1
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Frost D, Cook K, Sanabria H. Time-heterogeneity of the Förster Radius from Dipole Orientational Dynamics Explains Observed Dynamic Shift. ARXIV 2024:arXiv:2404.09883v1. [PMID: 38699162 PMCID: PMC11065046] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Grants] [Subscribe] [Scholar Register] [Indexed: 05/05/2024]
Abstract
Förster resonance energy transfer (FRET) is a quantum mechanical phenomenon involving the non-radiative transfer of energy between coupled electric dipoles. Due to the strong dependence of FRET on the distance between the dipoles, it is frequently used as a "molecular ruler" in biology, chemistry, and physics. This is done by placing dipolar molecules called dyes on molecules of interest. In time-resolved confocal single-molecule FRET (smFRET) experiments, the joint distribution of the FRET efficiency and the donor fluorescence lifetime can reveal underlying molecular conformational dynamics via deviation from their theoretical Förster relationship. This deviation is referred to as a dynamic shift. Quantifying the dynamic shift caused by the motion of the fluorescent dyes is essential to decoupling the dynamics of the studied molecules and the dyes. We develop novel Langevin models for the dye linker dynamics, including rotational dynamics, based on first physics principles and proper dye linker chemistry to match accessible volumes predicted by molecular dynamics simulations. By simulating the dyes' stochastic translational and rotational dynamics, we show that the observed dynamic shift can largely be attributed to the mutual orientational dynamics of the electric dipole moments associated with the dyes, not their accessible volume.
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Affiliation(s)
- David Frost
- School of Mathematical and Statistical Sciences, Clemson University
| | - Keisha Cook
- School of Mathematical and Statistical Sciences, Clemson University
| | - Hugo Sanabria
- Department of Physics and Astronomy, Clemson University
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2
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Kolimi N, Ballard J, Peulen T, Goutam R, Duffy FX, Ramírez-Sarmiento CA, Babul J, Medina E, Sanabria H. DNA controls the dimerization of the human FoxP1 forkhead domain. CELL REPORTS. PHYSICAL SCIENCE 2024; 5:101854. [PMID: 38585429 PMCID: PMC10997372 DOI: 10.1016/j.xcrp.2024.101854] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 04/09/2024]
Abstract
Transcription factors (TFs) regulate gene expression by binding to specific DNA sequences and gating access to genes. Even when the binding of TFs and their cofactors to DNA is reversible, indicating a reversible control of gene expression, there is little knowledge about the molecular effect DNA has on TFs. Using single-molecule multiparameter fluorescence spectroscopy, molecular dynamics simulations, and biochemical assays, we find that the monomeric form of the forkhead (FKH) domain of the human FoxP1 behaves as a disordered protein and increases its folded population when it dimerizes. Notably, DNA binding promotes a disordered FKH dimer bound to DNA, negatively controlling the stability of the dimeric FoxP1:DNA complex. The DNA-mediated reversible regulation on FKH dimers suggests that FoxP1-dependent gene suppression is unstable, and it must require the presence of other dimerization domains or cofactors to revert the negative impact exerted by the DNA.
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Affiliation(s)
- Narendar Kolimi
- Department of Physics and Astronomy, Clemson University, Clemson, SC 29634, USA
| | - Jake Ballard
- Department of Physics and Astronomy, Clemson University, Clemson, SC 29634, USA
| | - Thomas Peulen
- Rudolf-Virchow-Zentrum – Center for Integrative and Translational Bioimaging, Haus D15, Josef-Schneider-Straße 2, 97080 Würzburg Germany
| | - Rajen Goutam
- Department of Physics and Astronomy, Clemson University, Clemson, SC 29634, USA
| | - Francis X. Duffy
- Department of Physics and Astronomy, Clemson University, Clemson, SC 29634, USA
| | - César A. Ramírez-Sarmiento
- Institute for Biological and Medical Engineering, Schools of Engineering, Medicine and Biological Sciences, Pontificia Universidad Católica de Chile, Santiago 7820436, Chile
- ANID – Millennium Science Initiative Program – Millennium Institute for Integrative Biology (iBio), Santiago 8331150, Chile
| | - Jorge Babul
- Departamento de Biología, Facultad de Ciencias, Universidad de Chile, Las Palmeras 3425, Casilla 653, Santiago 7800003, Chile
| | - Exequiel Medina
- Department of Physics and Astronomy, Clemson University, Clemson, SC 29634, USA
- Departamento de Biología, Facultad de Ciencias, Universidad de Chile, Las Palmeras 3425, Casilla 653, Santiago 7800003, Chile
| | - Hugo Sanabria
- Department of Physics and Astronomy, Clemson University, Clemson, SC 29634, USA
- Lead contact
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3
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Klughammer N, Barth A, Dekker M, Fragasso A, Onck PR, Dekker C. Diameter dependence of transport through nuclear pore complex mimics studied using optical nanopores. eLife 2024; 12:RP87174. [PMID: 38376900 PMCID: PMC10942607 DOI: 10.7554/elife.87174] [Citation(s) in RCA: 1] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 02/21/2024] Open
Abstract
The nuclear pore complex (NPC) regulates the selective transport of large biomolecules through the nuclear envelope. As a model system for nuclear transport, we construct NPC mimics by functionalizing the pore walls of freestanding palladium zero-mode waveguides with the FG-nucleoporin Nsp1. This approach enables the measurement of single-molecule translocations through individual pores using optical detection. We probe the selectivity of Nsp1-coated pores by quantitatively comparing the translocation rates of the nuclear transport receptor Kap95 to the inert probe BSA over a wide range of pore sizes from 35 nm to 160 nm. Pores below 55 ± 5 nm show significant selectivity that gradually decreases for larger pores. This finding is corroborated by coarse-grained molecular dynamics simulations of the Nsp1 mesh within the pore, which suggest that leakage of BSA occurs by diffusion through transient openings within the dynamic mesh. Furthermore, we experimentally observe a modulation of the BSA permeation when varying the concentration of Kap95. The results demonstrate the potential of single-molecule fluorescence measurements on biomimetic NPCs to elucidate the principles of nuclear transport.
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Affiliation(s)
- Nils Klughammer
- Department of Bionanoscience, Kavli Institute of Nanoscience, Delft University of TechnologyDelftNetherlands
| | - Anders Barth
- Department of Bionanoscience, Kavli Institute of Nanoscience, Delft University of TechnologyDelftNetherlands
| | - Maurice Dekker
- Zernike Institute for Advanced Materials, University of GroningenGroningenNetherlands
| | - Alessio Fragasso
- Department of Bionanoscience, Kavli Institute of Nanoscience, Delft University of TechnologyDelftNetherlands
| | - Patrick R Onck
- Zernike Institute for Advanced Materials, University of GroningenGroningenNetherlands
| | - Cees Dekker
- Department of Bionanoscience, Kavli Institute of Nanoscience, Delft University of TechnologyDelftNetherlands
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4
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Bogetti X, Saxena S. Integrating Electron Paramagnetic Resonance Spectroscopy and Computational Modeling to Measure Protein Structure and Dynamics. Chempluschem 2024; 89:e202300506. [PMID: 37801003 DOI: 10.1002/cplu.202300506] [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: 09/07/2023] [Revised: 10/05/2023] [Accepted: 10/06/2023] [Indexed: 10/07/2023]
Abstract
Electron paramagnetic resonance (EPR) has become a powerful probe of conformational heterogeneity and dynamics of biomolecules. In this Review, we discuss different computational modeling techniques that enrich the interpretation of EPR measurements of dynamics or distance restraints. A variety of spin labels are surveyed to provide a background for the discussion of modeling tools. Molecular dynamics (MD) simulations of models containing spin labels provide dynamical properties of biomolecules and their labels. These simulations can be used to predict EPR spectra, sample stable conformations and sample rotameric preferences of label sidechains. For molecular motions longer than milliseconds, enhanced sampling strategies and de novo prediction software incorporating or validated by EPR measurements are able to efficiently refine or predict protein conformations, respectively. To sample large-amplitude conformational transition, a coarse-grained or an atomistic weighted ensemble (WE) strategy can be guided with EPR insights. Looking forward, we anticipate an integrative strategy for efficient sampling of alternate conformations by de novo predictions, followed by validations by systematic EPR measurements and MD simulations. Continuous pathways between alternate states can be further sampled by WE-MD including all intermediate states.
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Affiliation(s)
- Xiaowei Bogetti
- Department of Chemistry, University of Pittsburgh, 219 Parkman Avenue, Pittsburgh, PA, 15260, USA
| | - Sunil Saxena
- Department of Chemistry, University of Pittsburgh, 219 Parkman Avenue, Pittsburgh, PA, 15260, USA
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5
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Post M, Wolf S, Stock G. Investigation of Rare Protein Conformational Transitions via Dissipation-Corrected Targeted Molecular Dynamics. J Chem Theory Comput 2023; 19:8978-8986. [PMID: 38011829 DOI: 10.1021/acs.jctc.3c01017] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/29/2023]
Abstract
To sample rare events, dissipation-corrected targeted molecular dynamics (dcTMD) applies a constant velocity constraint along a one-dimensional reaction coordinate s, which drives an atomistic system from an initial state into a target state. Employing a cumulant approximation of Jarzynski's identity, the free energy ΔG(s) is calculated from the mean external work and dissipated work of the process. By calculating the friction coefficient Γ(s) from the dissipated work, in a second step, the equilibrium dynamics of the process can be studied by propagating a Langevin equation. While so far dcTMD has been mostly applied to study the unbinding of protein-ligand complexes, here its applicability to rare conformational transitions within a protein and the prediction of their kinetics are investigated. As this typically requires the introduction of multiple collective variables {xj} = x, a theoretical framework is outlined to calculate the associated free energy ΔG(x) and friction Γ(x) from dcTMD simulations along coordinate s. Adopting the α-β transition of alanine dipeptide as well as the open-closed transition of T4 lysozyme as representative examples, the virtues and shortcomings of dcTMD to predict protein conformational transitions and the related kinetics are studied.
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Affiliation(s)
- Matthias Post
- Biomolecular Dynamics, Institute of Physics, University of Freiburg, Freiburg 79104, Germany
| | - Steffen Wolf
- Biomolecular Dynamics, Institute of Physics, University of Freiburg, Freiburg 79104, Germany
| | - Gerhard Stock
- Biomolecular Dynamics, Institute of Physics, University of Freiburg, Freiburg 79104, Germany
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6
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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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7
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Wolff AM, Nango E, Young ID, Brewster AS, Kubo M, Nomura T, Sugahara M, Owada S, Barad BA, Ito K, Bhowmick A, Carbajo S, Hino T, Holton JM, Im D, O'Riordan LJ, Tanaka T, Tanaka R, Sierra RG, Yumoto F, Tono K, Iwata S, Sauter NK, Fraser JS, Thompson MC. Mapping protein dynamics at high spatial resolution with temperature-jump X-ray crystallography. Nat Chem 2023; 15:1549-1558. [PMID: 37723259 PMCID: PMC10624634 DOI: 10.1038/s41557-023-01329-4] [Citation(s) in RCA: 1] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/16/2022] [Accepted: 08/17/2023] [Indexed: 09/20/2023]
Abstract
Understanding and controlling protein motion at atomic resolution is a hallmark challenge for structural biologists and protein engineers because conformational dynamics are essential for complex functions such as enzyme catalysis and allosteric regulation. Time-resolved crystallography offers a window into protein motions, yet without a universal perturbation to initiate conformational changes the method has been limited in scope. Here we couple a solvent-based temperature jump with time-resolved crystallography to visualize structural motions in lysozyme, a dynamic enzyme. We observed widespread atomic vibrations on the nanosecond timescale, which evolve on the submillisecond timescale into localized structural fluctuations that are coupled to the active site. An orthogonal perturbation to the enzyme, inhibitor binding, altered these dynamics by blocking key motions that allow energy to dissipate from vibrations into functional movements linked to the catalytic cycle. Because temperature jump is a universal method for perturbing molecular motion, the method demonstrated here is broadly applicable for studying protein dynamics.
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Affiliation(s)
- Alexander M Wolff
- Department of Chemistry and Biochemistry, University of California, Merced, Merced, CA, USA
| | - Eriko Nango
- RIKEN SPring-8 Center, Sayo-gun, Japan.
- Institute of Multidisciplinary Research for Advanced Materials, Tohoku University, Aoba-ku, Japan.
| | - Iris D Young
- Department of Bioengineering and Therapeutic Sciences, University of California, San Francisco, San Francisco, CA, USA
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, USA
| | - Aaron S Brewster
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, USA
| | - Minoru Kubo
- RIKEN SPring-8 Center, Sayo-gun, Japan
- Department of Life Science, Graduate School of Science, University of Hyogo, Hyogo, Japan
| | - Takashi Nomura
- RIKEN SPring-8 Center, Sayo-gun, Japan
- Department of Life Science, Graduate School of Science, University of Hyogo, Hyogo, Japan
| | | | | | - Benjamin A Barad
- Department of Bioengineering and Therapeutic Sciences, University of California, San Francisco, San Francisco, CA, USA
- Department of Integrative Structural and Computational Biology, Scripps Research, San Diego, CA, USA
| | - Kazutaka Ito
- Laboratory for Drug Discovery, Pharmaceuticals Research Center, Asahi Kasei Pharma Corporation, Izunokuni-shi, Japan
| | - Asmit Bhowmick
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, USA
| | - Sergio Carbajo
- SLAC National Accelerator Laboratory, Linac Coherent Light Source, Menlo Park, CA, USA
- Department of Electrical and Computer Engineering, University of California, Los Angeles, Los Angeles, CA, USA
| | - Tomoya Hino
- Department of Chemistry and Biotechnology, Graduate School of Engineering, Tottori University, Tottori, Japan
- Center for Research on Green Sustainable Chemistry, Tottori University, Tottori, Japan
| | - James M Holton
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, USA
- Department of Biochemistry and Biophysics, University of California San Francisco, San Francisco, CA, USA
- Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Laboratory, Menlo Park, CA, USA
| | - Dohyun Im
- Department of Cell Biology, Graduate School of Medicine, Kyoto University, Yoshidakonoe-cho, Sakyo-ku, Japan
| | - Lee J O'Riordan
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, USA
| | - Tomoyuki Tanaka
- RIKEN SPring-8 Center, Sayo-gun, Japan
- Department of Cell Biology, Graduate School of Medicine, Kyoto University, Yoshidakonoe-cho, Sakyo-ku, Japan
| | - Rie Tanaka
- RIKEN SPring-8 Center, Sayo-gun, Japan
- Department of Cell Biology, Graduate School of Medicine, Kyoto University, Yoshidakonoe-cho, Sakyo-ku, Japan
| | - Raymond G Sierra
- SLAC National Accelerator Laboratory, Linac Coherent Light Source, Menlo Park, CA, USA
| | - Fumiaki Yumoto
- Structural Biology Research Center, Institute of Materials Structure Science, KEK/High Energy Accelerator Research Organization, Tsukuba, Japan
- Ginward Japan K.K., Tokyo, Japan
| | - Kensuke Tono
- RIKEN SPring-8 Center, Sayo-gun, Japan
- Japan Synchrotron Radiation Research Institute, Hyogo, Japan
| | - So Iwata
- RIKEN SPring-8 Center, Sayo-gun, Japan
- Department of Cell Biology, Graduate School of Medicine, Kyoto University, Yoshidakonoe-cho, Sakyo-ku, Japan
| | - Nicholas K Sauter
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, USA
| | - James S Fraser
- Department of Bioengineering and Therapeutic Sciences, University of California, San Francisco, San Francisco, CA, USA
| | - Michael C Thompson
- Department of Chemistry and Biochemistry, University of California, Merced, Merced, CA, USA.
