1
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Steffen FD, Cunha RA, Sigel RKO, Börner R. FRET-guided modeling of nucleic acids. Nucleic Acids Res 2024; 52:e59. [PMID: 38869063 PMCID: PMC11260485 DOI: 10.1093/nar/gkae496] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/27/2023] [Accepted: 05/29/2024] [Indexed: 06/14/2024] Open
Abstract
The functional diversity of RNAs is encoded in their innate conformational heterogeneity. The combination of single-molecule spectroscopy and computational modeling offers new attractive opportunities to map structural transitions within nucleic acid ensembles. Here, we describe a framework to harmonize single-molecule Förster resonance energy transfer (FRET) measurements with molecular dynamics simulations and de novo structure prediction. Using either all-atom or implicit fluorophore modeling, we recreate FRET experiments in silico, visualize the underlying structural dynamics and quantify the reaction coordinates. Using multiple accessible-contact volumes as a post hoc scoring method for fragment assembly in Rosetta, we demonstrate that FRET can be used to filter a de novo RNA structure prediction ensemble by refuting models that are not compatible with in vitro FRET measurement. We benchmark our FRET-assisted modeling approach on double-labeled DNA strands and validate it against an intrinsically dynamic manganese(II)-binding riboswitch. We show that a FRET coordinate describing the assembly of a four-way junction allows our pipeline to recapitulate the global fold of the riboswitch displayed by the crystal structure. We conclude that computational fluorescence spectroscopy facilitates the interpretability of dynamic structural ensembles and improves the mechanistic understanding of nucleic acid interactions.
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Affiliation(s)
- Fabio D Steffen
- Department of Chemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Richard A Cunha
- Department of Chemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Roland K O Sigel
- Department of Chemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Richard Börner
- Department of Chemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
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2
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Pati AK, Kilic Z, Martin MI, Terry DS, Borgia A, Bar S, Jockusch S, Kiselev R, Altman RB, Blanchard SC. Recovering true FRET efficiencies from smFRET investigations requires triplet state mitigation. Nat Methods 2024; 21:1222-1230. [PMID: 38877317 PMCID: PMC11239528 DOI: 10.1038/s41592-024-02293-8] [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: 03/29/2023] [Accepted: 04/25/2024] [Indexed: 06/16/2024]
Abstract
Single-molecule fluorescence resonance energy transfer (smFRET) methods employed to quantify time-dependent compositional and conformational changes within biomolecules require elevated illumination intensities to recover robust photon emission streams from individual fluorophores. Here we show that outside the weak-excitation limit, and in regimes where fluorophores must undergo many rapid cycles of excitation and relaxation, non-fluorescing, excitation-induced triplet states with lifetimes orders of magnitude longer lived than photon-emitting singlet states degrade photon emission streams from both donor and acceptor fluorophores resulting in illumination-intensity-dependent changes in FRET efficiency. These changes are not commonly taken into consideration; therefore, robust strategies to suppress excited state accumulations are required to recover accurate and precise FRET efficiency, and thus distance, estimates. We propose both robust triplet state suppression and data correction strategies that enable the recovery of FRET efficiencies more closely approximating true values, thereby extending the spatial and temporal resolution of smFRET.
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Affiliation(s)
- Avik K Pati
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
- Department of Chemistry, Birla Institute of Technology and Science, Pilani, Rajasthan, India
| | - Zeliha Kilic
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
| | - Maxwell I Martin
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
- Department of Chemical Biology & Therapeutics, St. Jude Children's Research Hospital, Memphis, TN, USA
| | - Daniel S Terry
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
| | - Alessandro Borgia
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
| | - Sukanta Bar
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
- Department of Chemical Biology & Therapeutics, St. Jude Children's Research Hospital, Memphis, TN, USA
| | - Steffen Jockusch
- Center for Photochemical Sciences and Department of Chemistry, Bowling Green State University, Bowling Green, OH, USA
| | - Roman Kiselev
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
| | - Roger B Altman
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA
- Department of Chemical Biology & Therapeutics, St. Jude Children's Research Hospital, Memphis, TN, USA
| | - Scott C Blanchard
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, TN, USA.
- Department of Chemical Biology & Therapeutics, St. Jude Children's Research Hospital, Memphis, TN, USA.
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3
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Miller JJ, Mallimadugula UL, Zimmerman MI, Stuchell-Brereton MD, Soranno A, Bowman GR. Accounting for fast vs slow exchange in single molecule FRET experiments reveals hidden conformational states. BIORXIV : THE PREPRINT SERVER FOR BIOLOGY 2024:2024.06.03.597137. [PMID: 38895430 PMCID: PMC11185552 DOI: 10.1101/2024.06.03.597137] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/21/2024]
Abstract
Proteins are dynamic systems whose structural preferences determine their function. Unfortunately, building atomically detailed models of protein structural ensembles remains challenging, limiting our understanding of the relationships between sequence, structure, and function. Combining single molecule Förster resonance energy transfer (smFRET) experiments with molecular dynamics simulations could provide experimentally grounded, all-atom models of a protein's structural ensemble. However, agreement between the two techniques is often insufficient to achieve this goal. Here, we explore whether accounting for important experimental details like averaging across structures sampled during a given smFRET measurement is responsible for this apparent discrepancy. We present an approach to account for this time-averaging by leveraging the kinetic information available from Markov state models of a protein's dynamics. This allows us to accurately assess which timescales are averaged during an experiment. We find this approach significantly improves agreement between simulations and experiments in proteins with varying degrees of dynamics, including the well-ordered protein T4 lysozyme, the partially disordered protein apolipoprotein E (ApoE), and a disordered amyloid protein (Aβ40). We find evidence for hidden states that are not apparent in smFRET experiments because of time averaging with other structures, akin to states in fast exchange in NMR, and evaluate different force fields. Finally, we show how remaining discrepancies between computations and experiments can be used to guide additional simulations and build structural models for states that were previously unaccounted for. We expect our approach will enable combining simulations and experiments to understand the link between sequence, structure, and function in many settings.
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Affiliation(s)
- Justin J. Miller
- Departments of Biochemistry & Biophysics and Bioengineering, University of Pennsylvania, Philadelphia, PA 19104, United States
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, Missouri 63110, United States
| | - Upasana L. Mallimadugula
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, Missouri 63110, United States
| | - Maxwell I. Zimmerman
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, Missouri 63110, United States
| | - Melissa D. Stuchell-Brereton
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, Missouri 63110, United States
| | - Andrea Soranno
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, Missouri 63110, United States
| | - Gregory R. Bowman
- Departments of Biochemistry & Biophysics and Bioengineering, University of Pennsylvania, Philadelphia, PA 19104, United States
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, Missouri 63110, United States
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4
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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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5
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Montepietra D, Tesei G, Martins JM, Kunze MBA, Best RB, Lindorff-Larsen K. FRETpredict: a Python package for FRET efficiency predictions using rotamer libraries. Commun Biol 2024; 7:298. [PMID: 38461354 PMCID: PMC10925062 DOI: 10.1038/s42003-024-05910-6] [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: 06/14/2023] [Accepted: 02/12/2024] [Indexed: 03/11/2024] Open
Abstract
Förster resonance energy transfer (FRET) is a widely-used and versatile technique for the structural characterization of biomolecules. Here, we introduce FRETpredict, an easy-to-use Python software to predict FRET efficiencies from ensembles of protein conformations. FRETpredict uses a rotamer library approach to describe the FRET probes covalently bound to the protein. The software efficiently and flexibly operates on large conformational ensembles such as those generated by molecular dynamics simulations to facilitate the validation or refinement of molecular models and the interpretation of experimental data. We provide access to rotamer libraries for many commonly used dyes and linkers and describe a general methodology to generate new rotamer libraries for FRET probes. We demonstrate the performance and accuracy of the software for different types of systems: a rigid peptide (polyproline 11), an intrinsically disordered protein (ACTR), and three folded proteins (HiSiaP, SBD2, and MalE). FRETpredict is open source (GPLv3) and is available at github.com/KULL-Centre/FRETpredict and as a Python PyPI package at pypi.org/project/FRETpredict .
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Affiliation(s)
- Daniele Montepietra
- Department of Chemical, Life and Environmental Sustainability Sciences, University of Parma, Parma, 43125, Italy
- Istituto Nanoscienze - CNR-NANO, Center S3, via G. Campi 213/A, 41125, Modena, Italy
| | - Giulio Tesei
- Structural Biology and NMR Laboratory & the Linderstrøm-Lang Centre for Protein Science, Department of Biology, University of Copenhagen, Copenhagen, DK-2200, Denmark
| | - João M Martins
- Structural Biology and NMR Laboratory & the Linderstrøm-Lang Centre for Protein Science, Department of Biology, University of Copenhagen, Copenhagen, DK-2200, Denmark
| | - Micha B A Kunze
- Structural Biology and NMR Laboratory & the Linderstrøm-Lang Centre for Protein Science, Department of Biology, University of Copenhagen, Copenhagen, DK-2200, Denmark
| | - Robert B Best
- Laboratory of Chemical Physics, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, MD, 20892-0520, USA.
| | - Kresten Lindorff-Larsen
- Structural Biology and NMR Laboratory & the Linderstrøm-Lang Centre for Protein Science, Department of Biology, University of Copenhagen, Copenhagen, DK-2200, Denmark.
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6
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Waly SMA, Benniston AC, Harriman A. Deducing the conformational space for an octa-proline helix. Chem Sci 2024; 15:1657-1671. [PMID: 38303943 PMCID: PMC10829019 DOI: 10.1039/d3sc05287g] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/06/2023] [Accepted: 12/20/2023] [Indexed: 02/03/2024] Open
Abstract
A molecular dyad, PY-P8-PER, comprising a proline octamer sandwiched between pyrene and perylene terminals has been synthesized in order to address the dynamics of electronic energy transfer (EET) along the oligo-proline chain. A simple pyrene-based control compound equipped with a bis-proline attachment serves as a reference for spectroscopic studies. The N-H NMR signal at the terminal pyrene allows distinction between cis and trans amides and, although the crystal structure for the control has the trans conformation, temperature-dependent NMR studies provide clear evidence for trans/cis isomerisation in D6-DMSO. Polar solvents tend to stabilise the trans structure for the pyrene amide group, even for longer oligo-proline units. Circular dichroism shows that the proline spacer for PY-P8-PER exists mainly in the all-trans geometry in methanol. Preferential excitation of the pyrene chromophore is possible at wavelengths in the 320-350 nm range and, for the dyad, is followed by efficacious EET to the perylene emitter. The probability for intramolecular EET, obtained from analysis of steady-state spectroscopic data, is ca. 80-90% in solvents of disparate polarity. Comparison with the Förster critical distance suggests the terminals are ca. 18 Å apart. Time-resolved fluorescence spectroscopy, in conjunction with DFT calculations, indicates the dyad exists as a handful of conformers displaying a narrow range of EET rates. Optimisation of a distributive model allows accurate simulation of the EET dynamics in terms of reasonable structures based on isomerisation of certain amide groups.
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Affiliation(s)
- Sara M A Waly
- Molecular Photonics Laboratory, Bedson Building, School of Natural and Environmental Sciences, Newcastle University Newcastle upon Tyne NE1 7RU UK
| | - Andrew C Benniston
- Molecular Photonics Laboratory, Bedson Building, School of Natural and Environmental Sciences, Newcastle University Newcastle upon Tyne NE1 7RU UK
| | - Anthony Harriman
- Molecular Photonics Laboratory, Bedson Building, School of Natural and Environmental Sciences, Newcastle University Newcastle upon Tyne NE1 7RU UK
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7
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Mooren OL, Stuchell-Brereton MD, McConnell P, Yan C, Wilkerson EM, Goldfarb D, Cooper JA, Sept D, Soranno A. Biophysical Mechanism of Allosteric Regulation of Actin Capping Protein. J Mol Biol 2023; 435:168342. [PMID: 37924863 PMCID: PMC10872493 DOI: 10.1016/j.jmb.2023.168342] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/13/2023] [Revised: 10/28/2023] [Accepted: 10/30/2023] [Indexed: 11/06/2023]
Abstract
Actin capping protein (CP) can be regulated by steric and allosteric mechanisms. The molecular mechanism of the allosteric regulation at a biophysical level includes linkage between the binding sites for three ligands: F-actin, Capping-Protein-Interacting (CPI) motifs, and V-1/myotrophin, based on biochemical functional studies and solvent accessibility experiments. Here, we investigated the mechanism of allosteric regulation at the atomic level using single-molecule Förster resonance energy transfer (FRET) and molecular dynamics (MD) to assess the conformational and structural dynamics of CP in response to linked-binding site ligands. In the absence of ligand, both single-molecule FRET and MD revealed two distinct conformations of CP in solution; previous crystallographic studies revealed only one. Interaction with CPI-motif peptides induced conformations within CP that bring the cap and stalk closer, while interaction with V-1 moves them away from one another. Comparing CPI-motif peptides from different proteins, we identified variations in CP conformations and dynamics that are specific to each CPI motif. MD simulations for CP alone and in complex with a CPI motif and V-1 reveal atomistic details of the conformational changes. Analysis of the interaction of CP with wild-type (wt) and chimeric CPI-motif peptides using single-molecule FRET, isothermal calorimetry (ITC) and MD simulation indicated that conformational and affinity differences are intrinsic to the C-terminal portion of the CPI motif. We conclude that allosteric regulation of CP involves changes in conformation that disseminate across the protein to link distinct binding-site functions. Our results provide novel insights into the biophysical mechanism of the allosteric regulation of CP.
