fluorescence lifetime experiments. BIOPHYSICAL REPORTS 2022; 2. [PMID: 36204594 PMCID: PMC9534301 DOI: 10.1016/j.bpr.2022.100071] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Subscribe] [Scholar Register] [Indexed: 10/27/2022]
Abstract
Single-molecule spectroscopy has revolutionized molecular biophysics and provided means to probe how structural moieties within biomolecules spatially reorganize at different timescales. There are several single-molecule methodologies that probe local structural dynamics in the vicinity of a single dye-labeled residue, which rely on fluorescence lifetimes as readout. Nevertheless, an analytical framework to quantify dynamics in such single-molecule single dye fluorescence bursts, at timescales of microseconds to milliseconds, has not yet been demonstrated. Here, we suggest an analytical framework for identifying and quantifying within-burst lifetime-based dynamics, such as conformational dynamics recorded in single-molecule photo-isomerization-related fluorescence enhancement. After testing the capabilities of the analysis on simulations, we proceed to exhibit within-burst millisecond local structural dynamics in the unbound α-synuclein monomer. The analytical framework provided in this work paves the way for extracting a full picture of the energy landscape for the coordinate probed by fluorescence lifetime-based single-molecule measurements.
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Affiliation(s)
- Paul David Harris
- Department of Biological Chemistry, The Alexander Silberman Institute of Life Sciences, Faculty of Mathematics & Science, The Edmond J. Safra Campus, The Hebrew University of Jerusalem, Jerusalem 9190401, Israel
| | - Eitan Lerner
- Department of Biological Chemistry, The Alexander Silberman Institute of Life Sciences, Faculty of Mathematics & Science, The Edmond J. Safra Campus, The Hebrew University of Jerusalem, Jerusalem 9190401, Israel.,The Center for Nanoscience and Nanotechnology, The Hebrew University of Jerusalem, Jerusalem 9190401, Israel
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6
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Abyzov A, Blackledge M, Zweckstetter M. Conformational Dynamics of Intrinsically Disordered Proteins Regulate Biomolecular Condensate Chemistry. Chem Rev 2022; 122:6719-6748. [PMID: 35179885 PMCID: PMC8949871 DOI: 10.1021/acs.chemrev.1c00774] [Citation(s) in RCA: 36] [Impact Index Per Article: 18.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/14/2022]
Abstract
![]()
Motions in biomolecules
are critical for biochemical reactions.
In cells, many biochemical reactions are executed inside of biomolecular
condensates formed by ultradynamic intrinsically disordered proteins.
A deep understanding of the conformational dynamics of intrinsically
disordered proteins in biomolecular condensates is therefore of utmost
importance but is complicated by diverse obstacles. Here we review
emerging data on the motions of intrinsically disordered proteins
inside of liquidlike condensates. We discuss how liquid–liquid
phase separation modulates internal motions across a wide range of
time and length scales. We further highlight the importance of intermolecular
interactions that not only drive liquid–liquid phase separation
but appear as key determinants for changes in biomolecular motions
and the aging of condensates in human diseases. The review provides
a framework for future studies to reveal the conformational dynamics
of intrinsically disordered proteins in the regulation of biomolecular
condensate chemistry.
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Affiliation(s)
- Anton Abyzov
- Translational Structural Biology Group, German Center for Neurodegenerative Diseases (DZNE), 37075 Göttingen, Germany
| | - Martin Blackledge
- Université Grenoble Alpes, Institut de Biologie Structurale (IBS), 38044 Grenoble, France.,CEA, DSV, IBS, 38044 Grenoble, France.,CNRS, IBS, 38044 Grenoble, France
| | - Markus Zweckstetter
- Translational Structural Biology Group, German Center for Neurodegenerative Diseases (DZNE), 37075 Göttingen, Germany.,Department for NMR-based Structural Biology, Max Planck Institute for Biophysical Chemistry, 37077 Göttingen, Germany
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7
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Zosel F, Holla A, Schuler B. Labeling of Proteins for Single-Molecule Fluorescence Spectroscopy. Methods Mol Biol 2022; 2376:207-233. [PMID: 34845612 DOI: 10.1007/978-1-0716-1716-8_12] [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] [Indexed: 06/13/2023]
Abstract
Single-molecule fluorescence spectroscopy has become an important technique for studying the conformational dynamics and folding of proteins. A key step for performing such experiments is the availability of high-quality samples. This chapter describes a simple and widely applicable strategy for preparing proteins that are site-specifically labeled with a donor and an acceptor dye for single-molecule Förster resonance energy transfer (FRET) experiments. The method is based on introducing two cysteine residues that are labeled with maleimide-functionalized fluorophores, combined with high-resolution chromatography. We discuss how to optimize site-specific labeling even in the absence of orthogonal coupling chemistry and present purification strategies that are suitable for samples ranging from intrinsically disordered proteins to large folded proteins. We also discuss common problems in protein labeling, how to avoid them, and how to stringently control sample quality.
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Affiliation(s)
- Franziska Zosel
- Department of Biochemistry, University of Zurich, Zurich, Switzerland
- Novo Nordisk A/S, Måløv, Denmark
| | - Andrea Holla
- Department of Biochemistry, University of Zurich, Zurich, Switzerland
| | - Benjamin Schuler
- Department of Biochemistry, University of Zurich, Zurich, Switzerland.
- Department of Physics, University of Zurich, Zurich, Switzerland.
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8
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Gopich IV, Chung HS. Theory and Analysis of Single-Molecule FRET Experiments. Methods Mol Biol 2022; 2376:247-282. [PMID: 34845614 DOI: 10.1007/978-1-0716-1716-8_14] [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] [Indexed: 06/13/2023]
Abstract
Inter-dye distances and conformational dynamics can be studied using single-molecule FRET measurements. We consider two approaches to analyze sequences of photons with recorded photon colors and arrival times. The first approach is based on FRET efficiency histograms obtained from binned photon sequences. The experimental histograms are compared with the theoretical histograms obtained using the joint distribution of acceptor and donor photons or the Gaussian approximation. In the second approach, a photon sequence is analyzed without binning. The parameters of a model describing conformational dynamics are found by maximizing the appropriate likelihood function. The first approach is simpler, while the second one is more accurate, especially when the population of species is small and transition rates are fast. The likelihood-based analysis as well as the recoloring method has the advantage that diffusion of molecules through the laser focus can be rigorously handled.