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8
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Hartmann A, Sreenivasa K, Schenkel M, Chamachi N, Schake P, Krainer G, Schlierf M. An automated single-molecule FRET platform for high-content, multiwell plate screening of biomolecular conformations and dynamics. Nat Commun 2023; 14:6511. [PMID: 37845199 PMCID: PMC10579363 DOI: 10.1038/s41467-023-42232-3] [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: 03/08/2023] [Accepted: 10/03/2023] [Indexed: 10/18/2023] Open
Abstract
Single-molecule FRET (smFRET) has become a versatile tool for probing the structure and functional dynamics of biomolecular systems, and is extensively used to address questions ranging from biomolecular folding to drug discovery. Confocal smFRET measurements are amongst the widely used smFRET assays and are typically performed in a single-well format. Thus, sampling of many experimental parameters is laborious and time consuming. To address this challenge, we extend here the capabilities of confocal smFRET beyond single-well measurements by integrating a multiwell plate functionality to allow for continuous and automated smFRET measurements. We demonstrate the broad applicability of the multiwell plate assay towards DNA hairpin dynamics, protein folding, competitive and cooperative protein-DNA interactions, and drug-discovery, revealing insights that would be very difficult to achieve with conventional single-well format measurements. For the adaptation into existing instrumentations, we provide a detailed guide and open-source acquisition and analysis software.
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Affiliation(s)
- Andreas Hartmann
- B CUBE Center for Molecular Bioengineering, TU Dresden, Tatzberg 41, 01307, Dresden, Germany.
| | - Koushik Sreenivasa
- B CUBE Center for Molecular Bioengineering, TU Dresden, Tatzberg 41, 01307, Dresden, Germany
- Department of Bionanoscience, Delft University of Technology, 2629HZ, Delft, Netherlands
| | - Mathias Schenkel
- B CUBE Center for Molecular Bioengineering, TU Dresden, Tatzberg 41, 01307, Dresden, Germany
| | - Neharika Chamachi
- B CUBE Center for Molecular Bioengineering, TU Dresden, Tatzberg 41, 01307, Dresden, Germany
| | - Philipp Schake
- B CUBE Center for Molecular Bioengineering, TU Dresden, Tatzberg 41, 01307, Dresden, Germany
| | - Georg Krainer
- B CUBE Center for Molecular Bioengineering, TU Dresden, Tatzberg 41, 01307, Dresden, Germany
- Centre for Misfolding Diseases, Yusuf Hamied Department of Chemistry, University of Cambridge, Lensfield Road, CB2 1EW, Cambridge, UK
- Institute of Molecular Biosciences, University of Graz, Humboldtstrasse 50/III, 8010, Graz, Austria
| | - Michael Schlierf
- B CUBE Center for Molecular Bioengineering, TU Dresden, Tatzberg 41, 01307, Dresden, Germany.
- Physics of Life, DFG Cluster of Excellence, TU Dresden, 01062, Dresden, Germany.
- Faculty of Physics, TU Dresden, 01062, Dresden, Germany.
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9
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Coñuecar R, Asela I, Rivera M, Galaz-Davison P, González-Higueras J, Hamilton GL, Engelberger F, Ramírez-Sarmiento CA, Babul J, Sanabria H, Medina E. DNA facilitates heterodimerization between human transcription factors FoxP1 and FoxP2 by increasing their conformational flexibility. iScience 2023; 26:107228. [PMID: 37485372 PMCID: PMC10362293 DOI: 10.1016/j.isci.2023.107228] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/03/2022] [Revised: 05/15/2023] [Accepted: 06/20/2023] [Indexed: 07/25/2023] Open
Abstract
Transcription factors regulate gene expression by binding to DNA. They have disordered regions and specific DNA-binding domains. Binding to DNA causes structural changes, including folding and interactions with other molecules. The FoxP subfamily of transcription factors in humans is unique because they can form heterotypic interactions without DNA. However, it is unclear how they form heterodimers and how DNA binding affects their function. We used computational and experimental methods to study the structural changes in FoxP1's DNA-binding domain when it forms a heterodimer with FoxP2. We found that FoxP1 has complex and diverse conformational dynamics, transitioning between compact and extended states. Surprisingly, DNA binding increases the flexibility of FoxP1, contrary to the typical folding-upon-binding mechanism. In addition, we observed a 3-fold increase in the rate of heterodimerization after FoxP1 binds to DNA. These findings emphasize the importance of structural flexibility in promoting heterodimerization to form transcriptional complexes.
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Affiliation(s)
- Ricardo Coñuecar
- Departamento de Biología, Facultad de Ciencias, Universidad de Chile, Santiago 7800003, Chile
| | - Isabel Asela
- Departamento de Biología, Facultad de Ciencias, Universidad de Chile, Santiago 7800003, Chile
| | - Maira Rivera
- Institute for Biological and Medical Engineering, Schools of Engineering, Medicine and Biological Sciences, Pontificia Universidad Católica de Chile, Santiago 7820436, Chile
- ANID – Millennium Science Initiative Program – Millennium Institute for Integrative Biology (iBio), Santiago 8331150, Chile
| | - Pablo Galaz-Davison
- Institute for Biological and Medical Engineering, Schools of Engineering, Medicine and Biological Sciences, Pontificia Universidad Católica de Chile, Santiago 7820436, Chile
- ANID – Millennium Science Initiative Program – Millennium Institute for Integrative Biology (iBio), Santiago 8331150, Chile
| | - Jorge González-Higueras
- Institute for Biological and Medical Engineering, Schools of Engineering, Medicine and Biological Sciences, Pontificia Universidad Católica de Chile, Santiago 7820436, Chile
- ANID – Millennium Science Initiative Program – Millennium Institute for Integrative Biology (iBio), Santiago 8331150, Chile
| | - George L. Hamilton
- Department of Biochemistry and Molecular Pharmacology, New York University School of Medicine, New York, NY, USA
| | - Felipe Engelberger
- Institute for Drug Discovery, Leipzig University Medical School, 04107 Leipzig, Germany
| | - César A. Ramírez-Sarmiento
- Institute for Biological and Medical Engineering, Schools of Engineering, Medicine and Biological Sciences, Pontificia Universidad Católica de Chile, Santiago 7820436, Chile
- ANID – Millennium Science Initiative Program – Millennium Institute for Integrative Biology (iBio), Santiago 8331150, Chile
| | - Jorge Babul
- Departamento de Biología, Facultad de Ciencias, Universidad de Chile, Santiago 7800003, Chile
| | - Hugo Sanabria
- Department of Physics & Astronomy, Clemson University, Clemson, SC 29634, USA
| | - Exequiel Medina
- Departamento de Biología, Facultad de Ciencias, Universidad de Chile, Santiago 7800003, Chile
- Department of Physics & Astronomy, Clemson University, Clemson, SC 29634, USA
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10
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He J, Dmochowski IJ. Local Xenon-Protein Interaction Produces Global Conformational Change and Allosteric Inhibition in Lysozyme. Biochemistry 2023; 62:1659-1669. [PMID: 37192381 PMCID: PMC10821772 DOI: 10.1021/acs.biochem.3c00046] [Citation(s) in RCA: 1] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 05/18/2023]
Abstract
Noble gases have well-established biological effects, yet their molecular mechanisms remain poorly understood. Here, we investigated, both experimentally and computationally, the molecular modes of xenon (Xe) action in bacteriophage T4 lysozyme (T4L). By combining indirect gassing methods with a colorimetric lysozyme activity assay, a reversible, Xe-specific (20 ± 3)% inhibition effect was observed. Accelerated molecular dynamic simulations revealed that Xe exerts allosteric inhibition on the protein by expanding a C-terminal hydrophobic cavity. Xe-induced cavity expansion results in global conformational changes, with long-range transduction distorting the active site where peptidoglycan binds. Interestingly, the peptide substrate binding site that enables lysozyme specificity does not change conformation. Two T4L mutants designed to reshape the C-terminal Xe cavity established a correlation between cavity expansion and enzyme inhibition. This work also highlights the use of Xe flooding simulations to identify new cryptic binding pockets. These results enrich our understanding of Xe-protein interactions at the molecular level and inspire further biochemical investigations with noble gases.
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Affiliation(s)
- Jiayi He
- Department of Chemistry, University of Pennsylvania, Philadelphia, Pennsylvania 19104-6323, United States
| | - Ivan J Dmochowski
- Department of Chemistry, University of Pennsylvania, Philadelphia, Pennsylvania 19104-6323, United States
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11
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Abstract
Internal motions in proteins take place on a broad range of time- and space-scales. The potential roles of these dynamics in the biochemical functions of proteins have intrigued biophysicists for many years, and multiple mechanisms to couple motions to function have been proposed. Some of these mechanisms have relied on equilibrium concepts. For example, the modulation of dynamics was proposed to change the entropy of a protein, hence affecting processes such as binding. This so-called dynamic allostery scenario has been demonstrated in several recent experiments. Perhaps even more intriguing may be models that involve out-of-equilibrium operation, which by necessity require the input of energy. We discuss several recent experimental studies that expose such potential mechanisms for coupling dynamics and function. In Brownian ratchets, for example, directional motion is promoted by switching a protein between two free energy surfaces. An additional example involves the effect of microsecond domain-closure dynamics of an enzyme on its much slower chemical cycle. These observations lead us to propose a novel two-time-scale paradigm for the activity of protein machines: fast equilibrium fluctuations take place on the microsecond-millisecond time scale, while on a slower time scale, free energy is invested in order to push the system out of equilibrium and drive functional transitions. Motions on the two time scales affect each other and are essential for the overall function of these machines.
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Affiliation(s)
- Gilad Haran
- Department of Chemical and Biological Physics, Weizmann Institute of Science, Rehovot 7610001, Israel
| | - Inbal Riven
- Department of Chemical and Biological Physics, Weizmann Institute of Science, Rehovot 7610001, Israel
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12
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Agam G, Gebhardt C, Popara M, Mächtel R, Folz J, Ambrose B, Chamachi N, Chung SY, Craggs TD, de Boer M, Grohmann D, Ha T, Hartmann A, Hendrix J, Hirschfeld V, Hübner CG, Hugel T, Kammerer D, Kang HS, Kapanidis AN, Krainer G, Kramm K, Lemke EA, Lerner E, Margeat E, Martens K, Michaelis J, Mitra J, Moya Muñoz GG, Quast RB, Robb NC, Sattler M, Schlierf M, Schneider J, Schröder T, Sefer A, Tan PS, Thurn J, Tinnefeld P, van Noort J, Weiss S, Wendler N, Zijlstra N, Barth A, Seidel CAM, Lamb DC, Cordes T. Reliability and accuracy of single-molecule FRET studies for characterization of structural dynamics and distances in proteins. Nat Methods 2023; 20:523-535. [PMID: 36973549 PMCID: PMC10089922 DOI: 10.1038/s41592-023-01807-0] [Citation(s) in RCA: 19] [Impact Index Per Article: 19.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/02/2022] [Accepted: 01/31/2023] [Indexed: 03/29/2023]
Abstract
Single-molecule Förster-resonance energy transfer (smFRET) experiments allow the study of biomolecular structure and dynamics in vitro and in vivo. We performed an international blind study involving 19 laboratories to assess the uncertainty of FRET experiments for proteins with respect to the measured FRET efficiency histograms, determination of distances, and the detection and quantification of structural dynamics. Using two protein systems with distinct conformational changes and dynamics, we obtained an uncertainty of the FRET efficiency ≤0.06, corresponding to an interdye distance precision of ≤2 Å and accuracy of ≤5 Å. We further discuss the limits for detecting fluctuations in this distance range and how to identify dye perturbations. Our work demonstrates the ability of smFRET experiments to simultaneously measure distances and avoid the averaging of conformational dynamics for realistic protein systems, highlighting its importance in the expanding toolbox of integrative structural biology.