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Affiliation(s)
- Olivia L Mooren
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, United States
| | - Melissa D Stuchell-Brereton
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, United States; Center for Biomolecular Condensates, Washington University in St Louis, St. Louis, MO, United States
| | - Patrick McConnell
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, United States
| | - Chenbo Yan
- Department of Biophysics, University of Michigan, Ann Arbor, MI, United States
| | - Emily M Wilkerson
- Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, MO, United States; Institute for Informatics, Washington University School of Medicine, St. Louis, MO, United States
| | - Dennis Goldfarb
- Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, MO, United States; Institute for Informatics, Washington University School of Medicine, St. Louis, MO, United States
| | - John A Cooper
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, United States.
| | - David Sept
- Department of Biophysics, University of Michigan, Ann Arbor, MI, United States; Department of Biomedical Engineering, University of Michigan, Ann Arbor, MI, United States.
| | - Andrea Soranno
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, United States; Center for Biomolecular Condensates, Washington University in St Louis, St. Louis, MO, United States.
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8
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Mathur D, Díaz SA, Hildebrandt N, Pensack RD, Yurke B, Biaggne A, Li L, Melinger JS, Ancona MG, Knowlton WB, Medintz IL. Pursuing excitonic energy transfer with programmable DNA-based optical breadboards. Chem Soc Rev 2023; 52:7848-7948. [PMID: 37872857 PMCID: PMC10642627 DOI: 10.1039/d0cs00936a] [Citation(s) in RCA: 5] [Impact Index Per Article: 5.0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/15/2023] [Indexed: 10/25/2023]
Abstract
DNA nanotechnology has now enabled the self-assembly of almost any prescribed 3-dimensional nanoscale structure in large numbers and with high fidelity. These structures are also amenable to site-specific modification with a variety of small molecules ranging from drugs to reporter dyes. Beyond obvious application in biotechnology, such DNA structures are being pursued as programmable nanoscale optical breadboards where multiple different/identical fluorophores can be positioned with sub-nanometer resolution in a manner designed to allow them to engage in multistep excitonic energy-transfer (ET) via Förster resonance energy transfer (FRET) or other related processes. Not only is the ability to create such complex optical structures unique, more importantly, the ability to rapidly redesign and prototype almost all structural and optical analogues in a massively parallel format allows for deep insight into the underlying photophysical processes. Dynamic DNA structures further provide the unparalleled capability to reconfigure a DNA scaffold on the fly in situ and thus switch between ET pathways within a given assembly, actively change its properties, and even repeatedly toggle between two states such as on/off. Here, we review progress in developing these composite materials for potential applications that include artificial light harvesting, smart sensors, nanoactuators, optical barcoding, bioprobes, cryptography, computing, charge conversion, and theranostics to even new forms of optical data storage. Along with an introduction into the DNA scaffolding itself, the diverse fluorophores utilized in these structures, their incorporation chemistry, and the photophysical processes they are designed to exploit, we highlight the evolution of DNA architectures implemented in the pursuit of increased transfer efficiency and the key lessons about ET learned from each iteration. We also focus on recent and growing efforts to exploit DNA as a scaffold for assembling molecular dye aggregates that host delocalized excitons as a test bed for creating excitonic circuits and accessing other quantum-like optical phenomena. We conclude with an outlook on what is still required to transition these materials from a research pursuit to application specific prototypes and beyond.
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Affiliation(s)
- Divita Mathur
- Department of Chemistry, Case Western Reserve University, Cleveland OH 44106, USA
| | - Sebastián A Díaz
- Center for Bio/Molecular Science and Engineering, Code 6900, USA.
| | - Niko Hildebrandt
- Department of Chemistry, Seoul National University, Seoul 08826, South Korea
- Department of Engineering Physics, McMaster University, Hamilton, L8S 4L7, Canada
| | - Ryan D Pensack
- Micron School of Materials Science & Engineering, Boise State University, Boise, ID 83725, USA.
| | - Bernard Yurke
- Micron School of Materials Science & Engineering, Boise State University, Boise, ID 83725, USA.
| | - Austin Biaggne
- Micron School of Materials Science & Engineering, Boise State University, Boise, ID 83725, USA.
| | - Lan Li
- Micron School of Materials Science & Engineering, Boise State University, Boise, ID 83725, USA.
- Center for Advanced Energy Studies, Idaho Falls, ID 83401, USA
| | - Joseph S Melinger
- Electronics Science and Technology Division, Code 6800, U.S. Naval Research Laboratory, Washington, DC 20375, USA
| | - Mario G Ancona
- Electronics Science and Technology Division, Code 6800, U.S. Naval Research Laboratory, Washington, DC 20375, USA
- Department of Electrical and Computer Engineering, Florida State University, Tallahassee, FL 32310, USA
| | - William B Knowlton
- Micron School of Materials Science & Engineering, Boise State University, Boise, ID 83725, USA.
| | - Igor L Medintz
- Center for Bio/Molecular Science and Engineering, Code 6900, USA.
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9
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Tsangaris TE, Smyth S, Gomes GNW, Liu ZH, Milchberg M, Bah A, Wasney GA, Forman-Kay JD, Gradinaru CC. Delineating Structural Propensities of the 4E-BP2 Protein via Integrative Modeling and Clustering. J Phys Chem B 2023; 127:7472-7486. [PMID: 37595014 PMCID: PMC10858721 DOI: 10.1021/acs.jpcb.3c04052] [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: 08/20/2023]
Abstract
The intrinsically disordered 4E-BP2 protein regulates mRNA cap-dependent translation through interaction with the predominantly folded eukaryotic initiation factor 4E (eIF4E). Phosphorylation of 4E-BP2 dramatically reduces the level of eIF4E binding, in part by stabilizing a binding-incompatible folded domain. Here, we used a Rosetta-based sampling algorithm optimized for IDRs to generate initial ensembles for two phospho forms of 4E-BP2, non- and 5-fold phosphorylated (NP and 5P, respectively), with the 5P folded domain flanked by N- and C-terminal IDRs (N-IDR and C-IDR, respectively). We then applied an integrative Bayesian approach to obtain NP and 5P conformational ensembles that agree with experimental data from nuclear magnetic resonance, small-angle X-ray scattering, and single-molecule Förster resonance energy transfer (smFRET). For the NP state, inter-residue distance scaling and 2D maps revealed the role of charge segregation and pi interactions in driving contacts between distal regions of the chain (∼70 residues apart). The 5P ensemble shows prominent contacts of the N-IDR region with the two phosphosites in the folded domain, pT37 and pT46, and, to a lesser extent, delocalized interactions with the C-IDR region. Agglomerative hierarchical clustering led to partitioning of each of the two ensembles into four clusters with different global dimensions and contact maps. This helped delineate an NP cluster that, based on our smFRET data, is compatible with the eIF4E-bound state. 5P clusters were differentiated by interactions of C-IDR with the folded domain and of the N-IDR with the two phosphosites in the folded domain. Our study provides both a better visualization of fundamental structural poses of 4E-BP2 and a set of falsifiable insights on intrachain interactions that bias folding and binding of this protein.
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Affiliation(s)
- Thomas E Tsangaris
- Department of Physics, University of Toronto, Toronto, Ontario M5S 1A7, Canada
- Department of Chemical & Physical Sciences, University of Toronto Mississauga, Mississauga, Ontario L5L 1C6, Canada
| | - Spencer Smyth
- Department of Physics, University of Toronto, Toronto, Ontario M5S 1A7, Canada
- Department of Chemical & Physical Sciences, University of Toronto Mississauga, Mississauga, Ontario L5L 1C6, Canada
| | - Gregory-Neal W Gomes
- Department of Physics, University of Toronto, Toronto, Ontario M5S 1A7, Canada
- Department of Chemical & Physical Sciences, University of Toronto Mississauga, Mississauga, Ontario L5L 1C6, Canada
| | - Zi Hao Liu
- Program in Molecular Medicine, Hospital for Sick Children, Toronto, Ontario M5G 0A4, Canada
- Department of Biochemistry, University of Toronto, Toronto, Ontario M5S 1A8, Canada
| | - Moses Milchberg
- Program in Molecular Medicine, Hospital for Sick Children, Toronto, Ontario M5G 0A4, Canada
- Department of Biochemistry, University of Toronto, Toronto, Ontario M5S 1A8, Canada
| | - Alaji Bah
- Program in Molecular Medicine, Hospital for Sick Children, Toronto, Ontario M5G 0A4, Canada
- Department of Biochemistry, University of Toronto, Toronto, Ontario M5S 1A8, Canada
| | - Gregory A Wasney
- Peter Gilgan Centre for Research and Learning, Hospital for Sick Children, Toronto, Ontario M5G 0A4, Canada
| | - Julie D Forman-Kay
- Program in Molecular Medicine, Hospital for Sick Children, Toronto, Ontario M5G 0A4, Canada
- Department of Biochemistry, University of Toronto, Toronto, Ontario M5S 1A8, Canada
| | - Claudiu C Gradinaru
- Department of Physics, University of Toronto, Toronto, Ontario M5S 1A7, Canada
- Department of Chemical & Physical Sciences, University of Toronto Mississauga, Mississauga, Ontario L5L 1C6, Canada
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10
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Mooren OL, Stuchell-Brereton MD, McConnell P, Yan C, Wilkerson EM, Goldfarb D, Cooper JA, Sept D, Soranno A. Biophysical Mechanism of Allosteric Regulation of Actin Capping Protein. BIORXIV : THE PREPRINT SERVER FOR BIOLOGY 2023:2023.08.16.553570. [PMID: 37645735 PMCID: PMC10462145 DOI: 10.1101/2023.08.16.553570] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 08/31/2023]
Abstract
Actin capping protein (CP) can be regulated by steric and allosteric mechanisms. The molecular mechanism of the allosteric regulation at a biophysical level includes linkage between the binding sites for three ligands: F-actin, Capping-Protein-Interacting (CPI) motifs, and V-1/myotrophin, based on biochemical functional studies and solvent accessibility experiments. Here, we investigated the mechanism of allosteric regulation at the atomic level using single-molecule Förster resonance energy transfer (FRET) and molecular dynamics (MD) to assess the conformational and structural dynamics of CP in response to linked-binding site ligands. In the absence of ligand, both single-molecule FRET and MD revealed two distinct conformations of CP in solution; previous crystallographic studies revealed only one. CPI-motif peptide association induced conformational changes within CP that propagate in one direction, while V-1 association induced conformational changes in the opposite direction. Comparing CPI-motif peptides from different proteins, we identified variations in CP conformations and dynamics that are specific to each CPI motif. MD simulations for CP alone and in complex with a CPI motif and V-1 reveal atomistic details of the conformational changes. Analysis of the interaction of CP with wildtype (wt) and chimeric CPI-motif peptides using single-molecule FRET, isothermal calorimetry (ITC) and MD simulation indicated that conformational and affinity differences are intrinsic to the C-terminal portion of the CPI-motif. We conclude that allosteric regulation of CP involves changes in conformation that disseminate across the protein to link distinct binding-site functions. Our results provide novel insights into the biophysical mechanism of the allosteric regulation of CP.