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Affiliation(s)
- Irina V Gopich
- Laboratory of Chemical Physics, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, MD, USA.
| | - Hoi Sung Chung
- Laboratory of Chemical Physics, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, MD, USA
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9
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Nüesch MF, Ivanović MT, Claude JB, Nettels D, Best RB, Wenger J, Schuler B. Single-molecule Detection of Ultrafast Biomolecular Dynamics with Nanophotonics. J Am Chem Soc 2021; 144:52-56. [PMID: 34970909 DOI: 10.1021/jacs.1c09387] [Citation(s) in RCA: 9] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/20/2023]
Abstract
Single-molecule Förster resonance energy transfer (FRET) is a versatile technique for probing the structure and dynamics of biomolecules even in heterogeneous ensembles. However, because of the limited fluorescence brightness per molecule and the relatively long fluorescence lifetimes, probing ultrafast structural dynamics in the nanosecond time scale has thus far been very challenging. Here, we demonstrate that nanophotonic fluorescence enhancement in zero-mode waveguides enables measurements of previously inaccessible low-nanosecond dynamics by dramatically improving time resolution and reduces data acquisition times by more than an order of magnitude. As a prototypical example, we use this approach to probe the dynamics of a short intrinsically disordered peptide that were previously inaccessible with single-molecule FRET measurements. We show that we are now able to detect the low-nanosecond correlations in this peptide, and we obtain a detailed interpretation of the underlying distance distributions and dynamics in conjunction with all-atom molecular dynamics simulations, which agree remarkably well with the experiments. We expect this combined approach to be widely applicable to the investigation of very rapid biomolecular dynamics.
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Affiliation(s)
- Mark F Nüesch
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Miloš T Ivanović
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Jean-Benoît Claude
- Aix Marseille Univ, CNRS, Centrale Marseille, Institut Fresnel, 13013 Marseille, France
| | - Daniel Nettels
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Robert B Best
- Laboratory of Chemical Physics, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, Maryland 20892-0520, United States
| | - Jérôme Wenger
- Aix Marseille Univ, CNRS, Centrale Marseille, Institut Fresnel, 13013 Marseille, France
| | - Benjamin Schuler
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland.,Department of Physics, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
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10
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Klose D, Holla A, Gmeiner C, Nettels D, Ritsch I, Bross N, Yulikov M, Allain FHT, Schuler B, Jeschke G. Resolving distance variations by single-molecule FRET and EPR spectroscopy using rotamer libraries. Biophys J 2021; 120:4842-4858. [PMID: 34536387 PMCID: PMC8595751 DOI: 10.1016/j.bpj.2021.09.021] [Citation(s) in RCA: 15] [Impact Index Per Article: 5.0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/26/2021] [Revised: 07/09/2021] [Accepted: 09/13/2021] [Indexed: 01/14/2023] Open
Abstract
Förster resonance energy transfer (FRET) and electron paramagnetic resonance (EPR) spectroscopy are complementary techniques for quantifying distances in the nanometer range. Both approaches are commonly employed for probing the conformations and conformational changes of biological macromolecules based on site-directed fluorescent or paramagnetic labeling. FRET can be applied in solution at ambient temperature and thus provides direct access to dynamics, especially if used at the single-molecule level, whereas EPR requires immobilization or work at cryogenic temperatures but provides data that can be more reliably used to extract distance distributions. However, a combined analysis of the complementary data from the two techniques has been complicated by the lack of a common modeling framework. Here, we demonstrate a systematic analysis approach based on rotamer libraries for both FRET and EPR labels to predict distance distributions between two labels from a structural model. Dynamics of the fluorophores within these distance distributions are taken into account by diffusional averaging, which improves the agreement with experiment. Benchmarking this methodology with a series of surface-exposed pairs of sites in a structured protein domain reveals that the lowest resolved distance differences can be as small as ∼0.25 nm for both techniques, with quantitative agreement between experimental and simulated transfer efficiencies within a range of ±0.045. Rotamer library analysis thus establishes a coherent way of treating experimental data from EPR and FRET and provides a basis for integrative structural modeling, including studies of conformational distributions and dynamics of biological macromolecules using both techniques.
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Affiliation(s)
- Daniel Klose
- Department of Chemistry and Applied Biosciences, ETH Zurich, Zurich, Switzerland.
| | - Andrea Holla
- Department of Biochemistry, University of Zurich, Zurich, Switzerland
| | - Christoph Gmeiner
- Department of Chemistry and Applied Biosciences, ETH Zurich, Zurich, Switzerland
| | - Daniel Nettels
- Department of Biochemistry, University of Zurich, Zurich, Switzerland
| | - Irina Ritsch
- Department of Chemistry and Applied Biosciences, ETH Zurich, Zurich, Switzerland
| | - Nadja Bross
- Department of Chemistry, University of Zurich, Zurich, Switzerland
| | - Maxim Yulikov
- Department of Chemistry and Applied Biosciences, ETH Zurich, Zurich, Switzerland
| | | | - Benjamin Schuler
- Department of Biochemistry, University of Zurich, Zurich, Switzerland; Department of Physics, University of Zurich, Zurich, Switzerland.
| | - Gunnar Jeschke
- Department of Chemistry and Applied Biosciences, ETH Zurich, Zurich, Switzerland
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11
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Abstract
Flexibility in complexes between intrinsically disordered proteins and folded ligands is widespread in nature. However, timescales and spatial amplitudes of such dynamics remained unexplored for most systems. Our results show that the disordered cytoplasmic tail of the cell adhesion protein E-cadherin diffuses across the entire surface of its folded binding partner β-catenin at fast submillisecond timescales. The nanometer amplitude of these motions could allow kinases to access their recognition motifs without requiring a dissociation of the complex. We expect that the rugged energy landscape found in the E-cadherin/β-catenin complex is a defining feature of dynamic and partially disordered protein complexes. Intrinsically disordered proteins often form dynamic complexes with their ligands. Yet, the speed and amplitude of these motions are hidden in classical binding kinetics. Here, we directly measure the dynamics in an exceptionally mobile, high-affinity complex. We show that the disordered tail of the cell adhesion protein E-cadherin dynamically samples a large surface area of the protooncogene β-catenin. Single-molecule experiments and molecular simulations resolve these motions with high resolution in space and time. Contacts break and form within hundreds of microseconds without a dissociation of the complex. The energy landscape of this complex is rugged with many small barriers (3 to 4 kBT) and reconciles specificity, high affinity, and extreme disorder. A few persistent contacts provide specificity, whereas unspecific interactions boost affinity.
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12
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Dingfelder F, Macocco I, Benke S, Nettels D, Faccioli P, Schuler B. Slow Escape from a Helical Misfolded State of the Pore-Forming Toxin Cytolysin A. JACS AU 2021; 1:1217-1230. [PMID: 34467360 PMCID: PMC8397351 DOI: 10.1021/jacsau.1c00175] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 04/20/2021] [Indexed: 05/12/2023]
Abstract
The pore-forming toxin cytolysin A (ClyA) is expressed as a large α-helical monomer that, upon interaction with membranes, undergoes a major conformational rearrangement into the protomer conformation, which then assembles into a cytolytic pore. Here, we investigate the folding kinetics of the ClyA monomer with single-molecule Förster resonance energy transfer spectroscopy in combination with microfluidic mixing, stopped-flow circular dichroism experiments, and molecular simulations. The complex folding process occurs over a broad range of time scales, from hundreds of nanoseconds to minutes. The very slow formation of the native state occurs from a rapidly formed and highly collapsed intermediate with large helical content and nonnative topology. Molecular dynamics simulations suggest pronounced non-native interactions as the origin of the slow escape from this deep trap in the free-energy surface, and a variational enhanced path-sampling approach enables a glimpse of the folding process that is supported by the experimental data.
