1
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Ando T, Fukuda S, Ngo KX, Flechsig H. High-Speed Atomic Force Microscopy for Filming Protein Molecules in Dynamic Action. Annu Rev Biophys 2024; 53:19-39. [PMID: 38060998 DOI: 10.1146/annurev-biophys-030722-113353] [Citation(s) in RCA: 2] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 07/18/2024]
Abstract
Structural biology is currently undergoing a transformation into dynamic structural biology, which reveals the dynamic structure of proteins during their functional activity to better elucidate how they function. Among the various approaches in dynamic structural biology, high-speed atomic force microscopy (HS-AFM) is unique in the ability to film individual molecules in dynamic action, although only topographical information is acquirable. This review provides a guide to the use of HS-AFM for biomolecular imaging and showcases several examples, as well as providing information on up-to-date progress in HS-AFM technology. Finally, we discuss the future prospects of HS-AFM in the context of dynamic structural biology in the upcoming era.
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Affiliation(s)
- Toshio Ando
- Nano Life Science Institute (WPI-NanoLSI), Kanazawa University, Kanazawa, Japan;
| | - Shingo Fukuda
- Nano Life Science Institute (WPI-NanoLSI), Kanazawa University, Kanazawa, Japan;
| | - Kien X Ngo
- Nano Life Science Institute (WPI-NanoLSI), Kanazawa University, Kanazawa, Japan;
| | - Holger Flechsig
- Nano Life Science Institute (WPI-NanoLSI), Kanazawa University, Kanazawa, Japan;
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2
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Bussoletti M, Gallo M, Bottacchiari M, Abbondanza D, Casciola CM. Mesoscopic elasticity controls dynamin-driven fission of lipid tubules. Sci Rep 2024; 14:14003. [PMID: 38890460 PMCID: PMC11189461 DOI: 10.1038/s41598-024-64685-2] [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: 03/09/2024] [Accepted: 06/12/2024] [Indexed: 06/20/2024] Open
Abstract
Mesoscale physics bridges the gap between the microscopic degrees of freedom of a system and its large-scale continuous behavior and highlights the role of a few key quantities in complex and multiscale phenomena, like dynamin-driven fission of lipid membranes. The dynamin protein wraps the neck formed during clathrin-mediated endocytosis, for instance, and constricts it until severing occurs. Although ubiquitous and fundamental for life, the cooperation between the GTP-consuming conformational changes within the protein and the full-scale response of the underlying lipid substrate is yet to be unraveled. In this work, we build an effective mesoscopic model from constriction to fission of lipid tubules based on continuum membrane elasticity and implicitly accounting for ratchet-like power strokes of dynamins. Localization of the fission event, the overall geometry, and the energy expenditure we predict comply with the major experimental findings. This bolsters the idea that a continuous picture emerges soon enough to relate dynamin polymerization length and membrane rigidity and tension with the optimal pathway to fission. We therefore suggest that dynamins found in in vivo processes may optimize their structure accordingly. Ultimately, we shed light on real-time conductance measurements available in literature and predict the fission time dependency on elastic parameters.
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Affiliation(s)
- Marco Bussoletti
- Department of Mechanical and Aerospace Engineering, Sapienza University of Rome, Rome, Italy
| | - Mirko Gallo
- Department of Mechanical and Aerospace Engineering, Sapienza University of Rome, Rome, Italy
| | - Matteo Bottacchiari
- Department of Mechanical and Aerospace Engineering, Sapienza University of Rome, Rome, Italy
- Department of Basic and Applied Sciences for Engineering, Sapienza University of Rome, Rome, Italy
| | - Dario Abbondanza
- Department of Mechanical and Aerospace Engineering, Sapienza University of Rome, Rome, Italy
| | - Carlo Massimo Casciola
- Department of Mechanical and Aerospace Engineering, Sapienza University of Rome, Rome, Italy.