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11
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Yang S, Zhang S, Hu F, Han J, Li F. Circularly polarized luminescence polymers: From design to applications. Coord Chem Rev 2023. [DOI: 10.1016/j.ccr.2023.215116] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 03/30/2023]
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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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Tam LKB, Chu JCH, He L, Yang C, Han KC, Cheung PCK, Ng DKP, Lo PC. Enzyme-Responsive Double-Locked Photodynamic Molecular Beacon for Targeted Photodynamic Anticancer Therapy. J Am Chem Soc 2023; 145:7361-7375. [PMID: 36961946 PMCID: PMC10080691 DOI: 10.1021/jacs.2c13732] [Citation(s) in RCA: 21] [Impact Index Per Article: 21.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 03/26/2023]
Abstract
An advanced photodynamic molecular beacon (PMB) was designed and synthesized, in which a distyryl boron dipyrromethene (DSBDP)-based photosensitizer and a Black Hole Quencher 3 moiety were connected via two peptide segments containing the sequences PLGVR and GFLG, respectively, of a cyclic peptide. These two short peptide sequences are well-known substrates of matrix metalloproteinase-2 (MMP-2) and cathepsin B, respectively, both of which are overexpressed in a wide range of cancer cells either extracellularly (for MMP-2) or intracellularly (for cathepsin B). Owing to the efficient Förster resonance energy transfer between the two components, this PMB was fully quenched in the native form. Only upon interaction with both MMP-2 and cathepsin B, either in a buffer solution or in cancer cells, both of the segments were cleaved specifically, and the two components could be completely separated, thereby restoring the photodynamic activities of the DSBDP moiety. This PMB could also be activated in tumors, and it effectively suppressed the tumor growth in A549 tumor-bearing nude mice upon laser irradiation without causing notable side effects. In particular, it did not cause skin photosensitivity, which is a very common side effect of photodynamic therapy (PDT) using conventional "always-on" photosensitizers. The overall results showed that this "double-locked" PMB functioned as a biological AND logic gate that could only be unlocked by the coexistence of two tumor-associated enzymes, which could greatly enhance the tumor specificity in PDT.
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Affiliation(s)
- Leo K B Tam
- Department of Chemistry, The Chinese University of Hong Kong, Shatin, N.T., Hong Kong, China
| | - Jacky C H Chu
- Department of Chemistry, The Chinese University of Hong Kong, Shatin, N.T., Hong Kong, China
| | - Lin He
- Department of Biomedical Sciences and Tung Biomedical Sciences Centre, City University of Hong Kong, Tat Chee Avenue, Kowloon, Hong Kong, China
| | - Caixia Yang
- Department of Chemistry, The Chinese University of Hong Kong, Shatin, N.T., Hong Kong, China
| | - Kam-Chu Han
- Department of Clinical Pathology, Pamela Youde Nethersole Eastern Hospital, Chai Wan, Hong Kong, China
| | - Peter Chi Keung Cheung
- School of Life Sciences, The Chinese University of Hong Kong, Shatin, N.T., Hong Kong, China
| | - Dennis K P Ng
- Department of Chemistry, The Chinese University of Hong Kong, Shatin, N.T., Hong Kong, China
| | - Pui-Chi Lo
- Department of Biomedical Sciences and Tung Biomedical Sciences Centre, City University of Hong Kong, Tat Chee Avenue, Kowloon, Hong Kong, China
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14
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Montepietra D, Tesei G, Martins JM, Kunze MBA, Best RB, Lindorff-Larsen K. FRETpredict: A Python package for FRET efficiency predictions using rotamer libraries. BIORXIV : THE PREPRINT SERVER FOR BIOLOGY 2023:2023.01.27.525885. [PMID: 36789411 PMCID: PMC9928041 DOI: 10.1101/2023.01.27.525885] [Citation(s) in RCA: 3] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Figures] [Subscribe] [Scholar Register] [Indexed: 01/30/2023]
Abstract
Here, we introduce FRETpredict, a Python software program to predict FRET efficiencies from ensembles of protein conformations. FRETpredict uses an established Rotamer Library Approach to describe the FRET probes covalently bound to the protein. The software efficiently operates on large conformational ensembles such as those generated by molecular dynamics simulations to facilitate the validation or refinement of molecular models and the interpretation of experimental data. We demonstrate the performance and accuracy of the software for different types of systems: a relatively structured peptide (polyproline 11), an intrinsically disordered protein (ACTR), and three folded proteins (HiSiaP, SBD2, and MalE). We also describe a general approach to generate new rotamer libraries for FRET probes of interest. FRETpredict is open source (GPLv3) and is available at github.com/KULL-Centre/FRETpredict and as a Python PyPI package at pypi.org/project/FRETpredict.
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Affiliation(s)
- Daniele Montepietra
- Department of Physics, Computer Science and Mathematics, University of Modena and Reggio Emilia, Via Campi 213/A 41125 Modena, Italy
- Istituto Nanoscienze – CNR-NANO, Center S3, via G. Campi 213/A, 41125 Modena, Italy
| | - Giulio Tesei
- Structural Biology and NMR Laboratory & the Linderstrøm-Lang Centre for Protein Science, Department of Biology, University of Copenhagen, Copenhagen, Denmark
| | - João M. Martins
- Structural Biology and NMR Laboratory & the Linderstrøm-Lang Centre for Protein Science, Department of Biology, University of Copenhagen, Copenhagen, Denmark
| | - Micha B. A. Kunze
- Structural Biology and NMR Laboratory & the Linderstrøm-Lang Centre for Protein Science, Department of Biology, University of Copenhagen, Copenhagen, Denmark
| | - Robert B. Best
- Laboratory of Chemical Physics, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, Maryland, United States of America
| | - Kresten Lindorff-Larsen
- Structural Biology and NMR Laboratory & the Linderstrøm-Lang Centre for Protein Science, Department of Biology, University of Copenhagen, Copenhagen, Denmark
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15
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Fujimoto KJ, Miyashita T, Dewa T, Yanai T. Determination of FRET orientation factor between artificial fluorophore and photosynthetic light-harvesting 2 complex (LH2). Sci Rep 2022; 12:15091. [PMID: 36065053 PMCID: PMC9445053 DOI: 10.1038/s41598-022-19375-2] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/23/2022] [Accepted: 08/29/2022] [Indexed: 11/30/2022] Open
Abstract
The orientation factor of fluorescence resonance energy transfer (FRET) between photosynthetic light-harvesting 2 complex (LH2) and artificial fluorophore (Alexa Fluor 647: A647) was theoretically investigated. The orientation factor of 2/3, i.e., the isotropic mean, is widely used to predict the donor–acceptor distance from FRET measurements. However, this approximation seems inappropriate because the movement of A647 is possibly restricted by the bifunctional linker binding to LH2. In this study, we performed molecular dynamics (MD) simulations and electronic coupling calculations on the LH2-A647 conjugate to analyze its orientation factor. The MD results showed that A647 keeps a position approximately 26 Å away from the bacteriochlorophyll (BChl) assembly in LH2. The effective orientation factor was extracted from the electronic coupling calculated using the transition charge from electrostatic potential (TrESP) method. With MD snapshots, an averaged orientation factor was predicted to be 1.55, significantly different from the isotropic mean value. The analysis also suggested that the value of the refractive index employed in the previous studies is not suitable for this system. Furthermore, optimal orientations of A647 with larger orientation factors to improve FRET efficiency were searched using Euler angles. The present approach is useful for extending the applicability of FRET analysis.
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Affiliation(s)
- Kazuhiro J Fujimoto
- Institute of Transformative Bio-Molecules (WPI-ITbM), Nagoya University, Furocho, Chikusa, Nagoya, 464-8601, Japan. .,Department of Chemistry, Graduate School of Science, Nagoya University, Furocho, Chikusa, Nagoya, 464-8601, Japan.
| | - Tomoya Miyashita
- Department of Chemistry, Graduate School of Science, Nagoya University, Furocho, Chikusa, Nagoya, 464-8601, Japan
| | - Takehisa Dewa
- Department of Life Science and Applied Chemistry, Graduate School of Engineering, Nagoya Institute of Technology, Gokiso-cho, Showa, Nagoya, 466-8555, Japan
| | - Takeshi Yanai
- Institute of Transformative Bio-Molecules (WPI-ITbM), Nagoya University, Furocho, Chikusa, Nagoya, 464-8601, Japan. .,Department of Chemistry, Graduate School of Science, Nagoya University, Furocho, Chikusa, Nagoya, 464-8601, Japan.
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16
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Zhang B, Lyu G, Kelly EA, Evans RC. Förster Resonance Energy Transfer in Luminescent Solar Concentrators. ADVANCED SCIENCE (WEINHEIM, BADEN-WURTTEMBERG, GERMANY) 2022; 9:e2201160. [PMID: 35678107 PMCID: PMC9376834 DOI: 10.1002/advs.202201160] [Citation(s) in RCA: 6] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 03/07/2022] [Revised: 04/27/2022] [Indexed: 05/20/2023]
Abstract
Luminescent solar concentrators (LSCs) are an emerging technology to collect and channel light from a large absorption area into a smaller one. They are a complementary technology for traditional solar photovoltaics (PV), particularly suitable for application in urban or indoor environments where their custom colors and form factors, and performance under diffuse light conditions may be advantageous. Förster resonance energy transfer (FRET) has emerged as a valuable approach to overcome some of the intrinsic limitations of conventional single lumophore LSCs, such as reabsorption or reduced quantum efficiency. This review outlines the potential of FRET to boost LSC performance, using highlights from the literature to illustrate the key criteria that must be considered when designing an FRET-LSC, including both the photophysical requirements of the FRET lumophores and their interaction with the host material. Based on these criteria, a list of design guidelines intended to aid researchers when they approach the design of a new FRET-LSC system is presented. By highlighting the unanswered questions in this field, the authors aim to demonstrate the potential of FRET-LSCs for both conventional solar-harvesting and emerging LSC-inspired technologies and hope to encourage participation from a diverse researcher base to address this exciting challenge.
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Affiliation(s)
- Bolong Zhang
- Department of Materials Science and MetallurgyUniversity of Cambridge27 Charles Babbage RoadCambridgeCB3 0FSUK
- CAS Key Laboratory of Design and Assembly of Functional Nanostructures, and Fujian Provincial Key Laboratory of Nanomaterials, Fujian Institute of Research on the Structure of MaterialsChinese Academy of SciencesFuzhouFujian350002China
| | - Guanpeng Lyu
- Department of Materials Science and MetallurgyUniversity of Cambridge27 Charles Babbage RoadCambridgeCB3 0FSUK
| | - Elaine A. Kelly
- Department of Materials Science and MetallurgyUniversity of Cambridge27 Charles Babbage RoadCambridgeCB3 0FSUK
| | - Rachel C. Evans
- Department of Materials Science and MetallurgyUniversity of Cambridge27 Charles Babbage RoadCambridgeCB3 0FSUK
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17
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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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18
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Cervantes-Salguero K, Biaggne A, Youngsman JM, Ward BM, Kim YC, Li L, Hall JA, Knowlton WB, Graugnard E, Kuang W. Strategies for Controlling the Spatial Orientation of Single Molecules Tethered on DNA Origami Templates Physisorbed on Glass Substrates: Intercalation and Stretching. Int J Mol Sci 2022; 23:ijms23147690. [PMID: 35887059 PMCID: PMC9323263 DOI: 10.3390/ijms23147690] [Citation(s) in RCA: 11] [Impact Index Per Article: 5.5] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/16/2022] [Revised: 07/08/2022] [Accepted: 07/10/2022] [Indexed: 11/18/2022] Open
Abstract
Nanoarchitectural control of matter is crucial for next-generation technologies. DNA origami templates are harnessed to accurately position single molecules; however, direct single molecule evidence is lacking regarding how well DNA origami can control the orientation of such molecules in three-dimensional space, as well as the factors affecting control. Here, we present two strategies for controlling the polar (θ) and in-plane azimuthal (ϕ) angular orientations of cyanine Cy5 single molecules tethered on rationally-designed DNA origami templates that are physically adsorbed (physisorbed) on glass substrates. By using dipolar imaging to evaluate Cy5′s orientation and super-resolution microscopy, the absolute spatial orientation of Cy5 is calculated relative to the DNA template. The sequence-dependent partial intercalation of Cy5 is discovered and supported theoretically using density functional theory and molecular dynamics simulations, and it is harnessed as our first strategy to achieve θ control for a full revolution with dispersion as small as ±4.5°. In our second strategy, ϕ control is achieved by mechanically stretching the Cy5 from its two tethers, being the dispersion ±10.3° for full stretching. These results can in principle be applied to any single molecule, expanding in this way the capabilities of DNA as a functional templating material for single-molecule orientation control. The experimental and modeling insights provided herein will help engineer similar self-assembling molecular systems based on polymers, such as RNA and proteins.
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Affiliation(s)
- Keitel Cervantes-Salguero
- Micron School of Materials Science and Engineering, Boise State University, Boise, ID 83725, USA; (A.B.); (J.M.Y.); (B.M.W.); (L.L.); (W.B.K.); (E.G.)