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Affiliation(s)
- Fabian Dingfelder
- Department
of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Iuri Macocco
- Department
of Physics, Trento University, Via Sommarive 14, 38123 Povo (Trento), Italy
- SISSA, Via Bonomea 265, 34136 Trieste, Italy
| | - Stephan Benke
- Department
of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Daniel Nettels
- Department
of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Pietro Faccioli
- Department
of Physics, Trento University, Via Sommarive 14, 38123 Povo (Trento), Italy
- INFN-TIFPA, Via Sommarive 14, 38123 Povo (Trento), Italy
| | - Benjamin Schuler
- Department
of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
- Department
of Physics, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
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13
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Benke S, Holla A, Wunderlich B, Soranno A, Nettels D, Schuler B. Combining Rapid Microfluidic Mixing and Three-Color Single-Molecule FRET for Probing the Kinetics of Protein Conformational Changes. J Phys Chem B 2021; 125:6617-6628. [PMID: 34125545 DOI: 10.1021/acs.jpcb.1c02370] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/27/2023]
Abstract
Single-molecule Förster resonance energy transfer (FRET) is well suited for studying the kinetics of protein conformational changes, owing to its high sensitivity and ability to resolve individual subpopulations in heterogeneous systems. However, the most common approach employing two fluorophores can only monitor one distance at a time, and the use of three fluorophores for simultaneously monitoring multiple distances has largely been limited to equilibrium fluctuations. Here we show that three-color single-molecule FRET can be combined with rapid microfluidic mixing to investigate conformational changes in a protein from milliseconds to minutes. In combination with manual mixing, we extended the kinetics to 1 h, corresponding to a total range of 5 orders of magnitude in time. We studied the monomer-to-protomer conversion of the pore-forming toxin cytolysin A (ClyA), one of the largest protein conformational transitions known. Site-specific labeling of ClyA with three fluorophores enabled us to follow the kinetics of three intramolecular distances at the same time and revealed a previously undetected intermediate. The combination of three-color single-molecule FRET with rapid microfluidic mixing thus provides an approach for probing the mechanisms of complex biomolecular processes with high time resolution.
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Affiliation(s)
- Stephan Benke
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Andrea Holla
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Bengt Wunderlich
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Andrea Soranno
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland.,Department of Biochemistry and Molecular Biophysics, Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis, Missouri 63130, United States
| | - Daniel Nettels
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Benjamin Schuler
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland.,Department of Physics, University of Zurich, Winterthurerstrasse. 190, 8057 Zurich, Switzerland
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14
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Cubuk J, Alston JJ, Incicco JJ, Singh S, Stuchell-Brereton MD, Ward MD, Zimmerman MI, Vithani N, Griffith D, Wagoner JA, Bowman GR, Hall KB, Soranno A, Holehouse AS. The SARS-CoV-2 nucleocapsid protein is dynamic, disordered, and phase separates with RNA. Nat Commun 2021; 12:1936. [PMID: 33782395 PMCID: PMC8007728 DOI: 10.1038/s41467-021-21953-3] [Citation(s) in RCA: 278] [Impact Index Per Article: 92.7] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/30/2020] [Accepted: 02/18/2021] [Indexed: 12/15/2022] Open
Abstract
The SARS-CoV-2 nucleocapsid (N) protein is an abundant RNA-binding protein critical for viral genome packaging, yet the molecular details that underlie this process are poorly understood. Here we combine single-molecule spectroscopy with all-atom simulations to uncover the molecular details that contribute to N protein function. N protein contains three dynamic disordered regions that house putative transiently-helical binding motifs. The two folded domains interact minimally such that full-length N protein is a flexible and multivalent RNA-binding protein. N protein also undergoes liquid-liquid phase separation when mixed with RNA, and polymer theory predicts that the same multivalent interactions that drive phase separation also engender RNA compaction. We offer a simple symmetry-breaking model that provides a plausible route through which single-genome condensation preferentially occurs over phase separation, suggesting that phase separation offers a convenient macroscopic readout of a key nanoscopic interaction.
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Affiliation(s)
- Jasmine Cubuk
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA
- Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis, MO, USA
| | - Jhullian J Alston
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA
- Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis, MO, USA
| | - J Jeremías Incicco
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA
- Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis, MO, USA
| | - Sukrit Singh
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA
- Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis, MO, USA
| | - Melissa D Stuchell-Brereton
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA
- Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis, MO, USA
| | - Michael D Ward
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA
- Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis, MO, USA
| | - Maxwell I Zimmerman
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA
- Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis, MO, USA
| | - Neha Vithani
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA
- Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis, MO, USA
| | - Daniel Griffith
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA
- Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis, MO, USA
| | - Jason A Wagoner
- Laufer Center for Physical and Quantitative Biology, Stony Brook University, Stony Brook, NY, USA
| | - Gregory R Bowman
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA
- Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis, MO, USA
| | - Kathleen B Hall
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA
| | - Andrea Soranno
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA.
- Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis, MO, USA.
| | - Alex S Holehouse
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA.
- Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis, MO, USA.
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15
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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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16
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Klein F, Barrera EE, Pantano S. Assessing SIRAH's Capability to Simulate Intrinsically Disordered Proteins and Peptides. J Chem Theory Comput 2021; 17:599-604. [PMID: 33411518 DOI: 10.1021/acs.jctc.0c00948] [Citation(s) in RCA: 13] [Impact Index Per Article: 4.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/16/2023]
Abstract
The challenges posed by intrinsically disordered proteins (IDPs) to atomistic and coarse-grained (CG) simulations are boosting efforts to develop and reparametrize current force fields. An assessment of the dynamical behavior of IDPs' and unstructured peptides with the CG SIRAH force field suggests that the current version achieves a fair description of IDPs' conformational flexibility. Moreover, we found a remarkable capability to capture the effect of point mutations in loosely structured peptides.