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Affiliation(s)
- Ganesh Agam
- Department of Chemistry, Ludwig-Maximilians University München, München, Germany
| | - Christian Gebhardt
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians University München, Planegg-Martinsried, Germany
| | - Milana Popara
- Molecular Physical Chemistry, Heinrich-Heine University Düsseldorf, Düsseldorf, Germany
| | - Rebecca Mächtel
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians University München, Planegg-Martinsried, Germany
| | - Julian Folz
- Molecular Physical Chemistry, Heinrich-Heine University Düsseldorf, Düsseldorf, Germany
| | | | - Neharika Chamachi
- B CUBE - Center for Molecular Bioengineering, Technische Universität Dresden, Dresden, Germany
| | - Sang Yoon Chung
- Department of Chemistry and Biochemistry, University of California, Los Angeles, CA, USA
| | | | - Marijn de Boer
- Molecular Microscopy Research Group, Zernike Institute for Advanced Materials, University of Groningen, AG Groningen, the Netherlands
| | - Dina Grohmann
- Department of Biochemistry, Genetics and Microbiology, Institute of Microbiology, Single-Molecule Biochemistry Laboratory, University of Regensburg, Regensburg, Germany
| | - Taekjip Ha
- Department of Biophysics and Biophysical Chemistry, Johns Hopkins University School of Medicine and Howard Hughes Medical Institute, Baltimore, MD, USA
| | - Andreas Hartmann
- B CUBE - Center for Molecular Bioengineering, Technische Universität Dresden, Dresden, Germany
| | - Jelle Hendrix
- Dynamic Bioimaging Laboratory, Advanced Optical Microscopy Center and Biomedical Research Institute, Hasselt University, Agoralaan C (BIOMED), Hasselt, Belgium
- Department of Chemistry, KU Leuven, Leuven, Belgium
| | | | | | - Thorsten Hugel
- Institute of Physical Chemistry, University of Freiburg, Freiburg, Germany
- Signalling Research Centers BIOSS and CIBSS, University of Freiburg, Freiburg, Germany
| | - Dominik Kammerer
- Department of Physics, Clarendon Laboratory, University of Oxford, Oxford, UK
- Kavli Institute of Nanoscience Discovery, University of Oxford, Oxford, UK
| | - Hyun-Seo Kang
- Bayerisches NMR Zentrum, Department of Bioscience, School of Natural Sciences, Technical University of München, Garching, Germany
| | - Achillefs N Kapanidis
- Department of Physics, Clarendon Laboratory, University of Oxford, Oxford, UK
- Kavli Institute of Nanoscience Discovery, University of Oxford, Oxford, UK
| | - Georg Krainer
- B CUBE - Center for Molecular Bioengineering, Technische Universität Dresden, Dresden, Germany
- Yusuf Hamied Department of Chemistry, University of Cambridge, Cambridge, UK
| | - Kevin Kramm
- Department of Biochemistry, Genetics and Microbiology, Institute of Microbiology, Single-Molecule Biochemistry Laboratory, University of Regensburg, Regensburg, Germany
| | - Edward A Lemke
- Biocenter, Johannes Gutenberg University Mainz, Mainz, Germany
- Institute of Molecular Biology, Mainz, Germany
- Structural and Computational Biology Unit, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Eitan Lerner
- Department of Biological Chemistry, The Alexander Silberman Institute of Life Sciences, and The Center for Nanoscience and Nanotechnology, Faculty of Mathematics and Science, The Edmond J. Safra Campus, The Hebrew University of Jerusalem, Jerusalem, Israel
| | - Emmanuel Margeat
- Centre de Biologie Structurale (CBS), University of Montpellier, CNRS, INSERM, Montpellier, France
| | - Kirsten Martens
- Biological and Soft Matter Physics, Huygens-Kamerlingh Onnes Laboratory, Leiden University, Leiden, the Netherlands
| | | | - Jaba Mitra
- Department of Biophysics and Biophysical Chemistry, Johns Hopkins University School of Medicine and Howard Hughes Medical Institute, Baltimore, MD, USA
- Materials Science and Engineering, University of Illinois Urbana-Champaign, Urbana, IL, USA
| | - Gabriel G Moya Muñoz
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians University München, Planegg-Martinsried, Germany
| | - Robert B Quast
- Centre de Biologie Structurale (CBS), University of Montpellier, CNRS, INSERM, Montpellier, France
| | - Nicole C Robb
- Department of Physics, Clarendon Laboratory, University of Oxford, Oxford, UK
- Kavli Institute of Nanoscience Discovery, University of Oxford, Oxford, UK
- Warwick Medical School, The University of Warwick, Coventry, UK
| | - Michael Sattler
- Bayerisches NMR Zentrum, Department of Bioscience, School of Natural Sciences, Technical University of München, Garching, Germany
- Institute of Structural Biology, Molecular Targets and Therapeutics Center, Helmholtz Center Munich, Munich, Germany
| | - Michael Schlierf
- B CUBE - Center for Molecular Bioengineering, Technische Universität Dresden, Dresden, Germany
- Cluster of Excellence Physics of Life, Technische Universität Dresden, Dresden, Germany
| | - Jonathan Schneider
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians University München, Planegg-Martinsried, Germany
| | - Tim Schröder
- Department of Chemistry, Ludwig-Maximilians University München, München, Germany
| | - Anna Sefer
- Institute for Biophysics, Ulm University, Ulm, Germany
| | - Piau Siong Tan
- Biocenter, Johannes Gutenberg University Mainz, Mainz, Germany
- Institute of Molecular Biology, Mainz, Germany
| | - Johann Thurn
- Institute of Physical Chemistry, University of Freiburg, Freiburg, Germany
- Institute of Technical Physics, German Aerospace Center (DLR), Stuttgart, Germany
| | - Philip Tinnefeld
- Department of Chemistry, Ludwig-Maximilians University München, München, Germany
| | - John van Noort
- Biological and Soft Matter Physics, Huygens-Kamerlingh Onnes Laboratory, Leiden University, Leiden, the Netherlands
| | - Shimon Weiss
- Department of Chemistry and Biochemistry, University of California, Los Angeles, CA, USA
- California NanoSystems Institute, University of California, Los Angeles, CA, USA
| | - Nicolas Wendler
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians University München, Planegg-Martinsried, Germany
| | - Niels Zijlstra
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians University München, Planegg-Martinsried, Germany
| | - Anders Barth
- Molecular Physical Chemistry, Heinrich-Heine University Düsseldorf, Düsseldorf, Germany.
- Department of Bionanoscience, Kavli Institute of Nanoscience, Delft University of Technology, Delft, the Netherlands.
| | - Claus A M Seidel
- Molecular Physical Chemistry, Heinrich-Heine University Düsseldorf, Düsseldorf, Germany.
| | - Don C Lamb
- Department of Chemistry, Ludwig-Maximilians University München, München, Germany.
| | - Thorben Cordes
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians University München, Planegg-Martinsried, Germany.
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13
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Zhang A, Zhuang X, Liu J, Huang J, Lin L, Tang Y, Zhao S, Li R, Wang B, Fang B, Hong W. Catalytic cycle of formate dehydrogenase captured by single-molecule conductance. Nat Catal 2023. [DOI: 10.1038/s41929-023-00928-1] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 03/08/2023]
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14
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Shrinking gate fluorescence correlation spectroscopy yields equilibrium constants and separates photophysics from structural dynamics. Proc Natl Acad Sci U S A 2023; 120:e2211896120. [PMID: 36652471 PMCID: PMC9942831 DOI: 10.1073/pnas.2211896120] [Citation(s) in RCA: 1] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/19/2023] Open
Abstract
Fluorescence correlation spectroscopy is a versatile tool for studying fast conformational changes of biomolecules especially when combined with Förster resonance energy transfer (FRET). Despite the many methods available for identifying structural dynamics in FRET experiments, the determination of the forward and backward transition rate constants and thereby also the equilibrium constant is difficult when two intensity levels are involved. Here, we combine intensity correlation analysis with fluorescence lifetime information by including only a subset of photons in the autocorrelation analysis based on their arrival time with respect to the excitation pulse (microtime). By fitting the correlation amplitude as a function of microtime gate, the transition rate constants from two fluorescence-intensity level systems and the corresponding equilibrium constants are obtained. This shrinking-gate fluorescence correlation spectroscopy (sg-FCS) approach is demonstrated using simulations and with a DNA origami-based model system in experiments on immobilized and freely diffusing molecules. We further show that sg-FCS can distinguish photophysics from dynamic intensity changes even if a dark quencher, in this case graphene, is involved. Finally, we unravel the mechanism of a FRET-based membrane charge sensor indicating the broad potential of the method. With sg-FCS, we present an algorithm that does not require prior knowledge and is therefore easily implemented when an autocorrelation analysis is carried out on time-correlated single-photon data.
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15
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Godec A, Makarov DE. Challenges in Inferring the Directionality of Active Molecular Processes from Single-Molecule Fluorescence Resonance Energy Transfer Trajectories. J Phys Chem Lett 2023; 14:49-56. [PMID: 36566432 DOI: 10.1021/acs.jpclett.2c03244] [Citation(s) in RCA: 2] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 06/17/2023]
Abstract
We discuss some of the practical challenges that one faces in using stochastic thermodynamics to infer directionality of molecular machines from experimental single-molecule trajectories. Because of the limited spatiotemporal resolution of single-molecule experiments and because both forward and backward transitions between the same pairs of states cannot always be detected, differentiating between the forward and backward directions of, e.g., an ATP-consuming molecular machine that operates periodically, turns out to be a nontrivial task. Using a simple extension of a Markov-state model that is commonly employed to analyze single-molecule transition-path measurements, we illustrate how irreversibility can be hidden from such measurements but in some cases can be uncovered when non-Markov effects in low-dimensional single-molecule trajectories are considered.
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Affiliation(s)
- Aljaž Godec
- Mathematical bioPhysics Group, Max Planck Institute for Multidisciplinary Sciences, 37077Göttingen, Germany
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16
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Xi K, Zhu L. Automated Path Searching Reveals the Mechanism of Hydrolysis Enhancement by T4 Lysozyme Mutants. Int J Mol Sci 2022; 23:ijms232314628. [PMID: 36498954 PMCID: PMC9736071 DOI: 10.3390/ijms232314628] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/05/2022] [Revised: 11/16/2022] [Accepted: 11/19/2022] [Indexed: 11/25/2022] Open
Abstract
Bacteriophage T4 lysozyme (T4L) is a glycosidase that is widely applied as a natural antimicrobial agent in the food industry. Due to its wide applications and small size, T4L has been regarded as a model system for understanding protein dynamics and for large-scale protein engineering. Through structural insights from the single conformation of T4L, a series of mutations (L99A,G113A,R119P) have been introduced, which have successfully raised the fractional population of its only hydrolysis-competent excited state to 96%. However, the actual impact of these substitutions on its dynamics remains unclear, largely due to the lack of highly efficient sampling algorithms. Here, using our recently developed travelling-salesman-based automated path searching (TAPS), we located the minimum-free-energy path (MFEP) for the transition of three T4L mutants from their ground states to their excited states. All three mutants share a three-step transition: the flipping of F114, the rearrangement of α0/α1 helices, and final refinement. Remarkably, the MFEP revealed that the effects of the mutations are drastically beyond the expectations of their original design: (a) the G113A substitution not only enhances helicity but also fills the hydrophobic Cavity I and reduces the free energy barrier for flipping F114; (b) R119P barely changes the stability of the ground state but stabilizes the excited state through rarely reported polar contacts S117OG:N132ND2, E11OE1:R145NH1, and E11OE2:Q105NE2; (c) the residue W138 flips into Cavity I and further stabilizes the excited state for the triple mutant L99A,G113A,R119P. These novel insights that were unexpected in the original mutant design indicated the necessity of incorporating path searching into the workflow of rational protein engineering.
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17
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Henderson J, Havranek O, Ma MCJ, Herman V, Kupcova K, Chrbolkova T, Pacheco-Blanco M, Wang Z, Comer JM, Zal T, Davis RE. Detecting Förster resonance energy transfer in living cells by conventional and spectral flow cytometry. Cytometry A 2022; 101:818-834. [PMID: 34128311 DOI: 10.1002/cyto.a.24472] [Citation(s) in RCA: 6] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/21/2020] [Revised: 05/31/2021] [Accepted: 06/07/2021] [Indexed: 01/27/2023]
Abstract
Assays based on Förster resonance energy transfer (FRET) can be used to study many processes in cell biology. Although this is most often done with microscopy for fluorescence detection, we report two ways to measure FRET in living cells by flow cytometry. Using a conventional flow cytometer and the "3-cube method" for intensity-based calculation of FRET efficiency, we measured the enzymatic activity of specific kinases in cells expressing a genetically-encoded reporter. For both AKT and protein kinase A, the method measured kinase activity in time-course, dose-response, and kinetic assays. Using the Cytek Aurora spectral flow cytometer, which applies linear unmixing to emission measured in multiple wavelength ranges, FRET from the same reporters was measured with greater single-cell precision, in real time and in the presence of other fluorophores. Results from gene-knockout studies suggested that spectral flow cytometry might enable the sorting of cells on the basis of FRET. The methods we present provide convenient and flexible options for using FRET with flow cytometry in studies of cell biology.
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Affiliation(s)
- Jared Henderson
- Department of Lymphoma and Myeloma, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA
| | - Ondrej Havranek
- Department of Lymphoma and Myeloma, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA.,BIOCEV, First Faculty of Medicine, Charles University, Vestec, Czech Republic.,Department of Hematology, Charles University and General University Hospital, Prague, Czech Republic
| | - Man Chun John Ma
- Department of Lymphoma and Myeloma, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA
| | - Vaclav Herman
- BIOCEV, First Faculty of Medicine, Charles University, Vestec, Czech Republic.,Department of Hematology, Charles University and General University Hospital, Prague, Czech Republic
| | - Kristyna Kupcova
- BIOCEV, First Faculty of Medicine, Charles University, Vestec, Czech Republic
| | - Tereza Chrbolkova
- BIOCEV, First Faculty of Medicine, Charles University, Vestec, Czech Republic
| | | | - Zhiqiang Wang
- Department of Lymphoma and Myeloma, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA
| | - Justin M Comer
- Department of Lymphoma and Myeloma, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA
| | - Tomasz Zal
- Department of Leukemia, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA
| | - Richard Eric Davis
- Department of Lymphoma and Myeloma, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA.,Department of Translational Molecular Pathology, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA
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18
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Imam N, Choudhury S, Heinze KG, Schindelin H. Differential modulation of collybistin conformational dynamics by the closely related GTPases Cdc42 and TC10. Front Synaptic Neurosci 2022; 14:959875. [PMID: 35989712 PMCID: PMC9386560 DOI: 10.3389/fnsyn.2022.959875] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/02/2022] [Accepted: 07/06/2022] [Indexed: 11/13/2022] Open
Abstract
Interneuronal synaptic transmission relies on the proper spatial organization of presynaptic neurotransmitter release and its reception on the postsynaptic side by cognate neurotransmitter receptors. Neurotransmitter receptors are incorporated into and arranged within the plasma membrane with the assistance of scaffolding and adaptor proteins. At inhibitory GABAergic postsynapses, collybistin, a neuronal adaptor protein, recruits the scaffolding protein gephyrin and interacts with various neuronal factors including cell adhesion proteins of the neuroligin family, the GABAA receptor α2-subunit and the closely related small GTPases Cdc42 and TC10 (RhoQ). Most collybistin splice variants harbor an N-terminal SH3 domain and exist in an autoinhibited/closed state. Cdc42 and TC10, despite sharing 67.4% amino acid sequence identity, interact differently with collybistin. Here, we delineate the molecular basis of the collybistin conformational activation induced by TC10 with the aid of recently developed collybistin FRET sensors. Time-resolved fluorescence-based FRET measurements reveal that TC10 binds to closed/inactive collybistin leading to relief of its autoinhibition, contrary to Cdc42, which only interacts with collybistin when forced into an open state by the introduction of mutations destabilizing the closed state of collybistin. Taken together, our data describe a TC10-driven signaling mechanism in which collybistin switches from its autoinhibited closed state to an open/active state.