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3
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Kim GW, Ha JW. Direct Observation of In-Focus Plasmonic Cargos via Breaking Angular Degeneracy in Differential Interference Contrast Microscopy. JACS AU 2023; 3:3436-3445. [PMID: 38155657 PMCID: PMC10751767 DOI: 10.1021/jacsau.3c00594] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Download PDF] [Figures] [Subscribe] [Scholar Register] [Received: 10/03/2023] [Revised: 11/19/2023] [Accepted: 11/22/2023] [Indexed: 12/30/2023]
Abstract
Breaking the angular degeneracy arising from the 2-fold optical symmetry of plasmonic anisotropic nanoprobes is critical in biological studies. In this study, we propose differential interference contrast (DIC) microscopy-based focused orientation and position imaging (dFOPI) to break the angular degeneracy of single gold nanorods (AuNRs). Single in-focus AuNRs (39 nm × 123 nm) within a spherical mesoporous silica shell were characterized with high throughput and produced distinct doughnut-shaped DIC image patterns featuring two lobes in the peripheral region, attributed to the scattering contribution of the AuNRs with large scattering cross sections. Interestingly, rotation of the lobes was observed in the focal plane for a large AuNR (>100 nm) tilted by more than ∼20° from the horizontal plane as the rotational stage was moved by 10° in a rotational study. From the rotation-dependent characteristic patterns, we directly visualized counterclockwise/clockwise rotations without the angular degeneracy at the localized surface plasmon resonance wavelength. Therefore, our dFOPI method can be applied for in vivo studies of important biological systems. To validate this claim, we tracked the three-dimensional rotational behavior of transferrin-modified in-focus AuNRs during clathrin-mediated endocytosis in real time without sacrificing the temporal and spatial resolution. In the invagination and scission stage, one or two directed twist motions of the AuNR cargos detached the AuNR-containing vesicles from the cell membrane. Furthermore, the dFOPI method directly visualized and revealed the right-handed twisting action along the dynamin helix in dynamin-catalyzed fission in live cells.
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Affiliation(s)
- Geun Wan Kim
- Department
of Chemistry, University of Ulsan, 93 Daehak-ro, Nam-gu, Ulsan 44610, South Korea
| | - Ji Won Ha
- Department
of Chemistry, University of Ulsan, 93 Daehak-ro, Nam-gu, Ulsan 44610, South Korea
- Energy
Harvest-Storage Research Center (EHSRC), University of Ulsan, 93 Daehak-ro, Nam-gu, Ulsan 44610, South Korea
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4
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Noshiro D, Noda NN. Immobilization of lipid nanorods onto two-dimensional crystals of protein tamavidin 2 for high-speed atomic force microscopy. STAR Protoc 2023; 4:102633. [PMID: 38043055 PMCID: PMC10701432 DOI: 10.1016/j.xpro.2023.102633] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/27/2023] [Revised: 09/08/2023] [Accepted: 09/21/2023] [Indexed: 12/05/2023] Open
Abstract
High-speed atomic force microscopy is a technique that allows real-time observation of biomolecules and biological phenomena reconstituted on a substrate. Here, we present a protocol for immobilizing lipid nanorods onto two-dimensional crystals of biotin-binding protein tamavidin 2. We describe steps for the preparation of tamavidin 2 protein, lipid nanorods, and two-dimensional crystals of tamavidin 2 formed on mica. Immobilized lipid nanorods are one of the useful tools for observation of specific proteins in action. For complete details on the use and execution of this protocol, please refer to Fukuda et al. (2023).1.
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Affiliation(s)
- Daisuke Noshiro
- Institute for Genetic Medicine, Hokkaido University, Sapporo, Hokkaido 060-0815, Japan
| | - Nobuo N Noda
- Institute for Genetic Medicine, Hokkaido University, Sapporo, Hokkaido 060-0815, Japan; Institute of Microbial Chemistry (BIKAKEN), Shinagawa-ku, Tokyo 141-0021, Japan.