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Affiliation(s)
- Florencia Klein
- Institut Pasteur de Montevideo, Mataojo 2020, Montevideo, CP 11400, Uruguay.,Graduate Program in Chemistry, Facultad de Química, Universidad de la República, Montevideo 11800, Uruguay
| | - Exequiel E Barrera
- Institut Pasteur de Montevideo, Mataojo 2020, Montevideo, CP 11400, Uruguay.,Instituto de Histología y Embriología (IHEM) - Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET), CC56, Universidad Nacional de Cuyo (UNCuyo), M5500 Mendoza, Argentina
| | - Sergio Pantano
- Institut Pasteur de Montevideo, Mataojo 2020, Montevideo, CP 11400, Uruguay.,Shanghai Institute for Advanced Immunochemical Studies, ShanghaiTech University, Shanghai 201210, China
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17
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Cubuk J, Alston JJ, Incicco JJ, Singh S, Stuchell-Brereton MD, Ward MD, Zimmerman MI, Vithani N, Griffith D, Wagoner JA, Bowman GR, Hall KB, Soranno A, Holehouse AS. The SARS-CoV-2 nucleocapsid protein is dynamic, disordered, and phase separates with RNA. BIORXIV : THE PREPRINT SERVER FOR BIOLOGY 2020:2020.06.17.158121. [PMID: 32587966 PMCID: PMC7310622 DOI: 10.1101/2020.06.17.158121] [Citation(s) in RCA: 30] [Impact Index Per Article: 7.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 04/11/2023]
Abstract
The SARS-CoV-2 nucleocapsid (N) protein is an abundant RNA binding protein critical for viral genome packaging, yet the molecular details that underlie this process are poorly understood. Here we combine single-molecule spectroscopy with all-atom simulations to uncover the molecular details that contribute to N protein function. N protein contains three dynamic disordered regions that house putative transiently-helical binding motifs. The two folded domains interact minimally such that full-length N protein is a flexible and multivalent RNA binding protein. N protein also undergoes liquid-liquid phase separation when mixed with RNA, and polymer theory predicts that the same multivalent interactions that drive phase separation also engender RNA compaction. We offer a simple symmetry-breaking model that provides a plausible route through which single-genome condensation preferentially occurs over phase separation, suggesting that phase separation offers a convenient macroscopic readout of a key nanoscopic interaction.
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18
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Yoon J, Ferrie JJ, Petersson EJ. Improved Modeling of Thioamide FRET Quenching by Including Conformational Restriction and Coulomb Coupling. J Phys Chem B 2020; 124:10653-10662. [PMID: 33196192 DOI: 10.1021/acs.jpcb.0c06865] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/20/2022]
Abstract
Thioamide-containing amino acids have been shown to quench a wide range of fluorophores through distinct mechanisms. Here, we quantitatively analyze the mechanism through which the thioamide functional group quenches the fluorescence of p-cyanophenylalanine (Cnf), tyrosine (Tyr), and tryptophan (Trp). By comparing PyRosetta simulations to published experiments performed on polyproline ruler peptides, we corroborate previous findings that both Cnf and Tyr quenching occurs via Förster resonance energy transfer (FRET), while Trp quenching occurs through an alternate mechanism such as Dexter transfer. Additionally, optimization of the peptide sampling scheme and comparison of thioamides attached to the peptide backbone and side chain revealed that the significant conformational restriction associated with the thioamide moiety results in a high sensitivity of the apparent FRET efficiency to underlying conformational differences. Moreover, by computing FRET efficiencies from structural models using a variety of approaches, we find that quantitative accuracy in the role of Coulomb coupling is required to explain contributions to the observed quenching efficiency from individual structures on a detailed level. Last, we demonstrate that these additional considerations improve our ability to predict thioamide quenching efficiencies observed during binding of thioamide-labeled peptides to fluorophore-labeled variants of calmodulin.
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Affiliation(s)
- Jimin Yoon
- Department of Chemistry, University of Pennsylvania, Philadelphia, Pennsylvania 19104, United States
| | - John J Ferrie
- Department of Chemistry, University of Pennsylvania, Philadelphia, Pennsylvania 19104, United States
| | - E James Petersson
- Department of Chemistry, University of Pennsylvania, Philadelphia, Pennsylvania 19104, United States
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19
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Frigini EN, Barrera EE, Pantano S, Porasso RD. Role of membrane curvature on the activation/deactivation of Carnitine Palmitoyltransferase 1A: A coarse grain molecular dynamic study. BIOCHIMICA ET BIOPHYSICA ACTA-BIOMEMBRANES 2019; 1862:183094. [PMID: 31705849 DOI: 10.1016/j.bbamem.2019.183094] [Citation(s) in RCA: 4] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Subscribe] [Scholar Register] [Received: 05/07/2019] [Revised: 09/20/2019] [Accepted: 09/23/2019] [Indexed: 10/25/2022]
Abstract
Carnitine Palmitoyltransferase 1A (CPT 1A) is an enzyme anchored to the outer mitochondrial membrane (OMM), where it regulates the passage of fatty acids into the mitochondria and intervenes in the process of β-oxidation of long-chain fatty acids. Although CPT 1A is inhibited by malonyl-CoA, its activity is also modulated by the curvature of OMM. This modulation depends on the behavior of the N-terminal domain (NTD), which can be adsorbed onto the OMM (nonactive CPT 1A) or interacting with the C-terminal domain (active CPT 1A). Aimed to provide mechanistic insights on the regulatory mechanism of CPT 1A, we studied the influence of the bilayer curvature on the NTD behavior through a series of coarse-grained (CG) molecular dynamics simulations using curved and planar membranes. Comparative analysis suggests that the main determinant for the activation/deactivation of the enzyme is the tilt angle orientation of the transmembrane (TM) domains. Planar membranes induce a wide variation on the tilt angle orientation of TM helices, while curved geometries promote small angles with the membrane normal. Our results identify the first TM domain as an important component of the membrane sensing mechanism.
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Affiliation(s)
- Ezequiel N Frigini
- Instituto de Matemáticas Aplicada San Luis (IMASL), CONICET, Facultad de Quimica, Bioquimica y Farmacia, Facultad de Ciencias Físico Matemáticas y Naturales, Universidad Nacional de San Luis, Av. Ejército de los Andes 950, San Luis 5700, Argentina
| | - Exequiel E Barrera
- Biomolecular Simulations Group, Institut Pasteur de Montevideo, Mataojo 2020, Montevideo 11400, Uruguay
| | - Sergio Pantano
- Biomolecular Simulations Group, Institut Pasteur de Montevideo, Mataojo 2020, Montevideo 11400, Uruguay
| | - Rodolfo D Porasso
- Instituto de Matemáticas Aplicada San Luis (IMASL), CONICET, Facultad de Quimica, Bioquimica y Farmacia, Facultad de Ciencias Físico Matemáticas y Naturales, Universidad Nacional de San Luis, Av. Ejército de los Andes 950, San Luis 5700, Argentina.