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Affiliation(s)
- Nasir Imam
- Institute of Structural Biology, Rudolf Virchow Center for Integrative and Translational Bioimaging, University of Würzburg, Würzburg, Germany
| | - Susobhan Choudhury
- Molecular Microscopy, Rudolf Virchow Center for Integrative and Translational Bioimaging, University of Würzburg, Würzburg, Germany
| | - Katrin G. Heinze
- Molecular Microscopy, Rudolf Virchow Center for Integrative and Translational Bioimaging, University of Würzburg, Würzburg, Germany
- Katrin G. Heinze,
| | - Hermann Schindelin
- Institute of Structural Biology, Rudolf Virchow Center for Integrative and Translational Bioimaging, University of Würzburg, Würzburg, Germany
- *Correspondence: Hermann Schindelin,
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19
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Peter MF, Gebhardt C, Mächtel R, Muñoz GGM, Glaenzer J, Narducci A, Thomas GH, Cordes T, Hagelueken G. Cross-validation of distance measurements in proteins by PELDOR/DEER and single-molecule FRET. Nat Commun 2022; 13:4396. [PMID: 35906222 PMCID: PMC9338047 DOI: 10.1038/s41467-022-31945-6] [Citation(s) in RCA: 13] [Impact Index Per Article: 6.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/02/2020] [Accepted: 07/11/2022] [Indexed: 11/09/2022] Open
Abstract
Pulsed electron-electron double resonance spectroscopy (PELDOR/DEER) and single-molecule Förster resonance energy transfer spectroscopy (smFRET) are frequently used to determine conformational changes, structural heterogeneity, and inter probe distances in biological macromolecules. They provide qualitative information that facilitates mechanistic understanding of biochemical processes and quantitative data for structural modelling. To provide a comprehensive comparison of the accuracy of PELDOR/DEER and smFRET, we use a library of double cysteine variants of four proteins that undergo large-scale conformational changes upon ligand binding. With either method, we use established standard experimental protocols and data analysis routines to determine inter-probe distances in the presence and absence of ligands. The results are compared to distance predictions from structural models. Despite an overall satisfying and similar distance accuracy, some inconsistencies are identified, which we attribute to the use of cryoprotectants for PELDOR/DEER and label-protein interactions for smFRET. This large-scale cross-validation of PELDOR/DEER and smFRET highlights the strengths, weaknesses, and synergies of these two important and complementary tools in integrative structural biology.
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Affiliation(s)
- Martin F Peter
- Institute of Structural Biology, University of Bonn, Bonn, Germany
| | - Christian Gebhardt
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians-Universität München, Planegg-Martinsried, Germany
| | - Rebecca Mächtel
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians-Universität München, Planegg-Martinsried, Germany
| | - Gabriel G Moya Muñoz
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians-Universität München, Planegg-Martinsried, Germany
| | - Janin Glaenzer
- Institute of Structural Biology, University of Bonn, Bonn, Germany
| | - Alessandra Narducci
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians-Universität München, Planegg-Martinsried, Germany
| | - Gavin H Thomas
- Department of Biology (Area 10), University of York, York, UK
| | - Thorben Cordes
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians-Universität München, Planegg-Martinsried, Germany.
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20
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Chinnaraj M, Flaumenhaft R, Pozzi N. Reduction of protein disulfide isomerase results in open conformations and stimulates dynamic exchange between structural ensembles. J Biol Chem 2022; 298:102217. [PMID: 35780832 PMCID: PMC9352907 DOI: 10.1016/j.jbc.2022.102217] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/23/2022] [Revised: 06/25/2022] [Accepted: 06/27/2022] [Indexed: 11/19/2022] Open
Abstract
Human protein disulfide isomerase (PDI) is an essential redox-regulated enzyme required for oxidative protein folding. It comprises four thioredoxin domains, two catalytically active (a, a’) and two inactive (b, b’), organized to form a flexible abb’a’ U-shape. Snapshots of unbound oxidized and reduced PDI have been obtained by X-ray crystallography. Yet, how PDI’s structure changes in response to the redox environment and inhibitor binding remains controversial. Here, we used multiparameter confocal single-molecule FRET to track the movements of the two catalytic domains with high temporal resolution. We found that at equilibrium, PDI visits three structurally distinct conformational ensembles, two “open” (O1 and O2) and one “closed” (C). We show that the redox environment dictates the time spent in each ensemble and the rate at which they exchange. While oxidized PDI samples O1, O2, and C more evenly and in a slower fashion, reduced PDI predominantly populates O1 and O2 and exchanges between them more rapidly, on the submillisecond timescale. These findings were not expected based on crystallographic data. Using mutational analyses, we further demonstrate that the R300-W396 cation-π interaction and active site cysteines dictate, in unexpected ways, how the catalytic domains relocate. Finally, we show that irreversible inhibitors targeting the active sites of reduced PDI did not abolish these protein dynamics but rather shifted the equilibrium toward the closed ensemble. This work introduces a new structural framework that challenges current views of PDI dynamics, helps rationalize its multifaceted role in biology, and should be considered when designing PDI-targeted therapeutics.
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Affiliation(s)
- Mathivanan Chinnaraj
- Edward A. Doisy Department of Biochemistry and Molecular Biology, Saint Louis University School of Medicine, St. Louis, MO, 63104, USA
| | - Robert Flaumenhaft
- Division of Hemostasis and Thrombosis, Department of Medicine, Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, Massachusetts 02115, USA.
| | - Nicola Pozzi
- Edward A. Doisy Department of Biochemistry and Molecular Biology, Saint Louis University School of Medicine, St. Louis, MO, 63104, USA.
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21
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Opanasyuk O, Barth A, Peulen TO, Felekyan S, Kalinin S, Sanabria H, Seidel CA. Unraveling multi-state molecular dynamics in single-molecule FRET experiments. II. Quantitative analysis of multi-state kinetic networks. J Chem Phys 2022; 157:031501. [DOI: 10.1063/5.0095754] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/14/2022] Open
Abstract
Single-molecule Förster Resonance Energy Transfer (smFRET) experiments are ideally suited to resolve the structural dynamics of biomolecules. A significant challenge to date is capturing and quantifying the exchange between multiple conformational states, mainly when these dynamics occur on the sub-millisecond timescale. Many methods for quantitative analysis are challenged if more than two states are involved, and the appropriate choice of the number of states in the kinetic network is difficult. An additional complication arises if dynamically active molecules coexist with pseudo-static molecules in similar conformational states with undistinguishable FRET efficiencies. To address these problems, we developed a quantitative integrative analysis framework that combines the information from FRET-lines that relate average fluorescence lifetimes and intensities in two-dimensional burst frequency histograms, fluorescence decays obtained by time-correlated single photon counting, photon distribution analysis of the intensities and fluorescence correlation spectroscopy. Individually, these methodologies provide ambiguous results for the characterization of dynamics in complex kinetic networks. However, the global analysis approach enables accurate determination of the number of states, their kinetic connectivity, the transition rate constants, and species fractions. To challenge the potential of smFRET experiments studying multi-state kinetic networks, we apply our integrative framework using a set of synthetic data for three-state systems with different kinetic connectivity and exchange rates. Our methodology paves the way towards an integrated analysis of multiparameter smFRET experiments that spans all dimensions of the experimental data. Finally, we propose a workflow for the analysis and show examples that demonstrate the usefulness of this toolkit for dynamic structural biology.
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Affiliation(s)
| | | | | | - Suren Felekyan
- PC-II, Heinrich Heine University Düsseldorf Department of Chemistry, Germany
| | - Stanislav Kalinin
- Institut für Physikalische Chemie, Heinrich-Heine-Universität Düsseldorf, Germany
| | | | - Claus A.M. Seidel
- Institut fuer Physikalische Chemie, Heinrich-Heine-Universität Düsseldorf, Germany
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22
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Post M, Lickert B, Diez G, Wolf S, Stock G. Cooperative Protein Allosteric Transition Mediated by a Fluctuating Transmission Network. J Mol Biol 2022; 434:167679. [PMID: 35690098 DOI: 10.1016/j.jmb.2022.167679] [Citation(s) in RCA: 7] [Impact Index Per Article: 3.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/28/2022] [Revised: 06/02/2022] [Accepted: 06/06/2022] [Indexed: 12/13/2022]
Abstract
Allosteric communication between distant protein sites represents a key mechanism of biomolecular regulation and signal transduction. Compared to other processes such as protein folding, however, the dynamical evolution of allosteric transitions is still not well understood. As an example of allosteric coupling between distant protein regions, we consider the global open-closed motion of the two domains of T4 lysozyme, which is triggered by local motion in the hinge region. Combining extensive molecular dynamics simulations with a correlation analysis of interresidue contacts, we identify a network of interresidue distances that move in a concerted manner. The cooperative process originates from a cogwheel-like motion of the hydrophobic core in the hinge region, which constitutes an evolutionary conserved and flexible transmission network. Through rigid contacts and the protein backbone, the small local changes of the hydrophobic core are passed on to the distant terminal domains and lead to the emergence of a rare global conformational transition. As in an Ising-type model, the cooperativity of the allosteric transition can be explained via the interaction of local fluctuations.
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Affiliation(s)
- Matthias Post
- Biomolecular Dynamics, Institute of Physics, Albert Ludwigs University, 79104 Freiburg, Germany. https://twitter.com/@_posti
| | - Benjamin Lickert
- Biomolecular Dynamics, Institute of Physics, Albert Ludwigs University, 79104 Freiburg, Germany. https://twitter.com/@BenjaminLickert
| | - Georg Diez
- Biomolecular Dynamics, Institute of Physics, Albert Ludwigs University, 79104 Freiburg, Germany. https://twitter.com/@gegadiez
| | - Steffen Wolf
- Biomolecular Dynamics, Institute of Physics, Albert Ludwigs University, 79104 Freiburg, Germany.
| | - Gerhard Stock
- Biomolecular Dynamics, Institute of Physics, Albert Ludwigs University, 79104 Freiburg, Germany.
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23
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Barth A, Opanasyuk O, Peulen TO, Felekyan S, Kalinin S, Sanabria H, Seidel CAM. Unraveling multi-state molecular dynamics in single-molecule FRET experiments. I. Theory of FRET-lines. J Chem Phys 2022; 156:141501. [PMID: 35428384 PMCID: PMC9014241 DOI: 10.1063/5.0089134] [Citation(s) in RCA: 14] [Impact Index Per Article: 7.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/31/2023] Open
Abstract
Conformational dynamics of biomolecules are of fundamental importance for their function. Single-molecule studies of Förster Resonance Energy Transfer (smFRET) between a tethered donor and acceptor dye pair are a powerful tool to investigate the structure and dynamics of labeled molecules. However, capturing and quantifying conformational dynamics in intensity-based smFRET experiments remains challenging when the dynamics occur on the sub-millisecond timescale. The method of multiparameter fluorescence detection addresses this challenge by simultaneously registering fluorescence intensities and lifetimes of the donor and acceptor. Together, two FRET observables, the donor fluorescence lifetime τD and the intensity-based FRET efficiency E, inform on the width of the FRET efficiency distribution as a characteristic fingerprint for conformational dynamics. We present a general framework for analyzing dynamics that relates average fluorescence lifetimes and intensities in two-dimensional burst frequency histograms. We present parametric relations of these observables for interpreting the location of FRET populations in E–τD diagrams, called FRET-lines. To facilitate the analysis of complex exchange equilibria, FRET-lines serve as reference curves for a graphical interpretation of experimental data to (i) identify conformational states, (ii) resolve their dynamic connectivity, (iii) compare different kinetic models, and (iv) infer polymer properties of unfolded or intrinsically disordered proteins. For a simplified graphical analysis of complex kinetic networks, we derive a moment-based representation of the experimental data that decouples the motion of the fluorescence labels from the conformational dynamics of the biomolecule. Importantly, FRET-lines facilitate exploring complex dynamic models via easily computed experimental observables. We provide extensive computational tools to facilitate applying FRET-lines.
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Affiliation(s)
- Anders Barth
- Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, Düsseldorf, Germany
| | - Oleg Opanasyuk
- Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, Düsseldorf, Germany
| | - Thomas-Otavio Peulen
- Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, Düsseldorf, Germany
| | - Suren Felekyan
- Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, Düsseldorf, Germany
| | - Stanislav Kalinin
- Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, Düsseldorf, Germany
| | - Hugo Sanabria
- Department of Physics and Astronomy, Clemson University, Clemson, South Carolina 29631, USA
| | - Claus A. M. Seidel
- Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, Düsseldorf, Germany
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24
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Naudi-Fabra S, Tengo M, Jensen MR, Blackledge M, Milles S. Quantitative Description of Intrinsically Disordered Proteins Using Single-Molecule FRET, NMR, and SAXS. J Am Chem Soc 2021; 143:20109-20121. [PMID: 34817999 PMCID: PMC8662727 DOI: 10.1021/jacs.1c06264] [Citation(s) in RCA: 22] [Impact Index Per Article: 7.3] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/17/2021] [Indexed: 12/18/2022]
Abstract
Studying the conformational landscape of intrinsically disordered and partially folded proteins is challenging and only accessible to a few solution state techniques, such as nuclear magnetic resonance (NMR), small-angle scattering techniques, and single-molecule Förster resonance energy transfer (smFRET). While each of the techniques is sensitive to different properties of the disordered chain, such as local structural propensities, overall dimension, or intermediate- and long-range contacts, conformational ensembles describing intrinsically disordered proteins (IDPs) accurately should ideally respect all of these properties. Here we develop an integrated approach using a large set of FRET efficiencies and fluorescence lifetimes, NMR chemical shifts, and paramagnetic relaxation enhancements (PREs), as well as small-angle X-ray scattering (SAXS) to derive quantitative conformational ensembles in agreement with all parameters. Our approach is tested using simulated data (five sets of PREs and 15 FRET efficiencies) and validated experimentally on the example of the disordered domain of measles virus phosphoprotein, providing new insights into the conformational landscape of this viral protein that comprises transient structural elements and is more compact than an unfolded chain throughout its length. Rigorous cross-validation using FRET efficiencies, fluorescence lifetimes, and SAXS demonstrates the predictive nature of the calculated conformational ensembles and underlines the potential of this strategy in integrative dynamic structural biology.