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5
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Yang J, Jahnke K, Xin L, Jing X, Zhan P, Peil A, Griffo A, Škugor M, Yang D, Fan S, Göpfrich K, Yan H, Wang P, Liu N. Modulating Lipid Membrane Morphology by Dynamic DNA Origami Networks. NANO LETTERS 2023. [PMID: 37440701 DOI: 10.1021/acs.nanolett.3c00750] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 07/15/2023]
Abstract
Membrane morphology and its dynamic adaptation regulate many cellular functions, which are often mediated by membrane proteins. Advances in DNA nanotechnology have enabled the realization of various protein-inspired structures and functions with precise control at the nanometer level, suggesting a viable tool to artificially engineer membrane morphology. In this work, we demonstrate a DNA origami cross (DOC) structure that can be anchored onto giant unilamellar vesicles (GUVs) and subsequently polymerized into micrometer-scale reconfigurable one-dimensional (1D) chains or two-dimensional (2D) lattices. Such DNA origami-based networks can be switched between left-handed (LH) and right-handed (RH) conformations by DNA fuels and exhibit potent efficacy in remodeling the membrane curvatures of GUVs. This work sheds light on designing hierarchically assembled dynamic DNA systems for the programmable modulation of synthetic cells for useful applications.
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Affiliation(s)
- Juanjuan Yang
- Institute of Molecular Medicine, Department of Laboratory Medicine, Shanghai Key Laboratory for Nucleic Acid Chemistry and Nanomedicine, Renji Hospital, School of Medicine, Shanghai Jiao Tong University Shanghai 200127, People's Republic of China
- 2. Physics Institute, University of Stuttgart, Pfaffenwaldring 57, 70569, Stuttgart, Germany
| | - Kevin Jahnke
- Biophysical Engineering Group, Max Planck Institute for Medical Research Heidelberg, Jahnstr. 29, 69120 Heidelberg, Germany
| | - Ling Xin
- 2. Physics Institute, University of Stuttgart, Pfaffenwaldring 57, 70569, Stuttgart, Germany
| | - Xinxin Jing
- 2. Physics Institute, University of Stuttgart, Pfaffenwaldring 57, 70569, Stuttgart, Germany
| | - Pengfei Zhan
- 2. Physics Institute, University of Stuttgart, Pfaffenwaldring 57, 70569, Stuttgart, Germany
| | - Andreas Peil
- 2. Physics Institute, University of Stuttgart, Pfaffenwaldring 57, 70569, Stuttgart, Germany
| | - Alessandra Griffo
- Biophysical Engineering Group, Max Planck Institute for Medical Research Heidelberg, Jahnstr. 29, 69120 Heidelberg, Germany
| | - Marko Škugor
- 2. Physics Institute, University of Stuttgart, Pfaffenwaldring 57, 70569, Stuttgart, Germany
| | - Donglei Yang
- Institute of Molecular Medicine, Department of Laboratory Medicine, Shanghai Key Laboratory for Nucleic Acid Chemistry and Nanomedicine, Renji Hospital, School of Medicine, Shanghai Jiao Tong University Shanghai 200127, People's Republic of China
| | - Sisi Fan
- 2. Physics Institute, University of Stuttgart, Pfaffenwaldring 57, 70569, Stuttgart, Germany
| | - Kerstin Göpfrich
- Biophysical Engineering Group, Max Planck Institute for Medical Research Heidelberg, Jahnstr. 29, 69120 Heidelberg, Germany
- Center for Molecular Biology of Heidelberg University (ZMBH), Im Neuenheime Feld 329, 69120 Heidelberg, Germany
| | - Hao Yan