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20
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Chekmarev SF. How the dyes affect folding of small proteins in single-molecule FRET experiments: A simulation study. Biophys Chem 2019; 254:106243. [PMID: 31442765 DOI: 10.1016/j.bpc.2019.106243] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.2] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/16/2019] [Revised: 08/01/2019] [Accepted: 08/02/2019] [Indexed: 11/18/2022]
Abstract
A key question in the application of the single-molecule Förster resonance energy transfer (smFRET) technique to study protein folding is how the dyes affect the protein behavior. Understanding of these effects is particularly important for small proteins, for which the dyes, along with their linkers, can be comparable in size (mass) with the protein. Using a coarse-grained model, we simulated folding of BBL protein and two of its FRET constructs. The obtained results suggest that even for small proteins, such as the 45-residue BBL, the appearance of the excluded volume in the protein conformation space due to the presence of dyes does not change the overall picture of folding. At the same time, some deviations from folding of the original protein are observed, in particular, the FRET constructs fold considerably slower than the original protein because the protein collapse in the initial state of folding is slowed down due to the protein loading with relatively massive dyes.
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Affiliation(s)
- Sergei F Chekmarev
- Institute of Thermophysics, SB RAS, 630090 Novosibirsk, Russia; Department of Physics, Novosibirsk State University, 630090 Novosibirsk, Russia.
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21
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Goluguri RR, Sen S, Udgaonkar J. Microsecond sub-domain motions and the folding and misfolding of the mouse prion protein. eLife 2019; 8:e44766. [PMID: 31025940 PMCID: PMC6516828 DOI: 10.7554/elife.44766] [Citation(s) in RCA: 12] [Impact Index Per Article: 2.4] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/29/2018] [Accepted: 04/25/2019] [Indexed: 12/29/2022] Open
Abstract
Protein aggregation appears to originate from partially unfolded conformations that are sampled through stochastic fluctuations of the native protein. It has been a challenge to characterize these fluctuations, under native like conditions. Here, the conformational dynamics of the full-length (23-231) mouse prion protein were studied under native conditions, using photoinduced electron transfer coupled to fluorescence correlation spectroscopy (PET-FCS). The slowest fluctuations could be associated with the folding of the unfolded state to an intermediate state, by the use of microsecond mixing experiments. The two faster fluctuations observed by PET-FCS, could be attributed to fluctuations within the native state ensemble. The addition of salt, which is known to initiate the aggregation of the protein, resulted in an enhancement in the time scale of fluctuations in the core of the protein. The results indicate the importance of native state dynamics in initiating the aggregation of proteins.
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Affiliation(s)
- Rama Reddy Goluguri
- National Centre for Biological SciencesTata Institute of Fundamental ResearchBengaluruIndia
| | - Sreemantee Sen
- National Centre for Biological SciencesTata Institute of Fundamental ResearchBengaluruIndia
| | - Jayant Udgaonkar
- National Centre for Biological SciencesTata Institute of Fundamental ResearchBengaluruIndia
- Indian Institute of Science Education and ResearchPuneIndia
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22
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Holmstrom ED, Holla A, Zheng W, Nettels D, Best RB, Schuler B. Accurate Transfer Efficiencies, Distance Distributions, and Ensembles of Unfolded and Intrinsically Disordered Proteins From Single-Molecule FRET. Methods Enzymol 2018; 611:287-325. [PMID: 30471690 PMCID: PMC8018263 DOI: 10.1016/bs.mie.2018.09.030] [Citation(s) in RCA: 32] [Impact Index Per Article: 5.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/12/2022]
Abstract
Intrinsically disordered proteins (IDPs) sample structurally diverse ensembles. Characterizing the underlying distributions of conformations is a key step toward understanding the structural and functional properties of IDPs. One increasingly popular method for obtaining quantitative information on intramolecular distances and distributions is single-molecule Förster resonance energy transfer (FRET). Here we describe two essential elements of the quantitative analysis of single-molecule FRET data of IDPs: the sample-specific calibration of the single-molecule instrument that is required for determining accurate transfer efficiencies, and the use of state-of-the-art methods for inferring accurate distance distributions from these transfer efficiencies. First, we illustrate how to quantify the correction factors for instrument calibration with alternating donor and acceptor excitation measurements of labeled samples spanning a wide range of transfer efficiencies. Second, we show how to infer distance distributions based on suitably parameterized simple polymer models, and how to obtain conformational ensembles from Bayesian reweighting of molecular simulations or from parameter optimization in simplified coarse-grained models.
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Affiliation(s)
- Erik D Holmstrom
- Department of Biochemistry, University of Zurich, Zurich, Switzerland.
| | - Andrea Holla
- Department of Biochemistry, University of Zurich, Zurich, Switzerland
| | - Wenwei Zheng
- College of Integrative Sciences and Arts, Arizona State University, Mesa, AZ, United States.
| | - Daniel Nettels
- Department of Biochemistry, University of Zurich, Zurich, Switzerland
| | - Robert B Best
- Laboratory of Chemical Physics, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, MD, United States.
| | - Benjamin Schuler
- Department of Biochemistry, University of Zurich, Zurich, Switzerland; Department of Physics, University of Zurich, Zurich, Switzerland.
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23
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Lerner E, Cordes T, Ingargiol A, Alhadid Y, Chung S, Michalet X, Weiss S. Toward dynamic structural biology: Two decades of single-molecule Förster resonance energy transfer. Science 2018; 359:eaan1133. [PMID: 29348210 PMCID: PMC6200918 DOI: 10.1126/science.aan1133] [Citation(s) in RCA: 313] [Impact Index Per Article: 52.2] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/12/2022]
Abstract
Classical structural biology can only provide static snapshots of biomacromolecules. Single-molecule Förster resonance energy transfer (smFRET) paved the way for studying dynamics in macromolecular structures under biologically relevant conditions. Since its first implementation in 1996, smFRET experiments have confirmed previously hypothesized mechanisms and provided new insights into many fundamental biological processes, such as DNA maintenance and repair, transcription, translation, and membrane transport. We review 22 years of contributions of smFRET to our understanding of basic mechanisms in biochemistry, molecular biology, and structural biology. Additionally, building on current state-of-the-art implementations of smFRET, we highlight possible future directions for smFRET in applications such as biosensing, high-throughput screening, and molecular diagnostics.