- Correspondence: (K.C.-S.); (W.K.)
| | - Austin Biaggne
- Micron School of Materials Science and Engineering, Boise State University, Boise, ID 83725, USA; (A.B.); (J.M.Y.); (B.M.W.); (L.L.); (W.B.K.); (E.G.)
| | - John M. Youngsman
- Micron School of Materials Science and Engineering, Boise State University, Boise, ID 83725, USA; (A.B.); (J.M.Y.); (B.M.W.); (L.L.); (W.B.K.); (E.G.)
| | - Brett M. Ward
- Micron School of Materials Science and Engineering, Boise State University, Boise, ID 83725, USA; (A.B.); (J.M.Y.); (B.M.W.); (L.L.); (W.B.K.); (E.G.)
| | - Young C. Kim
- Materials Science and Technology Division, U.S. Naval Research Laboratory, Code 6300, Washington, DC 20375, USA;
| | - Lan Li
- Micron School of Materials Science and Engineering, Boise State University, Boise, ID 83725, USA; (A.B.); (J.M.Y.); (B.M.W.); (L.L.); (W.B.K.); (E.G.)
- Center for Advanced Energy Studies, Idaho Falls, ID 83401, USA
| | - John A. Hall
- Division of Research and Economic Development, Boise State University, Boise, ID 83725, USA;
| | - William B. Knowlton
- Micron School of Materials Science and Engineering, Boise State University, Boise, ID 83725, USA; (A.B.); (J.M.Y.); (B.M.W.); (L.L.); (W.B.K.); (E.G.)
- Department of Electrical and Computer Engineering, Boise State University, Boise, ID 83725, USA
| | - Elton Graugnard
- Micron School of Materials Science and Engineering, Boise State University, Boise, ID 83725, USA; (A.B.); (J.M.Y.); (B.M.W.); (L.L.); (W.B.K.); (E.G.)
- Center for Advanced Energy Studies, Idaho Falls, ID 83401, USA
| | - Wan Kuang
- Department of Electrical and Computer Engineering, Boise State University, Boise, ID 83725, USA
- Correspondence: (K.C.-S.); (W.K.)
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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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20
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Wang D, Wu S, Wang D, Song X, Yang M, Zhang W, Huang S, Weng J, Liu Z, Wang W. The importance of the compact disordered state in the fuzzy interactions between intrinsically disordered proteins. Chem Sci 2022; 13:2363-2377. [PMID: 35310482 PMCID: PMC8864705 DOI: 10.1039/d1sc06825c] [Citation(s) in RCA: 7] [Impact Index Per Article: 3.5] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/07/2021] [Accepted: 01/21/2022] [Indexed: 11/23/2022] Open
Abstract
The intrinsically disordered C-terminal domain (CTD) of protein 4.1G is able to specifically bind a 26-residue intrinsically disordered region of NuMA, forming a dynamic fuzzy complex. As one of a few cases of extremely fuzzy interactions between two intrinsically disordered proteins/regions (IDPs/IDRs) without induced folding, the principle of the binding is unknown. Here, we combined experimental and computational methods to explore the detailed mechanism of the interaction between 4.1G-CTD and NuMA. MD simulations suggest that the kinetic hub states in the structure ensemble of 4.1G-CTD are favorable in the fuzzy complex. The feature of these hub states is that the binding 'hot spot' motifs βA and βB exhibit β strand propensities and are well packed to each other. The binding between 4.1G-CTD and NuMA is disrupted at low pH, which changes the intramolecular packing of 4.1G-CTD and weakens the packing between βA and βB motifs. Low pH conditions also lead to increased hydrodynamic radius and acceleration of backbone dynamics of 4.1G-CTD. All these results underscore the importance of tertiary structural arrangements and overall compactness of 4.1G-CTD in its binding to NuMA, i.e. the compact disordered state of 4.1G-CTD is crucial for binding. Different from the short linear motifs (SLiMs) that are often found to mediate IDP interactions, 4.1G-CTD functions as an intrinsically disordered domain (IDD), which is a functional and structural unit similar to conventional protein domains. This work sheds light on the molecular recognition mechanism of IDPs/IDRs and expands the conventional structure-function paradigm in protein biochemistry.
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Affiliation(s)
- Dan Wang
- Department of Chemistry, Multiscale Research Institute of Complex Systems and Institute of Biomedical Sciences, Fudan University Shanghai 200438 China
| | - Shaowen Wu
- Guangdong Key Laboratory for Crop Germplasm Resources Preservation and Utilization, Agro-biological Gene Research Center, Guangdong Academy of Agricultural Sciences Guangzhou 510640 Guangdong China
| | | | - Xingyu Song
- Department of Chemistry, Multiscale Research Institute of Complex Systems and Institute of Biomedical Sciences, Fudan University Shanghai 200438 China
| | - Maohua Yang
- Department of Chemistry, Multiscale Research Institute of Complex Systems and Institute of Biomedical Sciences, Fudan University Shanghai 200438 China
| | - Wolun Zhang
- LightEdge Technologies Limited Zhongshan 528403 China
| | - Shaohui Huang
- Institute of Biophysics, Chinese Academy of Sciences Beijing 100101 China
- University of Chinese Academy of Science Beijing 101408 China
| | - Jingwei Weng
- Department of Chemistry, Multiscale Research Institute of Complex Systems and Institute of Biomedical Sciences, Fudan University Shanghai 200438 China
| | - Zhijun Liu
- National Facility for Protein Science in Shanghai, Zhangjiang Lab, Shanghai Advanced Research Institute, Chinese Academy of Sciences Shanghai 201210 China
| | - Wenning Wang
- Department of Chemistry, Multiscale Research Institute of Complex Systems and Institute of Biomedical Sciences, Fudan University Shanghai 200438 China
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21
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He W, Henning-Knechtel A, Kirmizialtin S. Visualizing RNA Structures by SAXS-Driven MD Simulations. FRONTIERS IN BIOINFORMATICS 2022; 2:781949. [PMID: 36304317 PMCID: PMC9580860 DOI: 10.3389/fbinf.2022.781949] [Citation(s) in RCA: 2] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/21/2021] [Accepted: 01/04/2022] [Indexed: 12/26/2022] Open
Abstract
The biological role of biomolecules is intimately linked to their structural dynamics. Experimental or computational techniques alone are often insufficient to determine accurate structural ensembles in atomic detail. We use all-atom molecular dynamics (MD) simulations and couple it to small-angle X-ray scattering (SAXS) experiments to resolve the structural dynamics of RNA molecules. To accomplish this task, we utilize a set of re-weighting and biasing techniques tailored for RNA molecules. To showcase our approach, we study two RNA molecules: a riboswitch that shows structural variations upon ligand binding, and a two-way junction RNA that displays structural heterogeneity and sensitivity to salt conditions. Integration of MD simulations and experiments allows the accurate construction of conformational ensembles of RNA molecules. We observe a dynamic change of the SAM-I riboswitch conformations depending on its binding partners. The binding of SAM and Mg2+ cations stabilizes the compact state. The absence of Mg2+ or SAM leads to the loss of tertiary contacts, resulting in a dramatic expansion of the riboswitch conformations. The sensitivity of RNA structures to the ionic strength demonstrates itself in the helix junction helix (HJH). The HJH shows non-monotonic compaction as the ionic strength increases. The physics-based picture derived from the experimentally guided MD simulations allows biophysical characterization of RNA molecules. All in all, SAXS-guided MD simulations offer great prospects for studying RNA structural dynamics.
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Affiliation(s)
- Weiwei He
- Chemistry Program, Science Division, New York University Abu Dhabi, Abu Dhabi, United Arab Emirates
- Department of Chemistry, New York University, New York, NY, United States
| | - Anja Henning-Knechtel
- Chemistry Program, Science Division, New York University Abu Dhabi, Abu Dhabi, United Arab Emirates
| | - Serdal Kirmizialtin
- Chemistry Program, Science Division, New York University Abu Dhabi, Abu Dhabi, United Arab Emirates
- *Correspondence: Serdal Kirmizialtin,
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22
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Conformational ensembles of intrinsically disordered proteins and flexible multidomain proteins. Biochem Soc Trans 2022; 50:541-554. [PMID: 35129612 DOI: 10.1042/bst20210499] [Citation(s) in RCA: 31] [Impact Index Per Article: 15.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/10/2021] [Revised: 01/13/2022] [Accepted: 01/17/2022] [Indexed: 12/29/2022]
Abstract
Intrinsically disordered proteins (IDPs) and multidomain proteins with flexible linkers show a high level of structural heterogeneity and are best described by ensembles consisting of multiple conformations with associated thermodynamic weights. Determining conformational ensembles usually involves the integration of biophysical experiments and computational models. In this review, we discuss current approaches to determine conformational ensembles of IDPs and multidomain proteins, including the choice of biophysical experiments, computational models used to sample protein conformations, models to calculate experimental observables from protein structure, and methods to refine ensembles against experimental data. We also provide examples of recent applications of integrative conformational ensemble determination to study IDPs and multidomain proteins and suggest future directions for research in the field.
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23
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Vermeer B, Schmid S. Can DyeCycling break the photobleaching limit in single-molecule FRET? NANO RESEARCH 2022; 15:9818-9830. [PMID: 35582137 PMCID: PMC9101981 DOI: 10.1007/s12274-022-4420-5] [Citation(s) in RCA: 5] [Impact Index Per Article: 2.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Subscribe] [Scholar Register] [Received: 02/04/2022] [Revised: 04/05/2022] [Accepted: 04/06/2022] [Indexed: 05/03/2023]
Abstract
UNLABELLED Biomolecular systems, such as proteins, crucially rely on dynamic processes at the nanoscale. Detecting biomolecular nanodynamics is therefore key to obtaining a mechanistic understanding of the energies and molecular driving forces that control biomolecular systems. Single-molecule fluorescence resonance energy transfer (smFRET) is a powerful technique to observe in real-time how a single biomolecule proceeds through its functional cycle involving a sequence of distinct structural states. Currently, this technique is fundamentally limited by irreversible photobleaching, causing the untimely end of the experiment and thus, a narrow temporal bandwidth of ≤ 3 orders of magnitude. Here, we introduce "DyeCycling", a measurement scheme with which we aim to break the photobleaching limit in smFRET. We introduce the concept of spontaneous dye replacement by simulations, and as an experimental proof-of-concept, we demonstrate the intermittent observation of a single biomolecule for one hour with a time resolution of milliseconds. Theoretically, DyeCycling can provide > 100-fold more information per single molecule than conventional smFRET. We discuss the experimental implementation of DyeCycling, its current and fundamental limitations, and specific biological use cases. Given its general simplicity and versatility, DyeCycling has the potential to revolutionize the field of time-resolved smFRET, where it may serve to unravel a wealth of biomolecular dynamics by bridging from milliseconds to the hour range. ELECTRONIC SUPPLEMENTARY MATERIAL Supplementary material is available for this article at 10.1007/s12274-022-4420-5 and is accessible for authorized users.
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Affiliation(s)
- Benjamin Vermeer
- NanoDynamicsLab, Laboratory of Biophysics, Wageningen University, Stippeneng 4, 6708WE Wageningen, The Netherlands
| | - Sonja Schmid
- NanoDynamicsLab, Laboratory of Biophysics, Wageningen University, Stippeneng 4, 6708WE Wageningen, The Netherlands
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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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Klose D, Holla A, Gmeiner C, Nettels D, Ritsch I, Bross N, Yulikov M, Allain FHT, Schuler B, Jeschke G. Resolving distance variations by single-molecule FRET and EPR spectroscopy using rotamer libraries. Biophys J 2021; 120:4842-4858. [PMID: 34536387 PMCID: PMC8595751 DOI: 10.1016/j.bpj.2021.09.021] [Citation(s) in RCA: 17] [Impact Index Per Article: 5.7] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/26/2021] [Revised: 07/09/2021] [Accepted: 09/13/2021] [Indexed: 01/14/2023] Open
Abstract
Förster resonance energy transfer (FRET) and electron paramagnetic resonance (EPR) spectroscopy are complementary techniques for quantifying distances in the nanometer range. Both approaches are commonly employed for probing the conformations and conformational changes of biological macromolecules based on site-directed fluorescent or paramagnetic labeling. FRET can be applied in solution at ambient temperature and thus provides direct access to dynamics, especially if used at the single-molecule level, whereas EPR requires immobilization or work at cryogenic temperatures but provides data that can be more reliably used to extract distance distributions. However, a combined analysis of the complementary data from the two techniques has been complicated by the lack of a common modeling framework. Here, we demonstrate a systematic analysis approach based on rotamer libraries for both FRET and EPR labels to predict distance distributions between two labels from a structural model. Dynamics of the fluorophores within these distance distributions are taken into account by diffusional averaging, which improves the agreement with experiment. Benchmarking this methodology with a series of surface-exposed pairs of sites in a structured protein domain reveals that the lowest resolved distance differences can be as small as ∼0.25 nm for both techniques, with quantitative agreement between experimental and simulated transfer efficiencies within a range of ±0.045. Rotamer library analysis thus establishes a coherent way of treating experimental data from EPR and FRET and provides a basis for integrative structural modeling, including studies of conformational distributions and dynamics of biological macromolecules using both techniques.