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Affiliation(s)
- Samuel Naudi-Fabra
- Institut de Biologie Structurale,
Université Grenoble Alpes-CEA-CNRS, 71, Avenue des Martyrs, 38044 Grenoble, France
| | - Maud Tengo
- Institut de Biologie Structurale,
Université Grenoble Alpes-CEA-CNRS, 71, Avenue des Martyrs, 38044 Grenoble, France
| | - Malene Ringkjøbing Jensen
- Institut de Biologie Structurale,
Université Grenoble Alpes-CEA-CNRS, 71, Avenue des Martyrs, 38044 Grenoble, France
| | - Martin Blackledge
- Institut de Biologie Structurale,
Université Grenoble Alpes-CEA-CNRS, 71, Avenue des Martyrs, 38044 Grenoble, France
| | - Sigrid Milles
- Institut de Biologie Structurale,
Université Grenoble Alpes-CEA-CNRS, 71, Avenue des Martyrs, 38044 Grenoble, France
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25
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Saikia N, Yanez-Orozco IS, Qiu R, Hao P, Milikisiyants S, Ou E, Hamilton GL, Weninger KR, Smirnova TI, Sanabria H, Ding F. Integrative structural dynamics probing of the conformational heterogeneity in synaptosomal-associated protein 25. CELL REPORTS. PHYSICAL SCIENCE 2021; 2:100616. [PMID: 34888535 PMCID: PMC8654206 DOI: 10.1016/j.xcrp.2021.100616] [Citation(s) in RCA: 9] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 05/28/2023]
Abstract
SNAP-25 (synaptosomal-associated protein of 25 kDa) is a prototypical intrinsically disordered protein (IDP) that is unstructured by itself but forms coiled-coil helices in the SNARE complex. With high conformational heterogeneity, detailed structural dynamics of unbound SNAP-25 remain elusive. Here, we report an integrative method to probe the structural dynamics of SNAP-25 by combining replica-exchange discrete molecular dynamics (rxDMD) simulations and label-based experiments at ensemble and single-molecule levels. The rxDMD simulations systematically characterize the coil-to-molten globular transition and reconstruct structural ensemble consistent with prior ensemble experiments. Label-based experiments using Förster resonance energy transfer and double electron-electron resonance further probe the conformational dynamics of SNAP-25. Agreements between simulations and experiments under both ensemble and single-molecule conditions allow us to assign specific helix-coil transitions in SNAP-25 that occur in submillisecond timescales and potentially play a vital role in forming the SNARE complex. We expect that this integrative approach may help further our understanding of IDPs.
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Affiliation(s)
- Nabanita Saikia
- Department of Physics and Astronomy, Clemson University, Clemson, SC 29634, USA
- Department of Chemistry, Navajo Technical University, Chinle, AZ 86503, USA
| | | | - Ruoyi Qiu
- Department of Physics, North Carolina State University, Raleigh, NC 27695, USA
| | - Pengyu Hao
- Department of Physics, North Carolina State University, Raleigh, NC 27695, USA
| | - Sergey Milikisiyants
- Department of Chemistry, North Carolina State University, Raleigh, NC 27695, USA
| | - Erkang Ou
- Department of Chemistry, North Carolina State University, Raleigh, NC 27695, USA
| | - George L. Hamilton
- Department of Physics and Astronomy, Clemson University, Clemson, SC 29634, USA
| | - Keith R. Weninger
- Department of Physics, North Carolina State University, Raleigh, NC 27695, USA
| | - Tatyana I. Smirnova
- Department of Chemistry, North Carolina State University, Raleigh, NC 27695, USA
| | - Hugo Sanabria
- Department of Physics and Astronomy, Clemson University, Clemson, SC 29634, USA
| | - Feng Ding
- Department of Physics and Astronomy, Clemson University, Clemson, SC 29634, USA
- Lead contact
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26
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Lv X, Jin K, Sun G, Ledesma-Amaro R, Liu L. Microscopy imaging of living cells in metabolic engineering. Trends Biotechnol 2021; 40:752-765. [PMID: 34799183 DOI: 10.1016/j.tibtech.2021.10.010] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/30/2021] [Revised: 10/25/2021] [Accepted: 10/25/2021] [Indexed: 01/23/2023]
Abstract
Microscopy imaging of living cells is becoming a pivotal, noninvasive, and highly specific tool in metabolic engineering to visualize molecular dynamics in industrial microorganisms. This review describes the different microscopy methods, from fluorescence to super resolution, with application in microbial bioengineering. Firstly, the role and importance of microscopy imaging is analyzed in the context of strain design. Then, the advantages and disadvantages of different microscopy technologies are discussed, including confocal laser scanning microscopy (CLSM), spatial light interference microscopy (SLIM), and super-resolution microscopy, followed by their applications in synthetic biology. Finally, the future perspectives of live-cell imaging and their potential to transform microbial systems are analyzed. This review provides theoretical guidance and highlights the importance of microscopy in understanding and engineering microbial metabolism.
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Affiliation(s)
- Xueqin Lv
- Key Laboratory of Carbohydrate Chemistry and Biotechnology, Ministry of Education, Jiangnan University, Wuxi 214122, China; Science Center for Future Foods, Jiangnan University, Wuxi 214122, China
| | - Ke Jin
- Key Laboratory of Carbohydrate Chemistry and Biotechnology, Ministry of Education, Jiangnan University, Wuxi 214122, China; Science Center for Future Foods, Jiangnan University, Wuxi 214122, China
| | - Guoyun Sun
- Key Laboratory of Carbohydrate Chemistry and Biotechnology, Ministry of Education, Jiangnan University, Wuxi 214122, China; Science Center for Future Foods, Jiangnan University, Wuxi 214122, China
| | - Rodrigo Ledesma-Amaro
- Department of Bioengineering and Imperial College Centre for Synthetic Biology, Imperial College London, London SW72AZ, UK
| | - Long Liu
- Key Laboratory of Carbohydrate Chemistry and Biotechnology, Ministry of Education, Jiangnan University, Wuxi 214122, China; Science Center for Future Foods, Jiangnan University, Wuxi 214122, China.
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27
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Carlon-Andres I, Malinauskas T, Padilla-Parra S. Structure dynamics of HIV-1 Env trimers on native virions engaged with living T cells. Commun Biol 2021; 4:1228. [PMID: 34707229 PMCID: PMC8551276 DOI: 10.1038/s42003-021-02658-1] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/23/2020] [Accepted: 09/09/2021] [Indexed: 11/29/2022] Open
Abstract
The HIV-1 envelope glycoprotein (Env) mediates viral entry into the host cell. Although the highly dynamic nature of Env intramolecular conformations has been shown with single molecule spectroscopy in vitro, the bona fide Env intra- and intermolecular mechanics when engaged with live T cells remains unknown. We used two photon fast fluorescence lifetime imaging detection of single-molecule Förster Resonance Energy Transfer occurring between fluorescent labels on HIV-1 Env on native virions. Our observations reveal Env dynamics at two levels: transitions between different intramolecular conformations and intermolecular interactions between Env within the viral membrane. Furthermore, we show that three broad neutralizing anti-Env antibodies directed to different epitopes restrict Env intramolecular dynamics and interactions between adjacent Env molecules when engaged with living T cells. Importantly, our results show that Env-Env interactions depend on efficient virus maturation, and that is disrupted upon binding of Env to CD4 or by neutralizing antibodies. Thus, this study illuminates how different intramolecular conformations and distribution of Env molecules mediate HIV-1 Env-T cell interactions in real time and therefore might control immune evasion.
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Affiliation(s)
- Irene Carlon-Andres
- Department of Infectious Diseases, King's College London, Faculty of Life Sciences & Medicine, London, United Kingdom.
- Randall Division of Cell and Molecular Biophysics, King's College London, London, United Kingdom.
- Division of Structural Biology, Wellcome Centre for Human Genetics, University of Oxford, Oxford, United Kingdom.
| | - Tomas Malinauskas
- Division of Structural Biology, Wellcome Centre for Human Genetics, University of Oxford, Oxford, United Kingdom
| | - Sergi Padilla-Parra
- Department of Infectious Diseases, King's College London, Faculty of Life Sciences & Medicine, London, United Kingdom.
- Randall Division of Cell and Molecular Biophysics, King's College London, London, United Kingdom.
- Division of Structural Biology, Wellcome Centre for Human Genetics, University of Oxford, Oxford, United Kingdom.
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28
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Ciferri A. Critical issues in molecular recognition: the enzyme-substrate association. SOFT MATTER 2021; 17:8585-8589. [PMID: 34553206 DOI: 10.1039/d1sm00470k] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/13/2023]
Abstract
Chemical and shape recognition are the main assembling mechanisms of complex bio-structures. A refined assessment of relevant thermodynamic parameters includes the consideration of a variety of contributions from different molecular motions and electronic interactions. An additional refinement includes the analysis of recognition involving multiple distant partners. This perspective note highlights some of the above issues in the cases of fibrillogenesis, and enzyme-substrate and antigen-antibody associations. Translational motions are found to be particularly relevant to the fibrillogenesis process. The assembly of enzyme-substrate complexes is discussed in terms of a dynamic equilibrium process.
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29
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Kolimi N, Pabbathi A, Saikia N, Ding F, Sanabria H, Alper J. Out-of-Equilibrium Biophysical Chemistry: The Case for Multidimensional, Integrated Single-Molecule Approaches. J Phys Chem B 2021; 125:10404-10418. [PMID: 34506140 PMCID: PMC8474109 DOI: 10.1021/acs.jpcb.1c02424] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/22/2022]
Abstract
![]()
Out-of-equilibrium
processes are ubiquitous across living organisms
and all structural hierarchies of life. At the molecular scale, out-of-equilibrium
processes (for example, enzyme catalysis, gene regulation, and motor
protein functions) cause biological macromolecules to sample an ensemble
of conformations over a wide range of time scales. Quantifying and
conceptualizing the structure–dynamics to function relationship
is challenging because continuously evolving multidimensional energy
landscapes are necessary to describe nonequilibrium biological processes
in biological macromolecules. In this perspective, we explore the
challenges associated with state-of-the-art experimental techniques
to understanding biological macromolecular function. We argue that
it is time to revisit how we probe and model functional out-of-equilibrium
biomolecular dynamics. We suggest that developing integrated single-molecule
multiparametric force–fluorescence instruments and using advanced
molecular dynamics simulations to study out-of-equilibrium biomolecules
will provide a path towards understanding the principles of and mechanisms
behind the structure–dynamics to function paradigm in biological
macromolecules.
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Affiliation(s)
- Narendar Kolimi
- Department of Physics and Astronomy, Clemson University, Clemson, South Carolina 29634, United States
| | - Ashok Pabbathi
- Department of Physics and Astronomy, Clemson University, Clemson, South Carolina 29634, United States
| | - Nabanita Saikia
- Department of Physics and Astronomy, Clemson University, Clemson, South Carolina 29634, United States
| | - Feng Ding
- Department of Physics and Astronomy, Clemson University, Clemson, South Carolina 29634, United States
| | - Hugo Sanabria
- Department of Physics and Astronomy, Clemson University, Clemson, South Carolina 29634, United States
| | - Joshua Alper
- Department of Physics and Astronomy, Clemson University, Clemson, South Carolina 29634, United States.,Department of Biological Sciences, Clemson University, Clemson, South Carolina 29634, United States
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30
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Steffen FD, Sigel RKO, Börner R. FRETraj: Integrating single-molecule spectroscopy with molecular dynamics. Bioinformatics 2021; 37:3953-3955. [PMID: 34478493 DOI: 10.1093/bioinformatics/btab615] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/28/2021] [Revised: 07/17/2021] [Accepted: 09/01/2021] [Indexed: 11/14/2022] Open
Abstract
SUMMARY Quantitative interpretation of single-molecule FRET experiments requires a model of the dye dynamics to link experimental energy transfer efficiencies to distances between atom positions. We have developed FRETraj, a Python module to predict FRET distributions based on accessible-contact volumes (ACV) and simulated photon statistics. FRETraj helps to identify optimal fluorophore positions on a biomolecule of interest by rapidly evaluating donor-acceptor distances. FRETraj is scalable and fully integrated into PyMOL and the Jupyter ecosystem. Here we describe the conformational dynamics of a DNA hairpin by computing multiple ACVs along a molecular dynamics trajectory and compare the predicted FRET distribution with single-molecule experiments. FRET-assisted modeling will accelerate the analysis of structural ensembles in particular dynamic, non-coding RNAs and transient protein-nucleic acid complexes. AVAILABILITY FRETraj is implemented as a cross-platform Python package available under the GPL-3.0 on Github (https://github.com/RNA-FRETools/fretraj) and is documented at https://RNA-FRETools.github.io/fretraj. SUPPLEMENTARY INFORMATION Supplementary data are available at Bioinformatics online.
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Affiliation(s)
| | | | - Richard Börner
- Department of Chemistry, University of Zurich, Switzerland
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31
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Torricella F, Bonucci A, Polykretis P, Cencetti F, Banci L. Rapid protein delivery to living cells for biomolecular investigation. Biochem Biophys Res Commun 2021; 570:82-88. [PMID: 34274850 DOI: 10.1016/j.bbrc.2021.07.006] [Citation(s) in RCA: 7] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Abstract] [Key Words] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/01/2021] [Accepted: 07/04/2021] [Indexed: 12/24/2022]
Abstract
The lack of a simple, fast and efficient method for protein delivery is limiting the widespread application of in-cell experiments, which are crucial for understanding the cellular function. We present here an innovative strategy to deliver proteins into both prokaryotic and eukaryotic cells, exploiting thermal vesiculation. This method allows to internalize substantial amounts of proteins, with different molecular weight and conformation, without compromising the structural properties and cell viability. Characterizing proteins in a physiological environment is essential as the environment can dramatically affect the conformation and dynamics of biomolecules as shown by in-cell EPR spectra vs those acquired in buffer solution. Considering its versatility, this method opens the possibility to scientists to study proteins directly in living cells through a wide range of techniques.