- School of Molecular Sciences and Center for Molecular Design and Biomimetics at Biodesign Institute, Arizona State University, Tempe, Arizona 85287, United States
| | - Pengfei Wang
- Institute of Molecular Medicine, Department of Laboratory Medicine, Shanghai Key Laboratory for Nucleic Acid Chemistry and Nanomedicine, Renji Hospital, School of Medicine, Shanghai Jiao Tong University Shanghai 200127, People's Republic of China
| | - Na Liu
- 2. Physics Institute, University of Stuttgart, Pfaffenwaldring 57, 70569, Stuttgart, Germany
- Max Planck Institute for Solid State Research, Heisenbergstrasse 1, D-70569 Stuttgart, Germany
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6
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Fukuda T, Furukawa K, Maruyama T, Yamashita SI, Noshiro D, Song C, Ogasawara Y, Okuyama K, Alam JM, Hayatsu M, Saigusa T, Inoue K, Ikeda K, Takai A, Chen L, Lahiri V, Okada Y, Shibata S, Murata K, Klionsky DJ, Noda NN, Kanki T. The mitochondrial intermembrane space protein mitofissin drives mitochondrial fission required for mitophagy. Mol Cell 2023; 83:2045-2058.e9. [PMID: 37192628 PMCID: PMC10330776 DOI: 10.1016/j.molcel.2023.04.022] [Citation(s) in RCA: 21] [Impact Index Per Article: 21.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/08/2022] [Revised: 01/30/2023] [Accepted: 04/21/2023] [Indexed: 05/18/2023]
Abstract
Mitophagy plays an important role in mitochondrial homeostasis by selective degradation of mitochondria. During mitophagy, mitochondria should be fragmented to allow engulfment within autophagosomes, whose capacity is exceeded by the typical mitochondria mass. However, the known mitochondrial fission factors, dynamin-related proteins Dnm1 in yeasts and DNM1L/Drp1 in mammals, are dispensable for mitophagy. Here, we identify Atg44 as a mitochondrial fission factor that is essential for mitophagy in yeasts, and we therefore term Atg44 and its orthologous proteins mitofissin. In mitofissin-deficient cells, a part of the mitochondria is recognized by the mitophagy machinery as cargo but cannot be enwrapped by the autophagosome precursor, the phagophore, due to a lack of mitochondrial fission. Furthermore, we show that mitofissin directly binds to lipid membranes and brings about lipid membrane fragility to facilitate membrane fission. Taken together, we propose that mitofissin acts directly on lipid membranes to drive mitochondrial fission required for mitophagy.
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Affiliation(s)
- Tomoyuki Fukuda
- Department of Cellular Physiology, Niigata University Graduate School of Medical and Dental Sciences, Niigata 951-8510, Japan
| | - Kentaro Furukawa
- Department of Cellular Physiology, Niigata University Graduate School of Medical and Dental Sciences, Niigata 951-8510, Japan
| | - Tatsuro Maruyama
- Institute of Microbial Chemistry (BIKAKEN), Shinagawa-ku, Tokyo 141-0021, Japan
| | - Shun-Ichi Yamashita
- Department of Cellular Physiology, Niigata University Graduate School of Medical and Dental Sciences, Niigata 951-8510, Japan
| | - Daisuke Noshiro
- Institute of Microbial Chemistry (BIKAKEN), Shinagawa-ku, Tokyo 141-0021, Japan; Institute for Genetic Medicine, Hokkaido University, Sapporo, Hokkaido 060-0815, Japan
| | - Chihong Song
- National Institute for Physiological Sciences (NIPS), National Institutes of Natural Sciences (NINS), Okazaki, Aichi 444-8585, Japan; Exploratory Research Center on Life and Living Systems (ExCELLS), National Institutes of Natural Sciences (NINS), Okazaki, Aichi 444-8585, Japan