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Affiliation(s)
- Daniel Klose
- Department of Chemistry and Applied Biosciences, ETH Zurich, Zurich, Switzerland.
| | - Andrea Holla
- Department of Biochemistry, University of Zurich, Zurich, Switzerland
| | - Christoph Gmeiner
- Department of Chemistry and Applied Biosciences, ETH Zurich, Zurich, Switzerland
| | - Daniel Nettels
- Department of Biochemistry, University of Zurich, Zurich, Switzerland
| | - Irina Ritsch
- Department of Chemistry and Applied Biosciences, ETH Zurich, Zurich, Switzerland
| | - Nadja Bross
- Department of Chemistry, University of Zurich, Zurich, Switzerland
| | - Maxim Yulikov
- Department of Chemistry and Applied Biosciences, ETH Zurich, Zurich, Switzerland
| | | | - Benjamin Schuler
- Department of Biochemistry, University of Zurich, Zurich, Switzerland; Department of Physics, University of Zurich, Zurich, Switzerland.
| | - Gunnar Jeschke
- Department of Chemistry and Applied Biosciences, ETH Zurich, Zurich, Switzerland
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26
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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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27
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Das D, Arora L, Mukhopadhyay S. Fluorescence Depolarization Kinetics Captures Short-Range Backbone Dihedral Rotations and Long-Range Correlated Dynamics of an Intrinsically Disordered Protein. J Phys Chem B 2021; 125:9708-9718. [PMID: 34415768 DOI: 10.1021/acs.jpcb.1c04426] [Citation(s) in RCA: 6] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/28/2022]
Abstract
Intrinsically disordered proteins (IDPs) do not autonomously fold into well-defined three-dimensional structures and are best described as a heterogeneous ensemble of rapidly interconverting conformers. It is challenging to elucidate their complex dynamic signatures using a single technique. In this study, we employed sensitive fluorescence depolarization kinetics by following picosecond time-resolved fluorescence anisotropy decays to directly capture the essential dynamical features of intrinsically disordered α-synuclein (α-syn) site-specifically labeled with thiol-active fluorophores. By utilizing a long-lifetime (≥10 ns) anisotropic label, we were able to discern three distinct rotational components of α-syn. The subnanosecond component represents the local wobbling-in-cone motion of the fluorophore, whereas the slower (∼1.4 ns) component corresponds to the short-range backbone dynamics governed by collective torsional fluctuations in the Ramachandran Φ-Ψ dihedral space. This backbone dihedral rotational time scale is sensitive to the local chain stiffness and slows down in the presence of an adjacent proline residue. We also observed a small-amplitude (≤10%) slower rotational correlation time (6-10 ns) that represents the long-range correlated dynamics involving a much longer segment of the polypeptide chain. These intrinsic dynamic signatures of IDPs will provide critical mechanistic underpinnings in a mosaic of biophysical phenomena involving internal friction, allosteric interactions, and phase separation.
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28
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Alston JJ, Soranno A, Holehouse AS. Integrating single-molecule spectroscopy and simulations for the study of intrinsically disordered proteins. Methods 2021; 193:116-135. [PMID: 33831596 PMCID: PMC8713295 DOI: 10.1016/j.ymeth.2021.03.018] [Citation(s) in RCA: 21] [Impact Index Per Article: 7.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/17/2020] [Revised: 03/25/2021] [Accepted: 03/31/2021] [Indexed: 12/21/2022] Open
Abstract
Over the last two decades, intrinsically disordered proteins and protein regions (IDRs) have emerged from a niche corner of biophysics to be recognized as essential drivers of cellular function. Various techniques have provided fundamental insight into the function and dysfunction of IDRs. Among these techniques, single-molecule fluorescence spectroscopy and molecular simulations have played a major role in shaping our modern understanding of the sequence-encoded conformational behavior of disordered proteins. While both techniques are frequently used in isolation, when combined they offer synergistic and complementary information that can help uncover complex molecular details. Here we offer an overview of single-molecule fluorescence spectroscopy and molecular simulations in the context of studying disordered proteins. We discuss the various means in which simulations and single-molecule spectroscopy can be integrated, and consider a number of studies in which this integration has uncovered biological and biophysical mechanisms.
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Affiliation(s)
- Jhullian J Alston
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis 63110, MO, USA; Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis 63130, MO, USA
| | - Andrea Soranno
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis 63110, MO, USA; Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis 63130, MO, USA.
| | - Alex S Holehouse
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis 63110, MO, USA; Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis 63130, MO, USA.
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29
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DNA sequence-dependent positioning of the linker histone in a nucleosome: A single-pair FRET study. Biophys J 2021; 120:3747-3763. [PMID: 34293303 DOI: 10.1016/j.bpj.2021.07.012] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/14/2021] [Revised: 06/25/2021] [Accepted: 07/13/2021] [Indexed: 01/01/2023] Open
Abstract
Linker histones (LHs) bind to nucleosomes with their globular domain (gH) positioned in either an on- or an off-dyad binding mode. Here, we study the effect of the linker DNA (L-DNA) sequence on the binding of a full-length LH, Xenopus laevis H1.0b, to a Widom 601 nucleosome core particle (NCP) flanked by two 40 bp long L-DNA arms, by single-pair FRET spectroscopy. We varied the sequence of the 11 bp of L-DNA adjoining the NCP on either side, making the sequence either A-tract, purely GC, or mixed with 64% AT. The labeled gH consistently exhibited higher FRET efficiency with the labeled L-DNA containing the A-tract than that with the pure-GC stretch, even when the stretches were swapped. However, it did not exhibit higher FRET efficiency with the L-DNA containing 64% AT-rich mixed DNA when compared to the pure-GC stretch. We explain our observations with a model that shows that the gH binds on dyad and that two arginines mediate recognition of the A-tract via its characteristically narrow minor groove. To investigate whether this on-dyad minor groove-based recognition was distinct from previously identified off-dyad major groove-based recognition, a nucleosome was designed with A-tracts on both the L-DNA arms. One A-tract was complementary to thymine and the other to deoxyuridine. The major groove of the thymine-tract was lined with methyl groups that were absent from the major groove of the deoxyuridine tract. The gH exhibited similar FRET for both these A-tracts, suggesting that it does not interact with the thymine methyl groups exposed on the major groove. Our observations thus complement previous studies that suggest that different LH isoforms may employ different ways of recognizing AT-rich DNA and A-tracts. This adaptability may enable the LH to universally compact scaffold-associated regions and constitutive heterochromatin, which are rich in such sequences.
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30
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Han MJ, He QT, Yang M, Chen C, Yao Y, Liu X, Wang Y, Zhu ZL, Zhu KK, Qu C, Yang F, Hu C, Guo X, Zhang D, Chen C, Sun JP, Wang J. Single-molecule FRET and conformational analysis of beta-arrestin-1 through genetic code expansion and a Se-click reaction. Chem Sci 2021; 12:9114-9123. [PMID: 34276941 PMCID: PMC8261736 DOI: 10.1039/d1sc02653d] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.7] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/14/2021] [Revised: 07/02/2021] [Accepted: 05/27/2021] [Indexed: 11/21/2022] Open
Abstract
Single-molecule Förster resonance energy transfer (smFRET) is a powerful tool for investigating the dynamic properties of biomacromolecules. However, the success of protein smFRET relies on the precise and efficient labeling of two or more fluorophores on the protein of interest (POI), which has remained highly challenging, particularly for large membrane protein complexes. Here, we demonstrate the site-selective incorporation of a novel unnatural amino acid (2-amino-3-(4-hydroselenophenyl) propanoic acid, SeF) through genetic expansion followed by a Se-click reaction to conjugate the Bodipy593 fluorophore on calmodulin (CaM) and β-arrestin-1 (βarr1). Using this strategy, we monitored the subtle but functionally important conformational change of βarr1 upon activation by the G-protein coupled receptor (GPCR) through smFRET for the first time. Our new method has broad applications for the site-specific labeling and smFRET measurement of membrane protein complexes, and the elucidation of their dynamic properties such as transducer protein selection.
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Affiliation(s)
- Ming-Jie Han
- Tianjin Institute of Industrial Biotechnology, Chinese Academy of Sciences Tianjin Airport Economic Area Tianjin 300308 China
| | - Qing-Tao He
- Key Laboratory Experimental Teratology of the Ministry of Education and Department of Biochemistry and Molecular Biology, School of Basic Medical Sciences, Cheeloo College of Medicine, Shandong University 44 Wenhua Xi Road Jinan 250012 Shandong China
- Department of Physiology and Pathophysiology, School of Basic Medical Sciences, Peking University, Key Laboratory of Molecular Cardiovascular Science, Ministry of Education Haidian District Beijing 100191 China
- Institute of Biophysics, Chinese Academy of Sciences Chaoyang District Beijing 100101 China
| | - Mengyi Yang
- School of Life Sciences, Tsinghua-Peking Joint Center for Life Sciences, Beijing Advanced Innovation Center for Structural Biology, Tsinghua University Haidian District Beijing 100084 China
| | - Chao Chen
- Tianjin Institute of Industrial Biotechnology, Chinese Academy of Sciences Tianjin Airport Economic Area Tianjin 300308 China
- University of the Chinese Academy of Sciences (UCAS) Shijingshan District Beijing 100049 China
| | - Yirong Yao
- School of Life Sciences, Tsinghua-Peking Joint Center for Life Sciences, Beijing Advanced Innovation Center for Structural Biology, Tsinghua University Haidian District Beijing 100084 China
| | - Xiaohong Liu
- Institute of Biophysics, Chinese Academy of Sciences Chaoyang District Beijing 100101 China
| | - Yuchuan Wang
- Shenzhen Institute of Transfusion Medicine, Shenzhen Blood Center Futian District Shenzhen 518052 China
| | - Zhong-Liang Zhu
- School of Life Sciences, University of Science and Technology of China Baohe District Anhui 230026 China
| | - Kong-Kai Zhu
- School of Biological Science and Technology, University of Jinan Jinan Shandong 250022 China
| | - Changxiu Qu
- Key Laboratory Experimental Teratology of the Ministry of Education and Department of Biochemistry and Molecular Biology, School of Basic Medical Sciences, Cheeloo College of Medicine, Shandong University 44 Wenhua Xi Road Jinan 250012 Shandong China
| | - Fan Yang
- Key Laboratory Experimental Teratology of the Ministry of Education and Department of Biochemistry and Molecular Biology, School of Basic Medical Sciences, Cheeloo College of Medicine, Shandong University 44 Wenhua Xi Road Jinan 250012 Shandong China
| | - Cheng Hu
- Institute of Biophysics, Chinese Academy of Sciences Chaoyang District Beijing 100101 China
| | - Xuzhen Guo
- Institute of Biophysics, Chinese Academy of Sciences Chaoyang District Beijing 100101 China
| | - Dawei Zhang
- Tianjin Institute of Industrial Biotechnology, Chinese Academy of Sciences Tianjin Airport Economic Area Tianjin 300308 China
| | - Chunlai Chen
- School of Life Sciences, Tsinghua-Peking Joint Center for Life Sciences, Beijing Advanced Innovation Center for Structural Biology, Tsinghua University Haidian District Beijing 100084 China
| | - Jin-Peng Sun
- Key Laboratory Experimental Teratology of the Ministry of Education and Department of Biochemistry and Molecular Biology, School of Basic Medical Sciences, Cheeloo College of Medicine, Shandong University 44 Wenhua Xi Road Jinan 250012 Shandong China
- Department of Physiology and Pathophysiology, School of Basic Medical Sciences, Peking University, Key Laboratory of Molecular Cardiovascular Science, Ministry of Education Haidian District Beijing 100191 China
| | - Jiangyun Wang
- Institute of Biophysics, Chinese Academy of Sciences Chaoyang District Beijing 100101 China
- University of the Chinese Academy of Sciences (UCAS) Shijingshan District Beijing 100049 China
- Shenzhen Institute of Transfusion Medicine, Shenzhen Blood Center Futian District Shenzhen 518052 China
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31
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Ucar H, Wagenknecht HA. DNA-templated control of chirality and efficient energy transport in supramolecular DNA architectures with aggregation-induced emission. Chem Sci 2021; 12:10048-10053. [PMID: 34377398 PMCID: PMC8317660 DOI: 10.1039/d1sc02351a] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.7] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/27/2021] [Accepted: 06/19/2021] [Indexed: 01/15/2023] Open
Abstract
Two conjugates of tetraphenylethylene with d-2′-deoxyuridine (1d) and l-2′-deoxyuridine (1l) were synthesized to construct new supramolecular DNA-architectures by self-assembly. The non-templated assemblies of 1d and 1l show strong aggregation-induced emission and their chirality is exclusively controlled by the configuration of their sugar part. In contrast, the chirality of the DNA-templated assemblies is governed by the configuration of the DNA, and there is no configuration-selective binding of 1d to d-A20 and 1l to l-A20. The quantum yield of the assembly of 1d along the single-stranded DNA A20 is 0.40; approximately every second available binding site on the DNA template is occupied by 1d. The strong aggregation-induced emission of these DNA architectures can be efficiently quenched and the excitation energy can be transported to Atto dyes at the 5′-terminus. A multistep energy transport “hopping” precedes the final energy transfer to the terminal acceptor. The building block 1d promotes this energy transport as stepping stones. This was elucidated by reference DNA double strands in which 1d was covalently incorporated at two distinct sites in the sequences, one near the Atto dye, and one farther away. This new type of completely self-assembled supramolecular DNA architecture is hierarchically ordered and the DNA template controls not only the binding but also the energy transport properties. The high intensity of the aggregation-induced emission and the excellent energy transport properties make these DNA-based materials promising candidates for optoelectronic applications. DNA architectures with tetraphenylethylene are assembled in a non-covalent way. The strong aggregation-induced emission of the chromophores is quenched and the energy is transported to Atto dyes by a multistep energy “hopping”.![]()
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Affiliation(s)
- Hülya Ucar
- Institute of Organic Chemistry, Karlsruhe Institute of Technology (KIT) Fritz-Haber-Weg 6 76131 Karlsruhe German
| | - Hans-Achim Wagenknecht
- Institute of Organic Chemistry, Karlsruhe Institute of Technology (KIT) Fritz-Haber-Weg 6 76131 Karlsruhe German
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32
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Guo X, Li Q, Xiang J, Liu M, Guan A, Tang Y, Sun H. A hybrid aggregate FRET probe from the mixed assembly of cyanine dyes for highly specific monitoring of mitochondria autophagy. Anal Chim Acta 2021; 1165:338561. [PMID: 33975703 DOI: 10.1016/j.aca.2021.338561] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/17/2021] [Revised: 03/27/2021] [Accepted: 04/19/2021] [Indexed: 10/21/2022]
Abstract
Mitochondria autophagy, also known as mitophagy, is a process in which mitochondria are wrapped by autophagosomes and fused with lysosomes for degradation. This process is essential for mitochondrial quality control. Here, we developed a hybrid aggregate FRET probe through mixed assembly of two cyanine dyes FMOTY and AMTC. In live cells, FMOTY and AMTC exist independently in lysosomes and mitochondria and will not produce interfering FRET background signals. The FRET signal is only generated when mitochondria is transported to lysosomes during mitophagy. This allows the hybridized aggregate to be used as a highly specific probe for monitoring mitophagy.