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Affiliation(s)
- Francesco Torricella
- Magnetic Resonance Center - CERM, University of Florence, via Luigi Sacconi 6, 50019, Sesto Fiorentino, Florence, Italy
| | - Alessio Bonucci
- Aix Marseille, Univ, CNRS, BIP, Laboratoire de Bioénergétique et Ingégnerie des protéines, Marseille, France
| | - Panagis Polykretis
- Interuniversity Consortium for Magnetic Resonance of Metallo Proteins (CIRMMP), via Luigi Sacconi 6, 50019, Sesto Fiorentino, Florence, Italy
| | - Francesca Cencetti
- Department of Experimental and Clinical Biomedical Sciences "Mario Serio", University of Florence, v.le GB Morgagni 50, 50134, Florence, Italy
| | - Lucia Banci
- Magnetic Resonance Center - CERM, University of Florence, via Luigi Sacconi 6, 50019, Sesto Fiorentino, Florence, Italy; Interuniversity Consortium for Magnetic Resonance of Metallo Proteins (CIRMMP), via Luigi Sacconi 6, 50019, Sesto Fiorentino, Florence, Italy; Department of Chemistry, University of Florence, via della Lastruccia 3, 50019, Sesto Fiorentino, Florence, Italy.
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32
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Dziuba D, Didier P, Ciaco S, Barth A, Seidel CAM, Mély Y. Fundamental photophysics of isomorphic and expanded fluorescent nucleoside analogues. Chem Soc Rev 2021; 50:7062-7107. [PMID: 33956014 DOI: 10.1039/d1cs00194a] [Citation(s) in RCA: 29] [Impact Index Per Article: 9.7] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/24/2022]
Abstract
Fluorescent nucleoside analogues (FNAs) are structurally diverse mimics of the natural essentially non-fluorescent nucleosides which have found numerous applications in probing the structure and dynamics of nucleic acids as well as their interactions with various biomolecules. In order to minimize disturbance in the labelled nucleic acid sequences, the FNA chromophoric groups should resemble the natural nucleobases in size and hydrogen-bonding patterns. Isomorphic and expanded FNAs are the two groups that best meet the criteria of non-perturbing fluorescent labels for DNA and RNA. Significant progress has been made over the past decades in understanding the fundamental photophysics that governs the spectroscopic and environmentally sensitive properties of these FNAs. Herein, we review recent advances in the spectroscopic and computational studies of selected isomorphic and expanded FNAs. We also show how this information can be used as a rational basis to design new FNAs, select appropriate sequences for optimal spectroscopic response and interpret fluorescence data in FNA applications.
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Affiliation(s)
- Dmytro Dziuba
- Laboratoire de Bioimagerie et Pathologies, UMR 7021, Université de Strasbourg, 74 route du Rhin, 67401 Illkirch, France.
| | - Pascal Didier
- Laboratoire de Bioimagerie et Pathologies, UMR 7021, Université de Strasbourg, 74 route du Rhin, 67401 Illkirch, France.
| | - Stefano Ciaco
- Laboratoire de Bioimagerie et Pathologies, UMR 7021, Université de Strasbourg, 74 route du Rhin, 67401 Illkirch, France. and Department of Biotechnology, Chemistry and Pharmacy, University of Siena, via Aldo Moro 2, 53100 Siena, Italy
| | - Anders Barth
- Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, 40225 Düsseldorf, Germany
| | - Claus A M Seidel
- Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, 40225 Düsseldorf, Germany
| | - Yves Mély
- Laboratoire de Bioimagerie et Pathologies, UMR 7021, Université de Strasbourg, 74 route du Rhin, 67401 Illkirch, France.
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33
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Zareie P, Szeto C, Farenc C, Gunasinghe SD, Kolawole EM, Nguyen A, Blyth C, Sng XYX, Li J, Jones CM, Fulcher AJ, Jacobs JR, Wei Q, Wojciech L, Petersen J, Gascoigne NRJ, Evavold BD, Gaus K, Gras S, Rossjohn J, La Gruta NL. Canonical T cell receptor docking on peptide-MHC is essential for T cell signaling. Science 2021; 372:372/6546/eabe9124. [PMID: 34083463 DOI: 10.1126/science.abe9124] [Citation(s) in RCA: 36] [Impact Index Per Article: 12.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/22/2020] [Accepted: 04/23/2021] [Indexed: 12/23/2022]
Abstract
T cell receptor (TCR) recognition of peptide-major histocompatibility complexes (pMHCs) is characterized by a highly conserved docking polarity. Whether this polarity is driven by recognition or signaling constraints remains unclear. Using "reversed-docking" TCRβ-variable (TRBV) 17+ TCRs from the naïve mouse CD8+ T cell repertoire that recognizes the H-2Db-NP366 epitope, we demonstrate that their inability to support T cell activation and in vivo recruitment is a direct consequence of reversed docking polarity and not TCR-pMHCI binding or clustering characteristics. Canonical TCR-pMHCI docking optimally localizes CD8/Lck to the CD3 complex, which is prevented by reversed TCR-pMHCI polarity. The requirement for canonical docking was circumvented by dissociating Lck from CD8. Thus, the consensus TCR-pMHC docking topology is mandated by T cell signaling constraints.
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Affiliation(s)
- Pirooz Zareie
- Infection and Immunity Program and Department of Biochemistry and Molecular Biology, Biomedicine Discovery Institute, Monash University, Clayton, Victoria, Australia
| | - Christopher Szeto
- Infection and Immunity Program and Department of Biochemistry and Molecular Biology, Biomedicine Discovery Institute, Monash University, Clayton, Victoria, Australia
| | - Carine Farenc
- Infection and Immunity Program and Department of Biochemistry and Molecular Biology, Biomedicine Discovery Institute, Monash University, Clayton, Victoria, Australia
| | - Sachith D Gunasinghe
- European Molecular Biology Laboratory (EMBL) Australia Node in Single Molecule Science and the ARC Centre of Excellence in Advanced Molecular Imaging, School of Medical Sciences, University of New South Wales, New South Wales, Australia
| | - Elizabeth M Kolawole
- Department of Pathology, University of Utah School of Medicine, Salt Lake City, UT, USA
| | - Angela Nguyen
- Infection and Immunity Program and Department of Biochemistry and Molecular Biology, Biomedicine Discovery Institute, Monash University, Clayton, Victoria, Australia
| | - Chantelle Blyth
- Infection and Immunity Program and Department of Biochemistry and Molecular Biology, Biomedicine Discovery Institute, Monash University, Clayton, Victoria, Australia
| | - Xavier Y X Sng
- Infection and Immunity Program and Department of Biochemistry and Molecular Biology, Biomedicine Discovery Institute, Monash University, Clayton, Victoria, Australia
| | - Jasmine Li
- Infection and Immunity Program and Department of Microbiology, Biomedicine Discovery Institute, Monash University, Clayton, Victoria, Australia
| | - Claerwen M Jones
- Infection and Immunity Program and Department of Biochemistry and Molecular Biology, Biomedicine Discovery Institute, Monash University, Clayton, Victoria, Australia
| | - Alex J Fulcher
- Monash Micro Imaging, Monash University, Clayton, Victoria, Australia
| | - Jesica R Jacobs
- Department of Pathology, University of Utah School of Medicine, Salt Lake City, UT, USA
| | - Qianru Wei
- Immunology Translational Research Programme and Department of Microbiology and Immunology, Yong Loo Lin School of Medicine, National University of Singapore, Singapore 117545
| | - Lukasz Wojciech
- Immunology Translational Research Programme and Department of Microbiology and Immunology, Yong Loo Lin School of Medicine, National University of Singapore, Singapore 117545
| | - Jan Petersen
- Infection and Immunity Program and Department of Biochemistry and Molecular Biology, Biomedicine Discovery Institute, Monash University, Clayton, Victoria, Australia.,Australian Research Council Centre of Excellence in Advanced Molecular Imaging, Monash University, Clayton, Victoria, Australia
| | - Nicholas R J Gascoigne
- Immunology Translational Research Programme and Department of Microbiology and Immunology, Yong Loo Lin School of Medicine, National University of Singapore, Singapore 117545
| | - Brian D Evavold
- Department of Pathology, University of Utah School of Medicine, Salt Lake City, UT, USA
| | - Katharina Gaus
- European Molecular Biology Laboratory (EMBL) Australia Node in Single Molecule Science and the ARC Centre of Excellence in Advanced Molecular Imaging, School of Medical Sciences, University of New South Wales, New South Wales, Australia
| | - Stephanie Gras
- Infection and Immunity Program and Department of Biochemistry and Molecular Biology, Biomedicine Discovery Institute, Monash University, Clayton, Victoria, Australia. .,Australian Research Council Centre of Excellence in Advanced Molecular Imaging, Monash University, Clayton, Victoria, Australia
| | - Jamie Rossjohn
- Infection and Immunity Program and Department of Biochemistry and Molecular Biology, Biomedicine Discovery Institute, Monash University, Clayton, Victoria, Australia. .,Australian Research Council Centre of Excellence in Advanced Molecular Imaging, Monash University, Clayton, Victoria, Australia.,Institute of Infection and Immunity, Cardiff University School of Medicine, Heath Park, Cardiff, UK
| | - Nicole L La Gruta
- Infection and Immunity Program and Department of Biochemistry and Molecular Biology, Biomedicine Discovery Institute, Monash University, Clayton, Victoria, Australia.
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34
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Miton CM, Buda K, Tokuriki N. Epistasis and intramolecular networks in protein evolution. Curr Opin Struct Biol 2021; 69:160-168. [PMID: 34077895 DOI: 10.1016/j.sbi.2021.04.007] [Citation(s) in RCA: 30] [Impact Index Per Article: 10.0] [Reference Citation Analysis] [Abstract] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/18/2021] [Revised: 04/01/2021] [Accepted: 04/21/2021] [Indexed: 12/01/2022]
Abstract
Proteins are molecular machines composed of complex, highly connected amino acid networks. Their functional optimization requires the reorganization of these intramolecular networks by evolution. In this review, we discuss the mechanisms by which epistasis, that is, the dependence of the effect of a mutation on the genetic background, rewires intramolecular interactions to alter protein function. Deciphering the biophysical basis of epistasis is crucial to our understanding of evolutionary dynamics and the elucidation of sequence-structure-function relationships. We featured recent studies that provide insights into the molecular mechanisms giving rise to epistasis, particularly at the structural level. These studies illustrate the convoluted and fascinating nature of the intramolecular networks co-opted by epistasis during the evolution of protein function.
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Affiliation(s)
- Charlotte M Miton
- Michael Smith Laboratories, University of British Columbia, Vancouver, V6T 1Z4, BC, Canada
| | - Karol Buda
- Michael Smith Laboratories, University of British Columbia, Vancouver, V6T 1Z4, BC, Canada
| | - Nobuhiko Tokuriki
- Michael Smith Laboratories, University of British Columbia, Vancouver, V6T 1Z4, BC, Canada.
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35
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Woo SO, Oh M, Alhalhooly L, Farmakes J, Rajapakse AJ, Yang Z, Collins PG, Choi Y. Different Single-Enzyme Conformational Dynamics upon Binding Hydrolyzable or Nonhydrolyzable Ligands. J Phys Chem B 2021; 125:5750-5756. [PMID: 34038124 DOI: 10.1021/acs.jpcb.1c01589] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/29/2022]
Abstract
Single-molecule measurements of protein dynamics help unveil the complex conformational changes and transitions that occur during ligand binding and catalytic processes. Using high-resolution single-molecule nanocircuit techniques, we have investigated differences in the conformational dynamics and transitions of lysozyme interacting with three ligands: peptidoglycan substrate, substrate-based chitin analogue, and indole derivative inhibitors. While processing peptidoglycan, lysozyme followed one of the two mechanistic pathways for the hydrolysis of the glycosidic bonds: a concerted mechanism inducing direct conformational changes from open to fully closed conformations or a nonconcerted mechanism involving transient pauses in intermediate conformations between the open and closed conformations. In the presence of either chitin or an indole inhibitor, lysozyme was unable to access the fully closed conformation where catalysis occurs. Instead, lysozymes' conformational closures terminated at slightly closed, "excited" conformations that were approximately one-quarter of the full hinge-bending range. With the indole inhibitor, lysozyme reached this excited conformation in a single step without any evidence of rate-liming intermediates, but the same conformational motions with chitin involved three hidden, intermediate processes and features similar to the nonconcerted peptidoglycan mechanism. The similarities suggest that these hidden processes involve attempts to accommodate imperfectly aligned polysaccharides in the active site. The results provide a detailed glimpse of the enzyme-ligand interplay at the crux of molecular recognition, enzyme specificity, and catalysis.