| | - Yuta Ogasawara
- Institute of Microbial Chemistry (BIKAKEN), Shinagawa-ku, Tokyo 141-0021, Japan; Institute for Genetic Medicine, Hokkaido University, Sapporo, Hokkaido 060-0815, Japan
| | - Kentaro Okuyama
- Division of Microscopic Anatomy, Niigata University Graduate School of Medical and Dental Sciences, Niigata 951-8510, Japan
| | - Jahangir Md Alam
- Institute of Microbial Chemistry (BIKAKEN), Shinagawa-ku, Tokyo 141-0021, Japan
| | - Manabu Hayatsu
- Division of Microscopic Anatomy, Niigata University Graduate School of Medical and Dental Sciences, Niigata 951-8510, Japan
| | - Tetsu Saigusa
- Department of Cellular Physiology, Niigata University Graduate School of Medical and Dental Sciences, Niigata 951-8510, Japan
| | - Keiichi Inoue
- Department of Cellular Physiology, Niigata University Graduate School of Medical and Dental Sciences, Niigata 951-8510, Japan
| | - Kazuho Ikeda
- Department of Cell Biology, Graduate School of Medicine, The University of Tokyo, Tokyo 113-0033, Japan; Laboratory for Cell Polarity Regulation, RIKEN Center for Biosystems Dynamics Research (BDR), Osaka 565-0874, Japan
| | - Akira Takai
- Department of Cell Biology, Graduate School of Medicine, The University of Tokyo, Tokyo 113-0033, Japan; Laboratory for Cell Polarity Regulation, RIKEN Center for Biosystems Dynamics Research (BDR), Osaka 565-0874, Japan
| | - Lin Chen
- National Institute for Physiological Sciences (NIPS), National Institutes of Natural Sciences (NINS), Okazaki, Aichi 444-8585, Japan; Exploratory Research Center on Life and Living Systems (ExCELLS), National Institutes of Natural Sciences (NINS), Okazaki, Aichi 444-8585, Japan
| | - Vikramjit Lahiri
- Life Sciences Institute and Department of Molecular, Cellular and Developmental Biology, University of Michigan, Ann Arbor, MI 48109, USA
| | - Yasushi Okada
- Department of Cell Biology, Graduate School of Medicine, The University of Tokyo, Tokyo 113-0033, Japan; Laboratory for Cell Polarity Regulation, RIKEN Center for Biosystems Dynamics Research (BDR), Osaka 565-0874, Japan; Department of Physics, Graduate School of Science, The University of Tokyo, Tokyo 113-0033, Japan; Universal Biology Institute (UBI) and International Research Center for Neurointelligence (WPI-IRCN), The University of Tokyo, Tokyo 113-0033, Japan
| | - Shinsuke Shibata
- Division of Microscopic Anatomy, Niigata University Graduate School of Medical and Dental Sciences, Niigata 951-8510, Japan
| | - Kazuyoshi Murata
- National Institute for Physiological Sciences (NIPS), National Institutes of Natural Sciences (NINS), Okazaki, Aichi 444-8585, Japan; Exploratory Research Center on Life and Living Systems (ExCELLS), National Institutes of Natural Sciences (NINS), Okazaki, Aichi 444-8585, Japan
| | - Daniel J Klionsky
- Life Sciences Institute and Department of Molecular, Cellular and Developmental Biology, University of Michigan, Ann Arbor, MI 48109, USA
| | - Nobuo N Noda
- Institute of Microbial Chemistry (BIKAKEN), Shinagawa-ku, Tokyo 141-0021, Japan; Institute for Genetic Medicine, Hokkaido University, Sapporo, Hokkaido 060-0815, Japan.
| | - Tomotake Kanki
- Department of Cellular Physiology, Niigata University Graduate School of Medical and Dental Sciences, Niigata 951-8510, Japan.