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Affiliation(s)
- Xiaomeng Guo
- Beijing National Laboratory for Molecular Sciences (BNLMS), State Key Laboratory for Structural Chemistry of Unstable and Stable Species, Institute of Chemistry Chinese Academy of Sciences, Beijing, 100190, PR China; University of Chinese Academy of Sciences, Beijing, 100049, PR China
| | - Qian Li
- Center for Physicochemical Analysis and Measurement, Institute of Chemistry Chinese Academy of Sciences, Beijing, 100190, PR China
| | - Junfeng Xiang
- Center for Physicochemical Analysis and Measurement, Institute of Chemistry Chinese Academy of Sciences, Beijing, 100190, PR China
| | - Meirong Liu
- Center for Physicochemical Analysis and Measurement, Institute of Chemistry Chinese Academy of Sciences, Beijing, 100190, PR China
| | - Aijiao Guan
- Beijing National Laboratory for Molecular Sciences (BNLMS), State Key Laboratory for Structural Chemistry of Unstable and Stable Species, Institute of Chemistry Chinese Academy of Sciences, Beijing, 100190, PR China
| | - Yalin Tang
- Beijing National Laboratory for Molecular Sciences (BNLMS), State Key Laboratory for Structural Chemistry of Unstable and Stable Species, Institute of Chemistry Chinese Academy of Sciences, Beijing, 100190, PR China; University of Chinese Academy of Sciences, Beijing, 100049, PR China
| | - Hongxia Sun
- Beijing National Laboratory for Molecular Sciences (BNLMS), State Key Laboratory for Structural Chemistry of Unstable and Stable Species, Institute of Chemistry Chinese Academy of Sciences, Beijing, 100190, PR China.
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33
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Hernández-Adame PL, Meza U, Rodríguez-Menchaca AA, Sánchez-Armass S, Ruiz-García J, Gomez E. Determination of the size of lipid rafts studied through single-molecule FRET simulations. Biophys J 2021; 120:2287-2295. [PMID: 33864789 DOI: 10.1016/j.bpj.2021.04.003] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.7] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/07/2020] [Revised: 02/16/2021] [Accepted: 04/08/2021] [Indexed: 11/16/2022] Open
Abstract
Fluorescence resonance energy transfer (FRET) is a high-resolution technique that allows the characterization of spatial and temporal properties of biological structures and mechanisms. In this work, we developed an in silico single-molecule FRET methodology to study the dynamics of fluorophores inside lipid rafts. We monitored the fluorescence of a single acceptor molecule in the presence of several donor molecules. By looking at the average fluorescence, we selected events with single acceptor and donor molecules, and we used them to determine the raft size in the range of 5-16 nm. We conclude that our method is robust and insensitive to variations in the diffusion coefficient, donor density, or selected fluorescence threshold.
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Affiliation(s)
| | - Ulises Meza
- Department of Physiology and Biophysics, School of Medicine, Universidad Autónoma de San Luis Potosí, San Luis Potosí, Mexico
| | - Aldo A Rodríguez-Menchaca
- Department of Physiology and Biophysics, School of Medicine, Universidad Autónoma de San Luis Potosí, San Luis Potosí, Mexico
| | - Sergio Sánchez-Armass
- Department of Physiology and Biophysics, School of Medicine, Universidad Autónoma de San Luis Potosí, San Luis Potosí, Mexico
| | - Jaime Ruiz-García
- Biological Physics Laboratory, Physics Institute, San Luis Potosí, Mexico.
| | - Eduardo Gomez
- Biological Physics Laboratory, Physics Institute, San Luis Potosí, Mexico.
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34
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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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35
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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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36
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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: 125] [Impact Index Per Article: 41.7] [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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37
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Hart SM, Chen WJ, Banal JL, Bricker WP, Dodin A, Markova L, Vyborna Y, Willard AP, Häner R, Bathe M, Schlau-Cohen GS. Engineering couplings for exciton transport using synthetic DNA scaffolds. Chem 2021. [DOI: 10.1016/j.chempr.2020.12.020] [Citation(s) in RCA: 12] [Impact Index Per Article: 4.0] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 02/08/2023]
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38
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Wolf S, Sohmen B, Hellenkamp B, Thurn J, Stock G, Hugel T. Hierarchical dynamics in allostery following ATP hydrolysis monitored by single molecule FRET measurements and MD simulations. Chem Sci 2021; 12:3350-3359. [PMID: 34164105 PMCID: PMC8179424 DOI: 10.1039/d0sc06134d] [Citation(s) in RCA: 14] [Impact Index Per Article: 4.7] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/07/2020] [Accepted: 01/14/2021] [Indexed: 02/06/2023] Open
Abstract
We report on a study that combines advanced fluorescence methods with molecular dynamics (MD) simulations to cover timescales from nanoseconds to milliseconds for a large protein. This allows us to delineate how ATP hydrolysis in a protein causes allosteric changes at a distant protein binding site, using the chaperone Hsp90 as test system. The allosteric process occurs via hierarchical dynamics involving timescales from nano- to milliseconds and length scales from Ångstroms to several nanometers. We find that hydrolysis of one ATP is coupled to a conformational change of Arg380, which in turn passes structural information via the large M-domain α-helix to the whole protein. The resulting structural asymmetry in Hsp90 leads to the collapse of a central folding substrate binding site, causing the formation of a novel collapsed state (closed state B) that we characterise structurally. We presume that similar hierarchical mechanisms are fundamental for information transfer induced by ATP hydrolysis through many other proteins.
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Affiliation(s)
- Steffen Wolf
- Biomolecular Dynamics, Institute of Physics, University of Freiburg Freiburg Germany +49 761 203 5883 +49 761 203 5913
| | - Benedikt Sohmen
- Institute of Physical Chemistry, University of Freiburg Freiburg Germany +49 761 203 6192
| | - Björn Hellenkamp
- Engineering and Applied Sciences, Columbia University New York USA
| | - Johann Thurn
- Institute of Physical Chemistry, University of Freiburg Freiburg Germany +49 761 203 6192
| | - Gerhard Stock
- Biomolecular Dynamics, Institute of Physics, University of Freiburg Freiburg Germany +49 761 203 5883 +49 761 203 5913
| | - Thorsten Hugel
- Institute of Physical Chemistry, University of Freiburg Freiburg Germany +49 761 203 6192
- Signalling Research Centers BIOSS and CIBSS, University of Freiburg Freiburg Germany
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39
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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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40
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Frommer J, Müller S. Changed reactivity of secondary hydroxy groups in C8-modified adenosine - lessons learned from silylation. Beilstein J Org Chem 2020; 16:2854-2861. [PMID: 33299483 PMCID: PMC7705864 DOI: 10.3762/bjoc.16.234] [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: 09/08/2020] [Accepted: 11/03/2020] [Indexed: 12/03/2022] Open
Abstract
Synthesis of site-specifically modified oligonucleotides has become a major tool for RNA structure and function studies. Reporter groups or specific functional entities are required to be attached at a pre-defined site of the oligomer. An attractive strategy is the incorporation of suitably functionalized building blocks that allow post-synthetic conjugation of the desired moiety. A C8-alkynyl-modified adenosine derivative was synthesized, reviving an old synthetic pathway for iodination of purine nucleobases. Silylation of the C8-alkynyl-modified adenosine revealed unexpected selectivity of the two secondary sugar hydroxy groups, with the 3'-O-isomer being preferentially formed. Optimization of the protection scheme lead to a new and economic route to the desired C8-alkynylated building block and its incorporation in RNA.
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Affiliation(s)
- Jennifer Frommer
- Institute for Biochemistry, University Greifswald, Felix-Hausdorff Str. 4, D-17487 Greifswald, Germany
- School of Chemistry, University of Birmingham, Edgbaston, Birmingham B15 2TT, UK
| | - Sabine Müller
- Institute for Biochemistry, University Greifswald, Felix-Hausdorff Str. 4, D-17487 Greifswald, Germany
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41
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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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42
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Liu J, Cheng R, Van Eps N, Wang N, Morizumi T, Ou WL, Klauser PC, Rozovsky S, Ernst OP, Wang L. Genetically Encoded Quinone Methides Enabling Rapid, Site-Specific, and Photocontrolled Protein Modification with Amine Reagents. J Am Chem Soc 2020; 142:17057-17068. [PMID: 32915556 DOI: 10.1021/jacs.0c06820] [Citation(s) in RCA: 21] [Impact Index Per Article: 5.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/19/2022]
Abstract
Site-specific modification of proteins with functional molecules provides powerful tools for researching and engineering proteins. Here we report a new chemical conjugation method which photocages highly reactive but chemically selective moieties, enabling the use of protein-inert amines for selective protein modification. New amino acids FnbY and FmnbY, bearing photocaged quinone methides (QMs), were genetically incorporated into proteins. Upon light activation, they generated highly reactive QM, which rapidly reacted with amine derivatives. This method features a rare combination of desired properties including fast kinetics, small and stable linkage, compatibility with low temperature, photocontrollability, and widely available reagents. Moreover, labeling via FnbY occurs on the β-carbon, affording the shortest linkage to protein backbone which is essential for advanced studies involving orientation and distance. We installed various functionalities onto proteins and attached a spin label as close as possible to the protein backbone, achieving high resolution in double electron-electron paramagnetic resonance distance measurements.