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Affiliation(s)
- Eitan Lerner
- Department of Chemistry and Biochemistry, University of California, Los Angeles, CA 90095, USA
| | - Thorben Cordes
- Molecular Microscopy Research Group, Zernike Institute for Advanced Materials, University of Groningen, 9747 AG Groningen, Netherlands
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians-Universität München, 82152 Planegg-Martinsried, Germany
| | - Antonino Ingargiol
- Department of Chemistry and Biochemistry, University of California, Los Angeles, CA 90095, USA
| | - Yazan Alhadid
- Department of Chemistry and Biochemistry, University of California, Los Angeles, CA 90095, USA
| | - SangYoon Chung
- Department of Chemistry and Biochemistry, University of California, Los Angeles, CA 90095, USA
| | - Xavier Michalet
- Department of Chemistry and Biochemistry, University of California, Los Angeles, CA 90095, USA
| | - Shimon Weiss
- Department of Chemistry and Biochemistry, University of California, Los Angeles, CA 90095, USA
- Department of Physiology, University of California, Los Angeles, CA 90095, USA
- California NanoSystems Institute, University of California, Los Angeles, CA 90095, USA
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24
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Zosel F, Haenni D, Soranno A, Nettels D, Schuler B. Combining short- and long-range fluorescence reporters with simulations to explore the intramolecular dynamics of an intrinsically disordered protein. J Chem Phys 2017; 147:152708. [DOI: 10.1063/1.4992800] [Citation(s) in RCA: 17] [Impact Index Per Article: 2.4] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 02/03/2023] Open
Affiliation(s)
- Franziska Zosel
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Dominik Haenni
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
- Center for Microscopy and Image Analysis, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Andrea Soranno
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, Missouri 63110, USA
| | - Daniel Nettels
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Benjamin Schuler
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
- Department of Physics, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
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25
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Holmstrom ED, Nettels D, Schuler B. Conformational Plasticity of Hepatitis C Virus Core Protein Enables RNA-Induced Formation of Nucleocapsid-like Particles. J Mol Biol 2017; 430:2453-2467. [PMID: 29045818 DOI: 10.1016/j.jmb.2017.10.010] [Citation(s) in RCA: 20] [Impact Index Per Article: 2.9] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/17/2017] [Revised: 09/29/2017] [Accepted: 10/02/2017] [Indexed: 02/08/2023]
Abstract
Many of the unanswered questions associated with hepatitis C virus assembly are related to the core protein (HCVcp), which forms an oligomeric nucleocapsid encompassing the viral genome. The structural properties of HCVcp have been difficult to quantify, at least in part because it is an intrinsically disordered protein. We have used single-molecule Förster Resonance Energy Transfer techniques to study the conformational dimensions and dynamics of the HCVcp nucleocapsid domain (HCVncd) at various stages during the RNA-induced formation of nucleocapsid-like particles. Our results indicate that HCVncd is a typical intrinsically disordered protein. When it forms small ribonucleoprotein complexes with various RNA hairpins from the 3' end of the HCV genome, it compacts but remains intrinsically disordered and conformationally dynamic. Above a critical RNA concentration, these ribonucleoprotein complexes rapidly and cooperatively assemble into large nucleocapsid-like particles, wherein the individual HCVncd subunits become substantially more extended.
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Affiliation(s)
- Erik D Holmstrom
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland.
| | - Daniel Nettels
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland.
| | - Benjamin Schuler
- Department of Biochemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland; Department of Physics, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland.
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26
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Rahamim G, Amir D, Haas E. Simultaneous Determination of Two Subdomain Folding Rates Using the "Transfer-Quench" Method. Biophys J 2017; 112:1786-1796. [PMID: 28494950 DOI: 10.1016/j.bpj.2017.01.037] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/10/2016] [Revised: 12/21/2016] [Accepted: 01/06/2017] [Indexed: 11/29/2022] Open
Abstract
The investigation of the mechanism of protein folding is complicated by the context dependence of the rates of intramolecular contact formation. Methods based on site-specific labeling and ultrafast spectroscopic detection of fluorescence signals were developed for monitoring the rates of individual subdomain folding transitions in situ, in the context of the whole molecule. However, each site-specific labeling modification might affect rates of folding of near-neighbor structural elements, and thus limit the ability to resolve fine differences in rates of folding of these elements. Therefore, it is highly desirable to be able to study the rates of folding of two or more neighboring subdomain structures using a single mutant to facilitate resolution of the order and interdependence of such steps. Here, we report the development of the "Transfer-Quench" method for measuring the rate of formation of two structural elements using a single triple-labeled mutant. This method is based on Förster resonance energy transfer combined with fluorescence quenching. We placed the donor and acceptor at the loop ends, and a quencher at an α-helical element involved in the node forming the loop. The folding of the triple-labeled mutant is monitored by the acceptor emission. The formation of nonlocal contact (loop closure) increases the time-dependent acceptor emission, while the closure of the labeled helix turn reduces this emission. The method was applied in a study of the folding mechanism of the common model protein, the B domain of staphylococcal protein A. Only natural amino acids were used as probes, and thus possible structural perturbations were minimized. Tyr and Trp residues served as donor and acceptor at the ends of a long loop between helices I and II, and a Cys residue as a quencher for the acceptor. We found that the closure of the loop (segment 14-33) occurs with the same rate constant as the nucleation of helix HII (segment 33-29), in line with the nucleation-condensation model.
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Affiliation(s)
- Gil Rahamim
- The Goodman Faculty of Life Sciences Bar Ilan University, Ramat Gan, Israel
| | - Dan Amir
- The Goodman Faculty of Life Sciences Bar Ilan University, Ramat Gan, Israel
| | - Elisha Haas
- The Goodman Faculty of Life Sciences Bar Ilan University, Ramat Gan, Israel.
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27
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Jazi AA, Ploetz E, Arizki M, Dhandayuthapani B, Waclawska I, Krämer R, Ziegler C, Cordes T. Caging and Photoactivation in Single-Molecule Förster Resonance Energy Transfer Experiments. Biochemistry 2017; 56:2031-2041. [PMID: 28362086 PMCID: PMC5390306 DOI: 10.1021/acs.biochem.6b00916] [Citation(s) in RCA: 12] [Impact Index Per Article: 1.7] [Reference Citation Analysis] [Abstract] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/28/2022]
Abstract
![]()
Caged
organic fluorophores are established tools for localization-based
super-resolution imaging. Their use relies on reversible deactivation
of standard organic fluorophores by chemical reduction or commercially
available caged dyes with ON switching of the fluorescent signal by
ultraviolet (UV) light. Here, we establish caging of cyanine fluorophores
and caged rhodamine dyes, i.e., chemical deactivation of fluorescence,
for single-molecule Förster resonance energy transfer (smFRET)
experiments with freely diffusing molecules. They allow temporal separation
and sorting of multiple intramolecular donor–acceptor pairs
during solution-based smFRET. We use this “caged FRET”
methodology for the study of complex biochemical species such as multisubunit
proteins or nucleic acids containing more than two fluorescent labels.
Proof-of-principle experiments and a characterization of the uncaging
process in the confocal volume are presented. These reveal that chemical
caging and UV reactivation allow temporal uncoupling of convoluted
fluorescence signals from, e.g., multiple spectrally similar donor
or acceptor molecules on nucleic acids. We also use caging without
UV reactivation to remove unwanted overlabeled species in experiments
with the homotrimeric membrane transporter BetP. We finally outline
further possible applications of the caged FRET methodology, such
as the study of weak biochemical interactions, which are otherwise
impossible with diffusion-based smFRET techniques because of the required
low concentrations of fluorescently labeled biomolecules.