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7
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Jukic N, Perrino AP, Redondo-Morata L, Scheuring S. Structure and dynamics of ESCRT-III membrane remodeling proteins by high-speed atomic force microscopy. J Biol Chem 2023; 299:104575. [PMID: 36870686 PMCID: PMC10074808 DOI: 10.1016/j.jbc.2023.104575] [Citation(s) in RCA: 2] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/29/2022] [Revised: 02/20/2023] [Accepted: 02/23/2023] [Indexed: 03/06/2023] Open
Abstract
Endosomal Sorting Complex Required for Transport (ESCRT) proteins assemble on the cytoplasmic leaflet of membranes and remodel them. ESCRT is involved in biological processes where membranes are bent away from the cytosol, constricted, and finally severed, such as in multi-vesicular body formation (in the endosomal pathway for protein sorting) or abscission during cell division. The ESCRT system is hijacked by enveloped viruses to allow buds of nascent virions to be constricted, severed and released. ESCRT-III proteins, the most downstream components of the ESCRT system, are monomeric and cytosolic in their autoinhibited conformation. They share a common architecture, a four-helix bundle with a fifth helix that interacts with this bundle to prevent polymerizing. Upon binding to negatively charged membranes, the ESCRT-III components adopt an activated state that allows them to polymerize into filaments and spirals, and to interact with the AAA-ATPase Vps4 for polymer remodeling. ESCRT-III has been studied with electron microscopy (EM) and fluorescence microscopy (FM); these methods provided invaluable information about ESCRT assembly structures or their dynamics, respectively, but neither approach provides detailed insights into both aspects simultaneously. High-speed atomic force microscopy (HS-AFM) has overcome this shortcoming, providing movies at high spatio-temporal resolution of biomolecular processes, significantly increasing our understanding of ESCRT-III structure and dynamics. Here, we review the contributions of HS-AFM in the analysis of ESCRT-III, focusing on recent developments of non-planar and deformable HS-AFM supports. We divide the HS-AFM observations into four sequential steps in the ESCRT-III lifecycle: 1) polymerization, 2) morphology, 3) dynamics, and 4) depolymerization.
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Affiliation(s)
- Nebojsa Jukic
- Weill Cornell Medicine, Physiology, Biophysics and Systems Biology Graduate Program, New York, NY 10065, USA
| | - Alma P Perrino
- Weill Cornell Medicine, Department of Anesthesiology, 1300 York Avenue, New York, NY 10065, USA
| | - Lorena Redondo-Morata
- Université de Lille, CNRS, Inserm, CHU Lille, Institut Pasteur de Lille, U1019-UMR9017-CIIL-Centre d'Infection et d'Immunité de Lille, F-59000 Lille, France
| | - Simon Scheuring
- Weill Cornell Medicine, Department of Anesthesiology, 1300 York Avenue, New York, NY 10065, USA; Weill Cornell Medicine, Department of Physiology and Biophysics, 1300 York Avenue, New York, NY 10065, USA; Kavli Institute at Cornell for Nanoscale Science, Cornell University, Ithaca, New York, NY 14853, USA.
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8
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Gopaldass N, De Leo MG, Courtellemont T, Mercier V, Bissig C, Roux A, Mayer A. Retromer oligomerization drives SNX-BAR coat assembly and membrane constriction. EMBO J 2023; 42:e112287. [PMID: 36644906 PMCID: PMC9841331 DOI: 10.15252/embj.2022112287] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/04/2022] [Revised: 11/02/2022] [Accepted: 11/02/2022] [Indexed: 01/17/2023] Open
Abstract
Proteins exit from endosomes through tubular carriers coated by retromer, a complex that impacts cellular signaling, lysosomal biogenesis and numerous diseases. The coat must overcome membrane tension to form tubules. We explored the dynamics and driving force of this process by reconstituting coat formation with yeast retromer and the BAR-domain sorting nexins Vps5 and Vps17 on oriented synthetic lipid tubules. This coat oligomerizes bidirectionally, forming a static tubular structure that does not exchange subunits. High concentrations of sorting nexins alone constrict membrane tubes to an invariant radius of 19 nm. At lower concentrations, oligomers of retromer must bind and interconnect the sorting nexins to drive constriction. Constricting less curved membranes into tubes, which requires more energy, coincides with an increased surface density of retromer on the sorting nexin layer. Retromer-mediated crosslinking of sorting nexins at variable densities may thus tune the energy that the coat can generate to deform the membrane. In line with this, genetic ablation of retromer oligomerization impairs endosomal protein exit in yeast and human cells.