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Affiliation(s)
- Sung Oh Woo
- Department of Physics, North Dakota State University, Fargo, North Dakota 5810, United States
| | - Myungkeun Oh
- Materials and Nanotechnology Program, North Dakota State University, Fargo, North Dakota 58108, United States
| | - Lina Alhalhooly
- Department of Physics, North Dakota State University, Fargo, North Dakota 5810, United States
| | - Jasmin Farmakes
- Department of Chemistry and Biochemistry, North Dakota State University, Fargo, North Dakota 58108, United States
| | - Arith J Rajapakse
- Department of Physics and Astronomy, University of California, Irvine, California 92697, United States
| | - Zhongyu Yang
- Department of Chemistry and Biochemistry, North Dakota State University, Fargo, North Dakota 58108, United States
| | - Philip G Collins
- Department of Physics and Astronomy, University of California, Irvine, California 92697, United States
| | - Yongki Choi
- Department of Physics, North Dakota State University, Fargo, North Dakota 5810, United States.,Materials and Nanotechnology Program, North Dakota State University, Fargo, North Dakota 58108, United States
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36
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Ma J, Saikia N, Godar S, Hamilton GL, Ding F, Alper J, Sanabria H. Ensemble Switching Unveils a Kinetic Rheostat Mechanism of the Eukaryotic Thiamine Pyrophosphate Riboswitch. RNA (NEW YORK, N.Y.) 2021; 27:rna.075937.120. [PMID: 33863818 PMCID: PMC8208051 DOI: 10.1261/rna.075937.120] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 04/21/2020] [Accepted: 04/13/2021] [Indexed: 05/05/2023]
Abstract
Thiamine pyrophosphate (TPP) riboswitches regulate thiamine metabolism by inhibiting the translation of enzymes essential to thiamine synthesis pathways upon binding to thiamine pyrophosphate in cells across all domains of life. Recent work on the Arabidopsis thaliana TPP riboswitch suggests a multi-step TPP binding process involving multiple riboswitch configurational ensembles and that Mg2+ dependence underlies the mechanism of TPP recognition and subsequent transition to the expression-inhibiting state of the aptamer domain followed by changes in the expression platform. However, details of the relationship between TPP riboswitch conformational changes and interactions with TPP and Mg2+ ¬¬in the aptamer domain constituting this mechanism are unknown. Therefore, we integrated single-molecule multiparameter fluorescence and force spectroscopy with atomistic molecular dynamics simulations and found that conformational transitions within the aptamer domain's sensor helices associated with TPP and Mg2+ ligand binding occurred between at least five different ensembles on timescales ranging from µs to ms. These dynamics are orders of magnitude faster than the 10 second-timescale folding kinetics associated with expression-state switching in the switch sequence. Together, our results show that a TPP and Mg2+ dependent mechanism determines dynamic configurational state ensemble switching of the aptamer domain's sensor helices that regulates the stability of the switch helix, which ultimately may lead to the expression-inhibiting state of the riboswitch. Additionally, we propose that two pathways exist for ligand recognition and that this mechanism underlies a kinetic rheostat-like behavior of the Arabidopsis thaliana TPP riboswitch.
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Affiliation(s)
- Junyan Ma
- Department of Chemistry, Clemson University
| | | | - Subash Godar
- Department of Physics and Astronomy, Clemson University
| | | | - Feng Ding
- Department of Physics and Astronomy, Clemson University
| | - Joshua Alper
- Department of Physics and Astronomy, Clemson University
| | - Hugo Sanabria
- Department of Physics and Astronomy, Clemson University
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37
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Sanders JC, Holmstrom ED. Integrating single-molecule FRET and biomolecular simulations to study diverse interactions between nucleic acids and proteins. Essays Biochem 2021; 65:37-49. [PMID: 33600559 PMCID: PMC8052285 DOI: 10.1042/ebc20200022] [Citation(s) in RCA: 6] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/13/2020] [Revised: 01/17/2021] [Accepted: 01/26/2021] [Indexed: 12/12/2022]
Abstract
The conformations of biological macromolecules are intimately related to their cellular functions. Conveniently, the well-characterized dipole-dipole distance-dependence of Förster resonance energy transfer (FRET) makes it possible to measure and monitor the nanoscale spatial dimensions of these conformations using fluorescence spectroscopy. For this reason, FRET is often used in conjunction with single-molecule detection to study a wide range of conformationally dynamic biochemical processes. Written for those not yet familiar with the subject, this review aims to introduce biochemists to the methodology associated with single-molecule FRET, with a particular emphasis on how it can be combined with biomolecular simulations to study diverse interactions between nucleic acids and proteins. In the first section, we highlight several conceptual and practical considerations related to this integrative approach. In the second section, we review a few recent research efforts wherein various combinations of single-molecule FRET and biomolecular simulations were used to study the structural and dynamic properties of biochemical systems involving different types of nucleic acids (e.g., DNA and RNA) and proteins (e.g., folded and disordered).
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Affiliation(s)
- Joshua C Sanders
- Department of Chemistry, University of Kansas, Lawrence, KS, U.S.A
| | - Erik D Holmstrom
- Department of Chemistry, University of Kansas, Lawrence, KS, U.S.A
- Department of Molecular Biosciences, University of Kansas, Lawrence, KS, U.S.A
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38
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Wort JL, Ackermann K, Norman DG, Bode BE. A general model to optimise Cu II labelling efficiency of double-histidine motifs for pulse dipolar EPR applications. Phys Chem Chem Phys 2021; 23:3810-3819. [PMID: 33533341 DOI: 10.1039/d0cp06196d] [Citation(s) in RCA: 19] [Impact Index Per Article: 6.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/31/2022]
Abstract
Electron paramagnetic resonance (EPR) distance measurements are making increasingly important contributions to studies of biomolecules underpinning health and disease by providing highly accurate and precise geometric constraints. Combining double-histidine (dH) motifs with CuII spin labels shows promise for further increasing the precision of distance measurements, and for investigating subtle conformational changes. However, non-covalent coordination-based spin labelling is vulnerable to low binding affinity. Dissociation constants of dH motifs for CuII-nitrilotriacetic acid were previously investigated via relaxation induced dipolar modulation enhancement (RIDME), and demonstrated the feasibility of exploiting the dH motif for EPR applications at sub-μM protein concentrations. Herein, the feasibility of using modulation depth quantitation in CuII-CuII RIDME to simultaneously estimate a pair of non-identical independent KD values in such a tetra-histidine model protein is addressed. Furthermore, we develop a general speciation model to optimise CuII labelling efficiency, depending upon pairs of identical or disparate KD values and total CuII label concentration. We find the dissociation constant estimates are in excellent agreement with previously determined values, and empirical modulation depths support the proposed model.
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Affiliation(s)
- Joshua L Wort
- EaStCHEM School of Chemistry, Biomedical Sciences Research Complex, and Centre of Magnetic Resonance, University of St Andrews North Haugh, St Andrews KY16 9ST, UK.
| | - Katrin Ackermann
- EaStCHEM School of Chemistry, Biomedical Sciences Research Complex, and Centre of Magnetic Resonance, University of St Andrews North Haugh, St Andrews KY16 9ST, UK.
| | - David G Norman
- School of Life Sciences, University of Dundee, Medical Sciences Institute, Dundee, DD1 5EH, UK
| | - Bela E Bode
- EaStCHEM School of Chemistry, Biomedical Sciences Research Complex, and Centre of Magnetic Resonance, University of St Andrews North Haugh, St Andrews KY16 9ST, UK.
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39
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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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40
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Wort JL, Arya S, Ackermann K, Stewart AJ, Bode BE. Pulse Dipolar EPR Reveals Double-Histidine Motif Cu II-NTA Spin-Labeling Robustness against Competitor Ions. J Phys Chem Lett 2021. [PMID: 33715381 DOI: 10.17630/d7138874-55dd-4874-a2e8-c026fbc0b67f] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 05/09/2023]
Abstract
Pulse-dipolar EPR is an appealing strategy for structural characterization of complex systems in solution that complements other biophysical techniques. Significantly, the emergence of genetically encoded self-assembling spin labels exploiting exogenously introduced double-histidine motifs in conjunction with CuII-chelates offers high precision distance determination in systems nonpermissive to thiol-directed spin labeling. However, the noncovalency of this interaction exposes potential vulnerabilities to competition from adventitious divalent metal ions, and pH sensitivity. Herein, a combination of room-temperature isothermal titration calorimetry (ITC) and cryogenic relaxation-induced dipolar modulation enhancement (RIDME) measurements are applied to the model protein Streptococcus sp. group G. protein G, B1 domain (GB1). Results demonstrate double-histidine motif spin labeling using CuII-nitrilotriacetic acid (CuII-NTA) is robust against the competitor ligand ZnII-NTA at >1000-fold molar excess, and high nM binding affinity is surprisingly retained under acidic and basic conditions even though room temperature affinity shows a stronger pH dependence. This indicates the strategy is well-suited for diverse biological applications, with the requirement of other metal ion cofactors or slightly acidic pH not necessarily being prohibitive.
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Affiliation(s)
| | - Swati Arya
- School of Medicine, University of St. Andrews, North Haugh, St. Andrews, KY16 9TF, U.K
| | | | - Alan J Stewart
- School of Medicine, University of St. Andrews, North Haugh, St. Andrews, KY16 9TF, U.K
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41
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Wort J, Arya S, Ackermann K, Stewart AJ, Bode BE. Pulse Dipolar EPR Reveals Double-Histidine Motif Cu II-NTA Spin-Labeling Robustness against Competitor Ions. J Phys Chem Lett 2021; 12:2815-2819. [PMID: 33715381 PMCID: PMC8006131 DOI: 10.1021/acs.jpclett.1c00211] [Citation(s) in RCA: 25] [Impact Index Per Article: 8.3] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 05/09/2023]
Abstract
Pulse-dipolar EPR is an appealing strategy for structural characterization of complex systems in solution that complements other biophysical techniques. Significantly, the emergence of genetically encoded self-assembling spin labels exploiting exogenously introduced double-histidine motifs in conjunction with CuII-chelates offers high precision distance determination in systems nonpermissive to thiol-directed spin labeling. However, the noncovalency of this interaction exposes potential vulnerabilities to competition from adventitious divalent metal ions, and pH sensitivity. Herein, a combination of room-temperature isothermal titration calorimetry (ITC) and cryogenic relaxation-induced dipolar modulation enhancement (RIDME) measurements are applied to the model protein Streptococcus sp. group G. protein G, B1 domain (GB1). Results demonstrate double-histidine motif spin labeling using CuII-nitrilotriacetic acid (CuII-NTA) is robust against the competitor ligand ZnII-NTA at >1000-fold molar excess, and high nM binding affinity is surprisingly retained under acidic and basic conditions even though room temperature affinity shows a stronger pH dependence. This indicates the strategy is well-suited for diverse biological applications, with the requirement of other metal ion cofactors or slightly acidic pH not necessarily being prohibitive.
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Affiliation(s)
- Joshua
L. Wort
- EaStCHEM
School of Chemistry, Biomedical Sciences Research Complex, Centre
of Magnetic Resonance, University of St.
Andrews, North Haugh, St. Andrews, KY16 9ST, U.K.
| | - Swati Arya
- EaStCHEM
School of Chemistry, Biomedical Sciences Research Complex, Centre
of Magnetic Resonance, University of St.
Andrews, North Haugh, St. Andrews, KY16 9ST, U.K.
- School
of Medicine, University of St. Andrews, North Haugh, St. Andrews, KY16 9TF, U.K.
| | - Katrin Ackermann
- EaStCHEM
School of Chemistry, Biomedical Sciences Research Complex, Centre
of Magnetic Resonance, University of St.
Andrews, North Haugh, St. Andrews, KY16 9ST, U.K.
| | - Alan J. Stewart
- EaStCHEM
School of Chemistry, Biomedical Sciences Research Complex, Centre
of Magnetic Resonance, University of St.
Andrews, North Haugh, St. Andrews, KY16 9ST, U.K.
- School
of Medicine, University of St. Andrews, North Haugh, St. Andrews, KY16 9TF, U.K.
| | - Bela E. Bode
- EaStCHEM
School of Chemistry, Biomedical Sciences Research Complex, Centre
of Magnetic Resonance, University of St.
Andrews, North Haugh, St. Andrews, KY16 9ST, U.K.
- E-mail:
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42
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Kurisaki I, Tanaka S. Reaction Pathway Sampling and Free-Energy Analyses for Multimeric Protein Complex Disassembly by Employing Hybrid Configuration Bias Monte Carlo/Molecular Dynamics Simulation. ACS OMEGA 2021; 6:4749-4758. [PMID: 33644582 PMCID: PMC7905796 DOI: 10.1021/acsomega.0c05579] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.7] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Received: 11/16/2020] [Accepted: 01/27/2021] [Indexed: 05/08/2023]
Abstract
Physicochemical characterization of multimeric biomacromolecule assembly and disassembly processes is a milestone to understand the mechanisms for biological phenomena at the molecular level. Mass spectroscopy (MS) and structural bioinformatics (SB) approaches have become feasible to identify subcomplexes involved in assembly and disassembly, while they cannot provide atomic information sufficient for free-energy calculation to characterize transition mechanism between two different sets of subcomplexes. To combine observations derived from MS and SB approaches with conventional free-energy calculation protocols, we here designed a new reaction pathway sampling method by employing hybrid configuration bias Monte Carlo/molecular dynamics (hcbMC/MD) scheme and applied it to simulate the disassembly process of serum amyloid P component (SAP) pentamer. The results we obtained are consistent with those of the earlier MS and SB studies with respect to SAP subcomplex species and the initial stage of SAP disassembly processes. Furthermore, we observed a novel dissociation event, ring-opening reaction of SAP pentamer. Employing free-energy calculation combined with the hcbMC/MD reaction pathway trajectories, we moreover obtained experimentally testable observations on (1) reaction time of the ring-opening reaction and (2) importance of Asp42 and Lys117 for stable formation of SAP oligomer.
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43
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Medina E, R Latham D, Sanabria H. Unraveling protein's structural dynamics: from configurational dynamics to ensemble switching guides functional mesoscale assemblies. Curr Opin Struct Biol 2021; 66:129-138. [PMID: 33246199 PMCID: PMC7965259 DOI: 10.1016/j.sbi.2020.10.016] [Citation(s) in RCA: 13] [Impact Index Per Article: 4.3] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/19/2020] [Revised: 10/16/2020] [Accepted: 10/19/2020] [Indexed: 12/18/2022]
Abstract
Evidence regarding protein structure and function manifest the imperative role that dynamics play in proteins, underlining reconsideration of the unanimated sequence-to-structure-to-function paradigm. Structural dynamics portray a heterogeneous energy landscape described by conformational ensembles where each structural representation can be responsible for unique functions or enable macromolecular assemblies. Using the human p27/Cdk2/Cyclin A ternary complex as an example, we highlight the vital role of intramolecular and intermolecular dynamics for target recognition, binding, and inhibition as a critical modulator of cell division. Rapidly sampling configurations is critical for the population of different conformational ensembles encoding functional roles. To garner this knowledge, we present how the integration of (sub)ensemble and single-molecule fluorescence spectroscopy with molecular dynamic simulations can characterize structural dynamics linking the heterogeneous ensembles to function. The incorporation of dynamics into the sequence-to-structure-to-function paradigm promises to assist in tackling various challenges, including understanding the formation and regulation of mesoscale assemblies inside cells.