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Affiliation(s)
- Jun Liu
- Department of Pharmaceutical Chemistry and the Cardiovascular Research Institute, University of California San Francisco, 555 Mission Bay Blvd. South, San Francisco, California 94158, United States
| | - Rujin Cheng
- Department of Chemistry and Biochemistry, University of Delaware, Newark, Delaware 19716, United States
| | - Ned Van Eps
- Department of Biochemistry, University of Toronto, Toronto, Ontario M5S 1A8, Canada
| | - Nanxi Wang
- Department of Pharmaceutical Chemistry and the Cardiovascular Research Institute, University of California San Francisco, 555 Mission Bay Blvd. South, San Francisco, California 94158, United States
| | - Takefumi Morizumi
- Department of Biochemistry, University of Toronto, Toronto, Ontario M5S 1A8, Canada
| | - Wei-Lin Ou
- Department of Biochemistry, University of Toronto, Toronto, Ontario M5S 1A8, Canada
| | - Paul C Klauser
- Department of Pharmaceutical Chemistry and the Cardiovascular Research Institute, University of California San Francisco, 555 Mission Bay Blvd. South, San Francisco, California 94158, United States
| | - Sharon Rozovsky
- Department of Chemistry and Biochemistry, University of Delaware, Newark, Delaware 19716, United States
| | - Oliver P Ernst
- Department of Biochemistry, University of Toronto, Toronto, Ontario M5S 1A8, Canada.,Department of Molecular Genetics, University of Toronto, Toronto, Ontario M5S 1A8, Canada
| | - Lei Wang
- Department of Pharmaceutical Chemistry and the Cardiovascular Research Institute, University of California San Francisco, 555 Mission Bay Blvd. South, San Francisco, California 94158, United States
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43
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Gomes GNW, Krzeminski M, Namini A, Martin EW, Mittag T, Head-Gordon T, Forman-Kay JD, Gradinaru CC. Conformational Ensembles of an Intrinsically Disordered Protein Consistent with NMR, SAXS, and Single-Molecule FRET. J Am Chem Soc 2020; 142:15697-15710. [PMID: 32840111 PMCID: PMC9987321 DOI: 10.1021/jacs.0c02088] [Citation(s) in RCA: 95] [Impact Index Per Article: 23.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/20/2022]
Abstract
Intrinsically disordered proteins (IDPs) have fluctuating heterogeneous conformations, which makes their structural characterization challenging. Although challenging, characterization of the conformational ensembles of IDPs is of great interest, since their conformational ensembles are the link between their sequences and functions. An accurate description of IDP conformational ensembles depends crucially on the amount and quality of the experimental data, how it is integrated, and if it supports a consistent structural picture. We used integrative modeling and validation to apply conformational restraints and assess agreement with the most common structural techniques for IDPs: Nuclear Magnetic Resonance (NMR) spectroscopy, Small-angle X-ray Scattering (SAXS), and single-molecule Förster Resonance Energy Transfer (smFRET). Agreement with such a diverse set of experimental data suggests that details of the generated ensembles can now be examined with a high degree of confidence. Using the disordered N-terminal region of the Sic1 protein as a test case, we examined relationships between average global polymeric descriptions and higher-moments of their distributions. To resolve apparent discrepancies between smFRET and SAXS inferences, we integrated SAXS data with NMR data and reserved the smFRET data for independent validation. Consistency with smFRET, which was not guaranteed a priori, indicates that, globally, the perturbative effects of NMR or smFRET labels on the Sic1 ensemble are minimal. Analysis of the ensembles revealed distinguishing features of Sic1, such as overall compactness and large end-to-end distance fluctuations, which are consistent with biophysical models of Sic1's ultrasensitive binding to its partner Cdc4. Our results underscore the importance of integrative modeling and validation in generating and drawing conclusions from IDP conformational ensembles.
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Affiliation(s)
- Gregory-Neal W Gomes
- Department of Physics, University of Toronto, Toronto, Ontario M5G 1X8, Canada.,Department of Chemical and Physical Sciences, University of Toronto Mississauga, Mississauga, Ontario L5L 1C6, Canada
| | - Mickaël Krzeminski
- Molecular Medicine Program, Hospital for Sick Children, Toronto, Ontario M5S 1A8, Canada.,Department of Biochemistry, University of Toronto, Toronto, Ontario M5G 1X8, Canada
| | - Ashley Namini
- Department of Physics, University of Toronto, Toronto, Ontario M5G 1X8, Canada.,Department of Chemical and Physical Sciences, University of Toronto Mississauga, Mississauga, Ontario L5L 1C6, Canada
| | - Erik W Martin
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, Tennessee 38105, United States
| | - Tanja Mittag
- Department of Structural Biology, St. Jude Children's Research Hospital, Memphis, Tennessee 38105, United States
| | - Teresa Head-Gordon
- Departments of Chemistry, Bioengineering, Chemical and Biomolecular Engineering University of California, Berkeley, California 94720, United States
| | - Julie D Forman-Kay
- Molecular Medicine Program, Hospital for Sick Children, Toronto, Ontario M5S 1A8, Canada.,Department of Biochemistry, University of Toronto, Toronto, Ontario M5G 1X8, Canada
| | - Claudiu C Gradinaru
- Department of Physics, University of Toronto, Toronto, Ontario M5G 1X8, Canada.,Department of Chemical and Physical Sciences, University of Toronto Mississauga, Mississauga, Ontario L5L 1C6, Canada
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44
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Hirashima S, Sugiyama H, Park S. Construction of a FRET System in a Double-Stranded DNA Using Fluorescent Thymidine and Cytidine Analogs. J Phys Chem B 2020; 124:8794-8800. [DOI: 10.1021/acs.jpcb.0c06879] [Citation(s) in RCA: 6] [Impact Index Per Article: 1.5] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/20/2023]
Affiliation(s)
- Shingo Hirashima
- Department of Chemistry, Graduate School of Science, Kyoto University, Sakyo, Kyoto 606-8502, Japan
| | - Hiroshi Sugiyama
- Department of Chemistry, Graduate School of Science, Kyoto University, Sakyo, Kyoto 606-8502, Japan
- Institute for Integrated Cell-Material Science (WPI-iCeMS), Kyoto University, Sakyo, Kyoto 606-8501, Japan
| | - Soyoung Park
- Department of Chemistry, Graduate School of Science, Kyoto University, Sakyo, Kyoto 606-8502, Japan
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45
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Paudel BP, Moye AL, Abou Assi H, El-Khoury R, Cohen SB, Holien JK, Birrento ML, Samosorn S, Intharapichai K, Tomlinson CG, Teulade-Fichou MP, González C, Beck JL, Damha MJ, van Oijen AM, Bryan TM. A mechanism for the extension and unfolding of parallel telomeric G-quadruplexes by human telomerase at single-molecule resolution. eLife 2020; 9:56428. [PMID: 32723475 PMCID: PMC7426096 DOI: 10.7554/elife.56428] [Citation(s) in RCA: 29] [Impact Index Per Article: 7.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/28/2020] [Accepted: 07/28/2020] [Indexed: 02/07/2023] Open
Abstract
Telomeric G-quadruplexes (G4) were long believed to form a protective structure at telomeres, preventing their extension by the ribonucleoprotein telomerase. Contrary to this belief, we have previously demonstrated that parallel-stranded conformations of telomeric G4 can be extended by human and ciliate telomerase. However, a mechanistic understanding of the interaction of telomerase with structured DNA remained elusive. Here, we use single-molecule fluorescence resonance energy transfer (smFRET) microscopy and bulk-phase enzymology to propose a mechanism for the resolution and extension of parallel G4 by telomerase. Binding is initiated by the RNA template of telomerase interacting with the G-quadruplex; nucleotide addition then proceeds to the end of the RNA template. It is only through the large conformational change of translocation following synthesis that the G-quadruplex structure is completely unfolded to a linear product. Surprisingly, parallel G4 stabilization with either small molecule ligands or by chemical modification does not always inhibit G4 unfolding and extension by telomerase. These data reveal that telomerase is a parallel G-quadruplex resolvase.
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Affiliation(s)
- Bishnu P Paudel
- Molecular Horizons and School of Chemistry and Molecular Bioscience, University of Wollongong, Wollongong, Australia.,Illawara Health and Medical Research Institute, Wollongong, Australia
| | - Aaron Lavel Moye
- Children's Medical Research Institute, University of Sydney, Westmead, Australia
| | - Hala Abou Assi
- Department of Chemistry, McGill University, Montreal, Canada
| | | | - Scott B Cohen
- Children's Medical Research Institute, University of Sydney, Westmead, Australia
| | - Jessica K Holien
- School of Science, College of Science, Engineering and Health, RMIT University, Melbourne, Australia
| | - Monica L Birrento
- Molecular Horizons and School of Chemistry and Molecular Bioscience, University of Wollongong, Wollongong, Australia.,Illawara Health and Medical Research Institute, Wollongong, Australia
| | - Siritron Samosorn
- Department of Chemistry and Center of Excellence for Innovation in Chemistry, Faculty of Science, Srinakharinwirot University, Bangkok, Thailand
| | - Kamthorn Intharapichai
- Department of Biobased Materials Science, Kyoto Institute of Technology, Matsugasaki, Sakyo-ku, Kyoto, Japan
| | | | - Marie-Paule Teulade-Fichou
- Institut Curie, PSL Research University, Orsay, France.,Université Paris Sud, Université Paris-Saclay, Orsay, France
| | - Carlos González
- Instituto de Química Física 'Rocasolano', CSIC, Madrid, Spain
| | - Jennifer L Beck
- Molecular Horizons and School of Chemistry and Molecular Bioscience, University of Wollongong, Wollongong, Australia.,Illawara Health and Medical Research Institute, Wollongong, Australia
| | - Masad J Damha
- Department of Chemistry, McGill University, Montreal, Canada
| | - Antoine M van Oijen
- Molecular Horizons and School of Chemistry and Molecular Bioscience, University of Wollongong, Wollongong, Australia.,Illawara Health and Medical Research Institute, Wollongong, Australia
| | - Tracy M Bryan
- Children's Medical Research Institute, University of Sydney, Westmead, Australia
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46
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Abstract
FRET is both a phenomenon and a spectroscopic technique, capable of measuring one geometric quantity: kappa-squared divided by the sixth power of the donor-acceptor distance. Kappa-squared is often replaced by a constant even though such a replacement may lead to serious errors. Kappaphobia, the fear of kappa or the reluctance to deal with kappa-squared adequately, is a looming presence in the FRET community. Unfortunately, this reluctance, or fear, is often tolerated, and sometimes encouraged. A decrease in kappaphobia will lead to an increase in the impact and success of FRET.
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Affiliation(s)
- B Wieb VanDerMeer
- Professor Emeritus of Physics and Biophysics, Western Kentucky University, Bowling Green, KY 42101, United States of America
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47
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Craggs TD, Sustarsic M, Plochowietz A, Mosayebi M, Kaju H, Cuthbert A, Hohlbein J, Domicevica L, Biggin PC, Doye JPK, Kapanidis AN. Substrate conformational dynamics facilitate structure-specific recognition of gapped DNA by DNA polymerase. Nucleic Acids Res 2020; 47:10788-10800. [PMID: 31544938 PMCID: PMC6846080 DOI: 10.1093/nar/gkz797] [Citation(s) in RCA: 30] [Impact Index Per Article: 7.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/15/2019] [Revised: 09/02/2019] [Accepted: 09/18/2019] [Indexed: 01/23/2023] Open
Abstract
DNA-binding proteins utilise different recognition mechanisms to locate their DNA targets; some proteins recognise specific DNA sequences, while others interact with specific DNA structures. While sequence-specific DNA binding has been studied extensively, structure-specific recognition mechanisms remain unclear. Here, we study structure-specific DNA recognition by examining the structure and dynamics of DNA polymerase I Klenow Fragment (Pol) substrates both alone and in DNA–Pol complexes. Using a docking approach based on a network of 73 distances collected using single-molecule FRET, we determined a novel solution structure of the single-nucleotide-gapped DNA–Pol binary complex. The structure resembled existing crystal structures with regards to the downstream primer-template DNA substrate, and revealed a previously unobserved sharp bend (∼120°) in the DNA substrate; this pronounced bend was present in living cells. MD simulations and single-molecule assays also revealed that 4–5 nt of downstream gap-proximal DNA are unwound in the binary complex. Further, experiments and coarse-grained modelling showed the substrate alone frequently adopts bent conformations with 1–2 nt fraying around the gap, suggesting a mechanism wherein Pol recognises a pre-bent, partially-melted conformation of gapped DNA. We propose a general mechanism for substrate recognition by structure-specific enzymes driven by protein sensing of the conformational dynamics of their DNA substrates.