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Affiliation(s)
- Navin Gopaldass
- Department of ImmunobiologyUniversity of LausanneEpalingesSwitzerland
| | | | | | - Vincent Mercier
- Department of BiochemistryUniversity of GenevaGenevaSwitzerland
| | - Christin Bissig
- Department of ImmunobiologyUniversity of LausanneEpalingesSwitzerland
| | - Aurélien Roux
- Department of BiochemistryUniversity of GenevaGenevaSwitzerland
- Swiss National Centre for Competence in Research Program Chemical BiologyGenevaSwitzerland
| | - Andreas Mayer
- Department of ImmunobiologyUniversity of LausanneEpalingesSwitzerland
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9
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Imoto Y, Raychaudhuri S, Ma Y, Fenske P, Sandoval E, Itoh K, Blumrich EM, Matsubayashi HT, Mamer L, Zarebidaki F, Söhl-Kielczynski B, Trimbuch T, Nayak S, Iwasa JH, Liu J, Wu B, Ha T, Inoue T, Jorgensen EM, Cousin MA, Rosenmund C, Watanabe S. Dynamin is primed at endocytic sites for ultrafast endocytosis. Neuron 2022; 110:2815-2835.e13. [PMID: 35809574 PMCID: PMC9464723 DOI: 10.1016/j.neuron.2022.06.010] [Citation(s) in RCA: 29] [Impact Index Per Article: 14.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/19/2021] [Revised: 03/24/2022] [Accepted: 06/09/2022] [Indexed: 02/06/2023]
Abstract
Dynamin mediates fission of vesicles from the plasma membrane during endocytosis. Typically, dynamin is recruited from the cytosol to endocytic sites, requiring seconds to tens of seconds. However, ultrafast endocytosis in neurons internalizes vesicles as quickly as 50 ms during synaptic vesicle recycling. Here, we demonstrate that Dynamin 1 is pre-recruited to endocytic sites for ultrafast endocytosis. Specifically, Dynamin 1xA, a splice variant of Dynamin 1, interacts with Syndapin 1 to form molecular condensates on the plasma membrane. Single-particle tracking of Dynamin 1xA molecules confirms the liquid-like property of condensates in vivo. When Dynamin 1xA is mutated to disrupt its interaction with Syndapin 1, the condensates do not form, and consequently, ultrafast endocytosis slows down by 100-fold. Mechanistically, Syndapin 1 acts as an adaptor by binding the plasma membrane and stores Dynamin 1xA at endocytic sites. This cache bypasses the recruitment step and accelerates endocytosis at synapses.
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Affiliation(s)
- Yuuta Imoto
- Department of Cell Biology, Johns Hopkins University, Baltimore, MD 21205, USA.