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Affiliation(s)
- Exequiel Medina
- Departamento de Biología, Facultad de Ciencias, Universidad de Chile, Las Palmeras 3425, Casilla 653, Santiago 7800003, Chile; Department of Physics and Astronomy, Clemson University, Clemson 29634, United States
| | - Danielle R Latham
- Department of Physics and Astronomy, Clemson University, Clemson 29634, United States
| | - Hugo Sanabria
- Department of Physics and Astronomy, Clemson University, Clemson 29634, United States.
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44
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Schepers B, Gohlke H. AMBER-DYES in AMBER: Implementation of fluorophore and linker parameters into AmberTools. J Chem Phys 2021; 152:221103. [PMID: 32534525 DOI: 10.1063/5.0007630] [Citation(s) in RCA: 13] [Impact Index Per Article: 4.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/23/2023] Open
Abstract
Molecular dynamics (MD) simulations of explicit representations of fluorescent dyes attached via a linker to a protein allow, e.g., probing commonly used approximations for dye localization and/or orientation or modeling Förster resonance energy transfer. However, setting up and performing such MD simulations with the AMBER suite of biomolecular simulation programs has remained challenging due to the unavailability of an easy-to-use set of parameters within AMBER. Here, we adapted the AMBER-DYES parameter set derived by Graen et al. [J. Chem. Theory Comput. 10, 5505 (2014)] into "AMBER-DYES in AMBER" to generate a force field applicable within AMBER for commonly used fluorescent dyes and linkers attached to a protein. In particular, the computationally efficient graphics processing unit (GPU) implementation of the AMBER MD engine can now be exploited to overcome sampling issues of dye movements. The implementation is compatible with state-of-the-art force fields such as GAFF, GAFF2, ff99SB, ff14SB, lipid17, and GLYCAM_06j, which allows simulating post-translationally modified proteins and/or protein-ligand complexes and/or proteins in membrane environments. It is applicable with frequently used water models such as TIP3P, TIP4P, TIP4P-Ew, and OPC. For ease of use, a LEaP-based workflow was created, which allows attaching (multiple) dye/linker combinations to a protein prior to further system preparation steps. Following the parameter development described by Graen et al. [J. Chem. Theory Comput. 10, 5505 (2014)] and the adaptation steps described here, AMBER-DYES in AMBER can be extended by additional linkers and fluorescent molecules.
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Affiliation(s)
- Bastian Schepers
- Institute for Pharmaceutical and Medicinal Chemistry, Heinrich Heine University Düsseldorf, 40225 Düsseldorf, Germany
| | - Holger Gohlke
- Institute for Pharmaceutical and Medicinal Chemistry, Heinrich Heine University Düsseldorf, 40225 Düsseldorf, Germany
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Chen H, Xie X, Chen TY. Single-molecule microscopy for in-cell quantification of protein oligomeric stoichiometry. Curr Opin Struct Biol 2020; 66:112-118. [PMID: 33242727 DOI: 10.1016/j.sbi.2020.10.022] [Citation(s) in RCA: 7] [Impact Index Per Article: 1.8] [Reference Citation Analysis] [Abstract] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/12/2020] [Revised: 10/14/2020] [Accepted: 10/19/2020] [Indexed: 12/11/2022]
Abstract
Protein organization modification plays a vital role in initiating signaling pathways, transcriptional regulation, and cell apoptosis regulation. Simultaneous quantification of oligomeric state and cellular parameters in the same cell, even though challenging, is required to understand their correlation at the molecular level. Recent advances of fluorescence protein and single-molecule localization microscopy enables the determination of localizations and oligomeric states of target proteins in cells. We reviewed the fluorescence intensity-based, localization-based, and photophysical property-based approaches for in-cell quantification of protein oligomeric stoichiometry. We discussed their working principles, applications, advantages, and limitations. These results also imply the combination of methodologies targeting different biological parameters at the single-cell level is essential to uncover the structure-function relationship at the molecular level.
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Affiliation(s)
- Huanhuan Chen
- Department of Chemistry, University of Houston, Houston, TX 77204, United States
| | - Xihong Xie
- Department of Chemistry, University of Houston, Houston, TX 77204, United States
| | - Tai-Yen Chen
- Department of Chemistry, University of Houston, Houston, TX 77204, United States.
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Dimura M, Peulen TO, Sanabria H, Rodnin D, Hemmen K, Hanke CA, Seidel CAM, Gohlke H. Automated and optimally FRET-assisted structural modeling. Nat Commun 2020; 11:5394. [PMID: 33106483 PMCID: PMC7589535 DOI: 10.1038/s41467-020-19023-1] [Citation(s) in RCA: 18] [Impact Index Per Article: 4.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/01/2020] [Accepted: 09/28/2020] [Indexed: 12/27/2022] Open
Abstract
FRET experiments can provide state-specific structural information of complex dynamic biomolecular assemblies. However, to overcome the sparsity of FRET experiments, they need to be combined with computer simulations. We introduce a program suite with (i) an automated design tool for FRET experiments, which determines how many and which FRET pairs should be used to minimize the uncertainty and maximize the accuracy of an integrative structure, (ii) an efficient approach for FRET-assisted coarse-grained structural modeling, and all-atom molecular dynamics simulations-based refinement, and (iii) a quantitative quality estimate for judging the accuracy of FRET-derived structures as opposed to precision. We benchmark our tools against simulated and experimental data of proteins with multiple conformational states and demonstrate an accuracy of ~3 Å RMSDCα against X-ray structures for sets of 15 to 23 FRET pairs. Free and open-source software for the introduced workflow is available at https://github.com/Fluorescence-Tools. A web server for FRET-assisted structural modeling of proteins is available at http://nmsim.de. To overcome the limitation of FRET data being too sparse to cover all structural details, FRET experiments need to be carefully designed and complemented with simulations. Here the authors present a toolkit for automated design of FRET experiments, which determines how many and which FRET pairs should be used to maximize the accuracy, and for FRET-assisted structural modeling and refinement at the atomistic level.
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Affiliation(s)
- Mykola Dimura
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany.,Institute for Pharmaceutical and Medicinal Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany
| | - Thomas-Otavio Peulen
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany
| | - Hugo Sanabria
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany.,Department of Physics and Astronomy, Clemson University, Clemson, SC, USA
| | - Dmitro Rodnin
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany
| | - Katherina Hemmen
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany
| | - Christian A Hanke
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany
| | - Claus A M Seidel
- Chair for Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany.
| | - Holger Gohlke
- Institute for Pharmaceutical and Medicinal Chemistry, Heinrich Heine University Düsseldorf, 40225, Düsseldorf, Germany. .,John von Neumann Institute for Computing (NIC), Jülich Supercomputing Centre (JSC), and Institute of Biological Information Processing (IBI-7: Structural Biochemistry), Forschungszentrum Jülich GmbH, 52425, Jülich, Germany.
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Cárdenas R, Martínez-Seoane J, Amero C. Combining Experimental Data and Computational Methods for the Non-Computer Specialist. Molecules 2020; 25:E4783. [PMID: 33081072 PMCID: PMC7594097 DOI: 10.3390/molecules25204783] [Citation(s) in RCA: 7] [Impact Index Per Article: 1.8] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/01/2020] [Revised: 08/25/2020] [Accepted: 08/28/2020] [Indexed: 01/01/2023] Open
Abstract
Experimental methods are indispensable for the study of the function of biological macromolecules, not just as static structures, but as dynamic systems that change conformation, bind partners, perform reactions, and respond to different stimulus. However, providing a detailed structural interpretation of the results is often a very challenging task. While experimental and computational methods are often considered as two different and separate approaches, the power and utility of combining both is undeniable. The integration of the experimental data with computational techniques can assist and enrich the interpretation, providing new detailed molecular understanding of the systems. Here, we briefly describe the basic principles of how experimental data can be combined with computational methods to obtain insights into the molecular mechanism and expand the interpretation through the generation of detailed models.
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Affiliation(s)
| | | | - Carlos Amero
- Laboratorio de Bioquímica y Resonancia Magnética Nuclear, Centro de Investigaciones Químicas, Instituto de Investigación en Ciencias Básicas y Aplicadas, Universidad Autónoma del Estado de Morelos, Cuernavaca, Morelos 62209, Mexico; (R.C.); (J.M.-S.)
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Haran G, Mazal H. How fast are the motions of tertiary-structure elements in proteins? J Chem Phys 2020; 153:130902. [PMID: 33032421 DOI: 10.1063/5.0024972] [Citation(s) in RCA: 10] [Impact Index Per Article: 2.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/18/2022] Open
Abstract
Protein motions occur on multiple time and distance scales. Large-scale motions of protein tertiary-structure elements, i.e., domains, are particularly intriguing as they are essential for the catalytic activity of many enzymes and for the functional cycles of protein machines and motors. Theoretical estimates suggest that domain motions should be very fast, occurring on the nanosecond or microsecond time scales. Indeed, free-energy barriers for domain motions are likely to involve salt bridges, which can break in microseconds. Experimental methods that can directly probe domain motions on fast time scales have appeared only in recent years. This Perspective discusses briefly some of these techniques, including nuclear magnetic resonance and single-molecule fluorescence spectroscopies. We introduce a few recent studies that demonstrate ultrafast domain motions and discuss their potential roles. Particularly surprising is the observation of tertiary-structure element dynamics that are much faster than the functional cycles in some protein machines. These swift motions can be rationalized on a case-by-case basis. For example, fast domain closure in multi-substrate enzymes may be utilized to optimize relative substrate orientation. Whether a large mismatch in time scales of conformational dynamics vs functional cycles is a general design principle in proteins remains to be determined.
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Affiliation(s)
- Gilad Haran
- Department of Chemical and Biological Physics, Weizmann Institute of Science, Rehovot, Israel
| | - Hisham Mazal
- Department of Chemical and Biological Physics, Weizmann Institute of Science, Rehovot, Israel
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Ciferri A. Molecular recognition mechanisms directing the self-assembly of biological structures. SOFT MATTER 2020; 16:8985-8995. [PMID: 33015695 DOI: 10.1039/d0sm01301c] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/11/2023]
Abstract
Self-assembling may be defined as the spontaneous association of material units into structures that are often capable of cyclic reorganization and functional behavior. Various molecular recognition processes stabilize assemblies of polymers and biological structures. The present article analyzes cases in which chemical, shape and other recognition mechanisms are individually or cooperatively operative. Simpler self-assembling theories reported in the literature are highlighted. Detailed processes for which chemical recognition is the prevailing, enthalpy-driven, process include the non-ideal component of miscibility, supramolecular polymerization, host-guest complexes and template polymerization. Also discussed are systems such as liquid crystalline closed polymers, ternary mesogenic systems and rigid crystalline polymers for which shape recognition is the prevailing entropy-driven process. Other recognition mechanisms include ion condensation effects, hydrophobic bonding and growth-coupled-to-orientation. Combinations of various recognition mechanisms are particularly evident in biological structures. Self-assembling mechanisms involved in the genesis of some biological systems can be scientifically identified, but much more needs to be known to describe the "engineered" assembling modes that support complex functional organs.
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Affiliation(s)
- Alberto Ciferri
- Chemistry Department, Duke University, DURHAM, NC 27708, USA.
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Krainer G, Hartmann A, Bogatyr V, Nielsen J, Schlierf M, Otzen DE. SDS-induced multi-stage unfolding of a small globular protein through different denatured states revealed by single-molecule fluorescence. Chem Sci 2020; 11:9141-9153. [PMID: 34123163 PMCID: PMC8163379 DOI: 10.1039/d0sc02100h] [Citation(s) in RCA: 11] [Impact Index Per Article: 2.8] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/14/2020] [Accepted: 08/07/2020] [Indexed: 12/03/2022] Open
Abstract
Ionic surfactants such as sodium dodecyl sulfate (SDS) unfold proteins in a much more diverse yet effective way than chemical denaturants such as guanidium chloride (GdmCl). But how these unfolding processes compare on a molecular level is poorly understood. Here, we address this question by scrutinising the unfolding pathway of the globular protein S6 in SDS and GdmCl with single-molecule Förster resonance energy transfer (smFRET) spectroscopy. We show that the unfolding mechanism in SDS is strikingly different and convoluted in comparison to denaturation in GdmCl. In contrast to the reversible two-state unfolding behaviour in GdmCl characterised by kinetics on the timescale of seconds, SDS demonstrated not one, but four distinct regimes of interactions with S6, dependent on the surfactant concentration. At ≤1 mM SDS, S6 and surfactant molecules form quasi-micelles on a minute timescale; at millimolar [SDS], the protein denatures through an unfolded/denatured ensemble of highly heterogeneous states on a multi-second timescale; at tens of millimolar of SDS, the protein unfolds into a micelle-packed conformation on the second timescale; and >50 mM SDS, the protein unfolds with millisecond timescale dynamics. We propose a detailed model for multi-stage unfolding of S6 in SDS, which involves at least three different types of denatured states with different level of compactness and dynamics and a continually changing landscape of interactions between protein and surfactant. Our results highlight the great potential of single-molecule fluorescence as a direct probe of nanoscale protein structure and dynamics in chemically complex surfactant environments.
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Affiliation(s)
- Georg Krainer
- B CUBE - Center for Molecular Bioengineering, TU Dresden Tatzberg 41 01307 Dresden Germany
| | - Andreas Hartmann
- B CUBE - Center for Molecular Bioengineering, TU Dresden Tatzberg 41 01307 Dresden Germany
| | - Vadim Bogatyr
- B CUBE - Center for Molecular Bioengineering, TU Dresden Tatzberg 41 01307 Dresden Germany
| | - Janni Nielsen
- Interdisciplinary Nanoscience Center (iNANO), Aarhus University Gustav Wieds Vej 14 8000 Aarhus Denmark
| | - Michael Schlierf
- B CUBE - Center for Molecular Bioengineering, TU Dresden Tatzberg 41 01307 Dresden Germany
- Cluster of Excellence Physics of Life, TU Dresden 01062 Dresden Germany
| | - Daniel E Otzen
- Interdisciplinary Nanoscience Center (iNANO), Aarhus University Gustav Wieds Vej 14 8000 Aarhus Denmark
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