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Affiliation(s)
- Timothy D Craggs
- Biological Physics Research Group, Clarendon Laboratory, Department of Physics, University of Oxford, Oxford OX1 3PU, UK
| | - Marko Sustarsic
- Biological Physics Research Group, Clarendon Laboratory, Department of Physics, University of Oxford, Oxford OX1 3PU, UK
| | - Anne Plochowietz
- Biological Physics Research Group, Clarendon Laboratory, Department of Physics, University of Oxford, Oxford OX1 3PU, UK
| | - Majid Mosayebi
- Physical and Theoretical Chemistry Laboratory, Department of Chemistry, University of Oxford, South Parks Road, Oxford OX1 3QZ, UK.,School of Physics, Institute for Research in Fundamental Sciences (IPM), Tehran 19538-33511, Iran
| | - Hendrik Kaju
- Physical and Theoretical Chemistry Laboratory, Department of Chemistry, University of Oxford, South Parks Road, Oxford OX1 3QZ, UK
| | - Andrew Cuthbert
- Biological Physics Research Group, Clarendon Laboratory, Department of Physics, University of Oxford, Oxford OX1 3PU, UK
| | - Johannes Hohlbein
- Laboratory of Biophysics, Wageningen University & Research, Wageningen 6708 WE, The Netherlands.,Microspectroscopy Research Facility Wageningen, Wageningen University & Research, Wageningen 6708 WE, The Netherlands
| | - Laura Domicevica
- Department of Biochemistry, University of Oxford, Oxford OX1 3QU, UK
| | - Philip C Biggin
- Department of Biochemistry, University of Oxford, Oxford OX1 3QU, UK
| | - Jonathan P K Doye
- Physical and Theoretical Chemistry Laboratory, Department of Chemistry, University of Oxford, South Parks Road, Oxford OX1 3QZ, UK
| | - Achillefs N Kapanidis
- Biological Physics Research Group, Clarendon Laboratory, Department of Physics, University of Oxford, Oxford OX1 3PU, UK
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48
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Calabrese AN, Schiffrin B, Watson M, Karamanos TK, Walko M, Humes JR, Horne JE, White P, Wilson AJ, Kalli AC, Tuma R, Ashcroft AE, Brockwell DJ, Radford SE. Inter-domain dynamics in the chaperone SurA and multi-site binding to its outer membrane protein clients. Nat Commun 2020; 11:2155. [PMID: 32358557 PMCID: PMC7195389 DOI: 10.1038/s41467-020-15702-1] [Citation(s) in RCA: 36] [Impact Index Per Article: 9.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/24/2019] [Accepted: 03/18/2020] [Indexed: 01/11/2023] Open
Abstract
The periplasmic chaperone SurA plays a key role in outer membrane protein (OMP) biogenesis. E. coli SurA comprises a core domain and two peptidylprolyl isomerase domains (P1 and P2), but its mechanisms of client binding and chaperone function have remained unclear. Here, we use chemical cross-linking, hydrogen-deuterium exchange mass spectrometry, single-molecule FRET and molecular dynamics simulations to map the client binding site(s) on SurA and interrogate the role of conformational dynamics in OMP recognition. We demonstrate that SurA samples an array of conformations in solution in which P2 primarily lies closer to the core/P1 domains than suggested in the SurA crystal structure. OMP binding sites are located primarily in the core domain, and OMP binding results in conformational changes between the core/P1 domains. Together, the results suggest that unfolded OMP substrates bind in a cradle formed between the SurA domains, with structural flexibility between domains assisting OMP recognition, binding and release.
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Affiliation(s)
- Antonio N Calabrese
- Astbury Centre for Structural Molecular Biology, School of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK
| | - Bob Schiffrin
- Astbury Centre for Structural Molecular Biology, School of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK
| | - Matthew Watson
- Astbury Centre for Structural Molecular Biology, School of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK
| | - Theodoros K Karamanos
- Astbury Centre for Structural Molecular Biology, School of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK
- National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, MD, 20892, USA
| | - Martin Walko
- Astbury Centre for Structural Molecular Biology, School of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK
- Astbury Centre for Structural Molecular Biology, School of Chemistry, University of Leeds, Leeds, LS2 9JT, UK
| | - Julia R Humes
- Astbury Centre for Structural Molecular Biology, School of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK
| | - Jim E Horne
- Astbury Centre for Structural Molecular Biology, School of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK
| | - Paul White
- Astbury Centre for Structural Molecular Biology, School of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK
| | - Andrew J Wilson
- Astbury Centre for Structural Molecular Biology, School of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK
- Astbury Centre for Structural Molecular Biology, School of Chemistry, University of Leeds, Leeds, LS2 9JT, UK
| | - Antreas C Kalli
- Astbury Centre for Structural Molecular Biology and School of Medicine, University of Leeds, Leeds, LS2 9JT, UK
| | - Roman Tuma
- Astbury Centre for Structural Molecular Biology, School of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK
- Faculty of Science, University of South Bohemia, Ceske Budejovice, Czech Republic
| | - Alison E Ashcroft
- Astbury Centre for Structural Molecular Biology, School of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK
| | - David J Brockwell
- Astbury Centre for Structural Molecular Biology, School of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK
| | - Sheena E Radford
- Astbury Centre for Structural Molecular Biology, School of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK.
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49
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Tsytlonok M, Hemmen K, Hamilton G, Kolimi N, Felekyan S, Seidel CAM, Tompa P, Sanabria H. Specific Conformational Dynamics and Expansion Underpin a Multi-Step Mechanism for Specific Binding of p27 with Cdk2/Cyclin A. J Mol Biol 2020; 432:2998-3017. [PMID: 32088186 PMCID: PMC7254055 DOI: 10.1016/j.jmb.2020.02.010] [Citation(s) in RCA: 17] [Impact Index Per Article: 4.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/22/2019] [Revised: 01/27/2020] [Accepted: 02/09/2020] [Indexed: 12/12/2022]
Abstract
The protein p27, a prominent regulatory protein in eukaryotes and an intrinsically disordered protein (IDP), regulates cell division by causing cell cycle arrest when bound in ternary complex with cyclin-dependent kinase (Cdk2) and cyclins (e.g., Cdk2/Cyclin A). We present an integrative study of p27 and its binding to Cdk2/Cyclin A complex by performing single-molecule multiparameter fluorescence spectroscopy, stopped-flow experiments, and molecular dynamics simulations. Our results suggest that unbound p27 adopts a compact conformation and undergoes conformational dynamics across several orders of magnitude in time (nano-to milliseconds), reflecting a multi-step mechanism for binding Cdk2/Cyclin A. Mutagenesis studies reveal that the region D1 in p27 plays a significant role in mediating the association kinetics, undergoing conformational rearrangement upon initial binding. Additionally, FRET experiments indicate an expansion of p27 throughout binding. The detected local and long-range structural dynamics suggest that p27 exhibits a limited binding surface in the unbound form, and stochastic conformational changes in D1 facilitate initial binding to Cdk2/Cyclin A complex. Furthermore, the post-kinase inhibitory domain (post-KID) region of p27 exchanges between distinct conformational ensembles: an extended regime exhibiting worm-like chain behavior, and a compact ensemble, which may protect p27 against nonspecific interactions. In summary, the binding interaction involves three steps: (i) D1 initiates binding, (ii) p27 wraps around Cdk2/Cyclin A and D2 binds, and (iii) the fully-formed fuzzy ternary complex is formed concomitantly with an extension of the post-KID region. An understanding of how the IDP nature of p27 underpins its functional interactions with Cdk2/Cyclin A provides insight into the complex binding mechanisms of IDPs and their regulatory mechanisms.
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Affiliation(s)
- Maksym Tsytlonok
- VIB-VUB Center for Structural Biology (CSB), Vrije Universiteit Brussel, Brussels, Belgium
| | - Katherina Hemmen
- Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, 40225, Düsseldorf, Germany; Rudolf Virchow Center for Experimental Biomedicine, University of Würzburg, 97078, Würzburg, Germany
| | - George Hamilton
- Department of Physics and Astronomy, Clemson University, Clemson, SC, 29634, USA
| | - Narendar Kolimi
- Department of Physics and Astronomy, Clemson University, Clemson, SC, 29634, USA
| | - Suren Felekyan
- Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, 40225, Düsseldorf, Germany
| | - Claus A M Seidel
- Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, 40225, Düsseldorf, Germany
| | - Peter Tompa
- VIB-VUB Center for Structural Biology (CSB), Vrije Universiteit Brussel, Brussels, Belgium; Institute of Enzymology, Research Centre for Natural Sciences, Budapest, Hungary
| | - Hugo Sanabria
- Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, 40225, Düsseldorf, Germany; Department of Physics and Astronomy, Clemson University, Clemson, SC, 29634, USA.
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50
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Guo H, Song S, Dai T, Sun K, Zhou G, Li M, Mann S, Dou H. Near-Infrared Fluorescent and Magnetic Resonance Dual-Imaging Coacervate Nanoprobes for Trypsin Mapping and Targeted Payload Delivery of Malignant Tumors. ACS APPLIED MATERIALS & INTERFACES 2020; 12:17302-17313. [PMID: 32212678 DOI: 10.1021/acsami.0c03433] [Citation(s) in RCA: 7] [Impact Index Per Article: 1.8] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/10/2023]
Abstract
Trypsin-responsive near-infrared fluorescent (NIRF) and magnetic resonance (MR) dual-imaging composite nanoparticle/polypeptide coacervate nanoprobes with tunable sizes, have been constructed herein via electrostatic interaction-induced self-assembly. Considering the requirements of in vivo metabolism on nanoparticle size, three coacervate nanoprobes with diameters of around 100, 200, and 300 nm were fabricated with a polydispersity of around 0.2. These coacervate nanoprobes consist of Fe3O4 magnetic nanoparticles surface-decorated with poly acrylic acid and Cy5.5-modified poly-l-lysine (PLL-g-Cy5.5) serving as MR imaging and trypsin-responsive substrate/NIRF agents, respectively. The notable fluorescence signal from PLL-g-Cy5.5 is self-quenched due to the short distances between the fluorescent Cy5.5 molecules after construction of the coacervate nanoprobes. Remarkably, coacervate nanoprobes with a diameter of around 100 nm are selectively disintegrated into fragmented segments upon the hydrolysis of PLL by trypsin, resulting in an 18-fold amplification of the NIRF intensity in comparison with the self-assembled coacervate nanoprobes in the quenched state. Moreover, the MR imaging enhancement is also related to the disintegration of the coacervate nanoprobes. Cellular experiments and in vivo studies demonstrate that the coacervate nanoprobes exhibit remarkable trypsin-sensitive NIRF and MR dual-imaging capabilities and thus have excellent potential to serve as dual-imaging nanoprobes for the efficient mapping of malignant tumors in which trypsin is often overexpressed. In consideration of their excellent capability to enrich charged molecules, the coacervate nanoprobes provide a conceptually novel and promising platform toward in vivo trypsin mapping and controlled delivery of targeted payloads.
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Affiliation(s)
- Heze Guo
- The State Key Laboratory of Metal Matrix Composites, School of Materials Science and Engineering, Shanghai Jiao Tong University, Shanghai 200240, P. R. China
| | - Sheng Song
- The State Key Laboratory of Metal Matrix Composites, School of Materials Science and Engineering, Shanghai Jiao Tong University, Shanghai 200240, P. R. China
| | - Tingting Dai
- Department of Plastic and Reconstructive Surgery, Shanghai Ninth People's Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai Key Laboratory of Tissue Engineering National Tissue Engineering Centre of China, Shanghai 200011, P. R. China
| | - Kang Sun
- The State Key Laboratory of Metal Matrix Composites, School of Materials Science and Engineering, Shanghai Jiao Tong University, Shanghai 200240, P. R. China
| | - Guangdong Zhou
- Department of Plastic and Reconstructive Surgery, Shanghai Ninth People's Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai Key Laboratory of Tissue Engineering National Tissue Engineering Centre of China, Shanghai 200011, P. R. China
| | - Mei Li
- Centre for Protolife Research and Centre for Organized Matter Chemistry, School of Chemistry, University of Bristol, Bristol BS8 1TS, U.K
| | - Stephen Mann
- Centre for Protolife Research and Centre for Organized Matter Chemistry, School of Chemistry, University of Bristol, Bristol BS8 1TS, U.K
| | - Hongjing Dou
- The State Key Laboratory of Metal Matrix Composites, School of Materials Science and Engineering, Shanghai Jiao Tong University, Shanghai 200240, P. R. China
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