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Affiliation(s)
- Atieh Aminian Jazi
- Molecular Microscopy Research Group, Zernike Institute for Advanced Materials, University of Groningen , Nijenborgh 4, 9747 AG Groningen, The Netherlands.,Institute of Biophysics and Biophysical Chemistry, Universität Regensburg , 95053 Regensburg, Germany
| | - Evelyn Ploetz
- Molecular Microscopy Research Group, Zernike Institute for Advanced Materials, University of Groningen , Nijenborgh 4, 9747 AG Groningen, The Netherlands
| | - Muhamad Arizki
- Molecular Microscopy Research Group, Zernike Institute for Advanced Materials, University of Groningen , Nijenborgh 4, 9747 AG Groningen, The Netherlands
| | | | - Izabela Waclawska
- Institute of Biophysics and Biophysical Chemistry, Universität Regensburg , 95053 Regensburg, Germany
| | - Reinhard Krämer
- Institute for Biochemistry, Universität Köln , 50674 Köln, Germany
| | - Christine Ziegler
- Institute of Biophysics and Biophysical Chemistry, Universität Regensburg , 95053 Regensburg, Germany
| | - Thorben Cordes
- Molecular Microscopy Research Group, Zernike Institute for Advanced Materials, University of Groningen , Nijenborgh 4, 9747 AG Groningen, The Netherlands
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28
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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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29
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Förster resonance energy transfer and protein-induced fluorescence enhancement as synergetic multi-scale molecular rulers. Sci Rep 2016; 6:33257. [PMID: 27641327 PMCID: PMC5027553 DOI: 10.1038/srep33257] [Citation(s) in RCA: 59] [Impact Index Per Article: 7.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/15/2016] [Accepted: 08/24/2016] [Indexed: 01/24/2023] Open
Abstract
Advanced microscopy methods allow obtaining information on (dynamic) conformational changes in biomolecules via measuring a single molecular distance in the structure. It is, however, extremely challenging to capture the full depth of a three-dimensional biochemical state, binding-related structural changes or conformational cross-talk in multi-protein complexes using one-dimensional assays. In this paper we address this fundamental problem by extending the standard molecular ruler based on Förster resonance energy transfer (FRET) into a two-dimensional assay via its combination with protein-induced fluorescence enhancement (PIFE). We show that donor brightness (via PIFE) and energy transfer efficiency (via FRET) can simultaneously report on e.g., the conformational state of double stranded DNA (dsDNA) following its interaction with unlabelled proteins (BamHI, EcoRV, and T7 DNA polymerase gp5/trx). The PIFE-FRET assay uses established labelling protocols and single molecule fluorescence detection schemes (alternating-laser excitation, ALEX). Besides quantitative studies of PIFE and FRET ruler characteristics, we outline possible applications of ALEX-based PIFE-FRET for single-molecule studies with diffusing and immobilized molecules. Finally, we study transcription initiation and scrunching of E. coli RNA-polymerase with PIFE-FRET and provide direct evidence for the physical presence and vicinity of the polymerase that causes structural changes and scrunching of the transcriptional DNA bubble.
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30
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Hamano N, Murata M, Kawano T, Piao JS, Narahara S, Nakata R, Akahoshi T, Ikeda T, Hashizume M. Förster Resonance Energy Transfer-Based Self-Assembled Nanoprobe for Rapid and Sensitive Detection of Postoperative Pancreatic Fistula. ACS APPLIED MATERIALS & INTERFACES 2016; 8:5114-5123. [PMID: 26845508 DOI: 10.1021/acsami.5b11902] [Citation(s) in RCA: 7] [Impact Index Per Article: 0.9] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/05/2023]
Abstract
Postoperative pancreatic fistula (POPF) is the most serious and challenging complication following gastroenterological surgery. Activated pancreatic juice leaking from the organ remnant contains proteases that attack the surrounding tissue, potentially leading to severe inflammation, tissue necrosis, and fistula formation. However, it is difficult to observe pancreatic leakage during surgery and to evaluate the protease activity of leaked fluid at the patient's bedside. This report describes a protein nanocage-based protease ratiometric sensor comprising a pancreatic protease-sensitive small heat-shock protein (HSP) 16.5, which is a naturally occurring protein in Methanococcus jannaschii that forms a spherical structure by self-assembly of 24 subunits, and a chemically conjugated donor-acceptor dye pair for Förster resonance energy transfer (FRET). The HSP-FRET probe was constructed by subunit exchange of each dye-labeled engineered HSP, resulting in a spherical nanocage of approximately 10 nm in diameter, which exhibited very high stability against degradation in blood plasma and no remarkable toxicity in mice. The efficiency of FRET was found to depend on both the dye orientation and the acceptor/donor ratio. Pancreatic proteases, including trypsin, α-chymotrypsin, and elastase, were quantitatively analyzed by fluorescence recovery with high specificity using the HSP-FRET nanoprobe. Furthermore, the HSP-FRET nanoprobe was sufficiently sensitive to detect POPF in the pancreatic juice of patients using only the naked eye within 10 min. Thus, this novel nanoprobe is proposed as an effective and convenient tool for the detection of POPF and the visualization of activated pancreatic juice during gastroenterological surgery.
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Affiliation(s)
- Nobuhito Hamano
- Center for Advanced Medical Innovation, ‡Department of Advanced Medical Initiatives, Faculty of Medical Sciences, and §Innovation Center for Medical Redox Navigation, Kyushu University , 3-1-1 Maidashi, Higashi-ku, Fukuoka 812-8582, Japan
| | - Masaharu Murata
- Center for Advanced Medical Innovation, ‡Department of Advanced Medical Initiatives, Faculty of Medical Sciences, and §Innovation Center for Medical Redox Navigation, Kyushu University , 3-1-1 Maidashi, Higashi-ku, Fukuoka 812-8582, Japan
| | - Takahito Kawano
- Center for Advanced Medical Innovation, ‡Department of Advanced Medical Initiatives, Faculty of Medical Sciences, and §Innovation Center for Medical Redox Navigation, Kyushu University , 3-1-1 Maidashi, Higashi-ku, Fukuoka 812-8582, Japan
| | - Jing Shu Piao
- Center for Advanced Medical Innovation, ‡Department of Advanced Medical Initiatives, Faculty of Medical Sciences, and §Innovation Center for Medical Redox Navigation, Kyushu University , 3-1-1 Maidashi, Higashi-ku, Fukuoka 812-8582, Japan
| | - Sayoko Narahara
- Center for Advanced Medical Innovation, ‡Department of Advanced Medical Initiatives, Faculty of Medical Sciences, and §Innovation Center for Medical Redox Navigation, Kyushu University , 3-1-1 Maidashi, Higashi-ku, Fukuoka 812-8582, Japan
| | - Ryosuke Nakata
- Center for Advanced Medical Innovation, ‡Department of Advanced Medical Initiatives, Faculty of Medical Sciences, and §Innovation Center for Medical Redox Navigation, Kyushu University , 3-1-1 Maidashi, Higashi-ku, Fukuoka 812-8582, Japan
| | - Tomohiko Akahoshi
- Center for Advanced Medical Innovation, ‡Department of Advanced Medical Initiatives, Faculty of Medical Sciences, and §Innovation Center for Medical Redox Navigation, Kyushu University , 3-1-1 Maidashi, Higashi-ku, Fukuoka 812-8582, Japan
| | - Tetsuo Ikeda
- Center for Advanced Medical Innovation, ‡Department of Advanced Medical Initiatives, Faculty of Medical Sciences, and §Innovation Center for Medical Redox Navigation, Kyushu University , 3-1-1 Maidashi, Higashi-ku, Fukuoka 812-8582, Japan
| | - Makoto Hashizume
- Center for Advanced Medical Innovation, ‡Department of Advanced Medical Initiatives, Faculty of Medical Sciences, and §Innovation Center for Medical Redox Navigation, Kyushu University , 3-1-1 Maidashi, Higashi-ku, Fukuoka 812-8582, Japan
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31
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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: 50] [Impact Index Per Article: 6.3] [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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32
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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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33
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Solntsev KM, Popov AV, Solovyeva VA, Al-Ainain SA, Il’ichev YV, Hernandez R, Kuzmin MG. Kinetics of intra- and intermolecular excited-state proton transfer ofω-(2-hydroxynaphthyl-1)-decanoic acid in homogeneous and micellar solutions. Methods Appl Fluoresc 2015. [DOI: 10.1088/2050-6120/4/1/014001] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.1] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/12/2022]
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34
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Kyrychenko A. Using fluorescence for studies of biological membranes: a review. Methods Appl Fluoresc 2015; 3:042003. [DOI: 10.1088/2050-6120/3/4/042003] [Citation(s) in RCA: 28] [Impact Index Per Article: 3.1] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/12/2022]
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35
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Kempe D, Schöne A, Fitter J, Gabba M. Accurate Fluorescence Quantum Yield Determination by Fluorescence Correlation Spectroscopy. J Phys Chem B 2015; 119:4668-72. [DOI: 10.1021/acs.jpcb.5b02170] [Citation(s) in RCA: 13] [Impact Index Per Article: 1.4] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/13/2022]
Affiliation(s)
- Daryan Kempe
- AG
Biophysik, I. Physikalisches Institut (IA), RWTH Aachen University, 52062 Aachen, Germany
| | - Antonie Schöne
- Molecular
Biophysics, Institute of Complex Systems (ICS-5), Forschungszentrum Jülich, 52428 Jülich, Germany
| | - Jörg Fitter
- AG
Biophysik, I. Physikalisches Institut (IA), RWTH Aachen University, 52062 Aachen, Germany
- Molecular
Biophysics, Institute of Complex Systems (ICS-5), Forschungszentrum Jülich, 52428 Jülich, Germany
| | - Matteo Gabba
- Molecular
Biophysics, Institute of Complex Systems (ICS-5), Forschungszentrum Jülich, 52428 Jülich, Germany
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36
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Darré L, Machado MR, Brandner AF, González HC, Ferreira S, Pantano S. SIRAH: A Structurally Unbiased Coarse-Grained Force Field for Proteins with Aqueous Solvation and Long-Range Electrostatics. J Chem Theory Comput 2015; 11:723-39. [DOI: 10.1021/ct5007746] [Citation(s) in RCA: 99] [Impact Index Per Article: 11.0] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/09/2023]
Affiliation(s)
- Leonardo Darré
- Institut Pasteur de Montevideo, Montevideo, Uruguay
- Department
of Chemistry, King’s College, London, United Kingdom
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37
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Nettels D, Haenni D, Maillot S, Gueye M, Barth A, Hirschfeld V, Hübner CG, Léonard J, Schuler B. Excited-state annihilation reduces power dependence of single-molecule FRET experiments. Phys Chem Chem Phys 2015; 17:32304-15. [DOI: 10.1039/c5cp05321h] [Citation(s) in RCA: 18] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/12/2023]
Abstract
Singlet–singlet annihilation between FRET dyes is evident in nanosecond fluorescence cross-correlation measurements.
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Affiliation(s)
- Daniel Nettels
- Department of Biochemistry
- University of Zurich
- 8057 Zurich
- Switzerland
| | - Dominik Haenni
- Department of Biochemistry
- University of Zurich
- 8057 Zurich
- Switzerland
| | - Sacha Maillot
- Institut de Physique et Chimie des Matériaux de Strasbourg & Labex NIE
- Université de Strasbourg
- 67034 Strasbourg Cedex 2
- France
| | - Moussa Gueye
- Institut de Physique et Chimie des Matériaux de Strasbourg & Labex NIE
- Université de Strasbourg
- 67034 Strasbourg Cedex 2
- France
| | - Anders Barth
- Institute of Physics
- University of Lübeck
- 23562 Lübeck
- Germany
| | | | | | - Jérémie Léonard
- Institut de Physique et Chimie des Matériaux de Strasbourg & Labex NIE
- Université de Strasbourg
- 67034 Strasbourg Cedex 2
- France
| | - Benjamin Schuler
- Department of Biochemistry
- University of Zurich
- 8057 Zurich
- Switzerland
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Single-molecule spectroscopy reveals chaperone-mediated expansion of substrate protein. Proc Natl Acad Sci U S A 2014; 111:13355-60. [PMID: 25165400 DOI: 10.1073/pnas.1407086111] [Citation(s) in RCA: 92] [Impact Index Per Article: 9.2] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/18/2022] Open
Abstract
Molecular chaperones are an essential part of the machinery that avoids protein aggregation and misfolding in vivo. However, understanding the molecular basis of how chaperones prevent such undesirable interactions requires the conformational changes within substrate proteins to be probed during chaperone action. Here we use single-molecule fluorescence spectroscopy to investigate how the DnaJ-DnaK chaperone system alters the conformational distribution of the denatured substrate protein rhodanese. We find that in a first step the ATP-independent binding of DnaJ to denatured rhodanese results in a compact denatured ensemble of the substrate protein. The following ATP-dependent binding of multiple DnaK molecules, however, leads to a surprisingly large expansion of denatured rhodanese. Molecular simulations indicate that hard-core repulsion between the multiple DnaK molecules provides the underlying mechanism for disrupting even strong interactions within the substrate protein and preparing it for processing by downstream chaperone systems.
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Brucale M, Schuler B, Samorì B. Single-molecule studies of intrinsically disordered proteins. Chem Rev 2014; 114:3281-317. [PMID: 24432838 DOI: 10.1021/cr400297g] [Citation(s) in RCA: 112] [Impact Index Per Article: 11.2] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/22/2022]
Affiliation(s)
- Marco Brucale
- Institute for the Study of Nanostructured Materials (ISMN), Italian National Council of Research (CNR) , Area della Ricerca Roma1, Via Salaria km 29.3 00015 Monterotondo (Rome), Italy
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