| | - Sumana Raychaudhuri
- Department of Cell Biology, Johns Hopkins University, Baltimore, MD 21205, USA
| | - Ye Ma
- Department of Biomedical Engineering, Johns Hopkins University, Baltimore, MD 21205, USA
| | - Pascal Fenske
- Institute of Neurophysiology, Charité Universitätsmedizin Berlin, Berlin, Germany
| | - Eduardo Sandoval
- Department of Cell Biology, Johns Hopkins University, Baltimore, MD 21205, USA
| | - Kie Itoh
- Department of Cell Biology, Johns Hopkins University, Baltimore, MD 21205, USA
| | - Eva-Maria Blumrich
- Centre for Discovery Brain Sciences, University of Edinburgh, Edinburgh, Scotland EH8 9XD, UK; The Muir Maxwell Epilepsy Centre, University of Edinburgh, Edinburgh, Scotland EH8 9XD, UK; Simons Initiatives for the Developing Brain, University of Edinburgh, Edinburgh, Scotland EH8 9XD, UK
| | - Hideaki T Matsubayashi
- Department of Cell Biology, Johns Hopkins University, Baltimore, MD 21205, USA; The Center for Cell Dynamics, Johns Hopkins University, Baltimore, MD 21205, USA
| | - Lauren Mamer
- Institute of Neurophysiology, Charité Universitätsmedizin Berlin, Berlin, Germany
| | - Fereshteh Zarebidaki
- Institute of Neurophysiology, Charité Universitätsmedizin Berlin, Berlin, Germany
| | | | - Thorsten Trimbuch
- Institute of Neurophysiology, Charité Universitätsmedizin Berlin, Berlin, Germany
| | - Shraddha Nayak
- Department of Biochemistry, University of Utah, Salt Lake City, UT 84112-0840, USA
| | - Janet H Iwasa
- Department of Biochemistry, University of Utah, Salt Lake City, UT 84112-0840, USA
| | - Jian Liu
- Department of Cell Biology, Johns Hopkins University, Baltimore, MD 21205, USA; Centre for Discovery Brain Sciences, University of Edinburgh, Edinburgh, Scotland EH8 9XD, UK
| | - Bin Wu
- The Center for Cell Dynamics, Johns Hopkins University, Baltimore, MD 21205, USA; Solomon H. Snyder Department of Neuroscience, Johns Hopkins University, Baltimore, MD 21205, USA; Department of Biophysics and Biophysical Chemistry, Johns Hopkins University, Baltimore, MD 21205, USA
| | - Taekjip Ha
- Department of Biomedical Engineering, Johns Hopkins University, Baltimore, MD 21205, USA; Department of Biophysics and Biophysical Chemistry, Johns Hopkins University, Baltimore, MD 21205, USA; Howard Hughes Medical Institute, Baltimore, MD 21205, USA
| | - Takanari Inoue
- Department of Cell Biology, Johns Hopkins University, Baltimore, MD 21205, USA; The Center for Cell Dynamics, Johns Hopkins University, Baltimore, MD 21205, USA
| | - Erik M Jorgensen
- HHMI, Department of Biology, University of Utah, Salt Lake City, UT 84112-0840, USA
| | - Michael A Cousin
- Centre for Discovery Brain Sciences, University of Edinburgh, Edinburgh, Scotland EH8 9XD, UK; The Muir Maxwell Epilepsy Centre, University of Edinburgh, Edinburgh, Scotland EH8 9XD, UK; Simons Initiatives for the Developing Brain, University of Edinburgh, Edinburgh, Scotland EH8 9XD, UK
| | - Christian Rosenmund
- Institute of Neurophysiology, Charité Universitätsmedizin Berlin, Berlin, Germany.
| | - Shigeki Watanabe
- Department of Cell Biology, Johns Hopkins University, Baltimore, MD 21205, USA; The Center for Cell Dynamics, Johns Hopkins University, Baltimore, MD 21205, USA; Solomon H. Snyder Department of Neuroscience, Johns Hopkins University, Baltimore, MD 21205, USA.
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10
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Fujise K, Noguchi S, Takeda T. Centronuclear Myopathy Caused by Defective Membrane Remodelling of Dynamin 2 and BIN1 Variants. Int J Mol Sci 2022; 23:ijms23116274. [PMID: 35682949 PMCID: PMC9181712 DOI: 10.3390/ijms23116274] [Citation(s) in RCA: 10] [Impact Index Per Article: 5.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/09/2022] [Revised: 05/31/2022] [Accepted: 06/01/2022] [Indexed: 02/01/2023] Open
Abstract
Centronuclear myopathy (CNM) is a congenital myopathy characterised by centralised nuclei in skeletal myofibers. T-tubules, sarcolemmal invaginations required for excitation-contraction coupling, are disorganised in the skeletal muscles of CNM patients. Previous studies showed that various endocytic proteins are involved in T-tubule biogenesis and their dysfunction is tightly associated with CNM pathogenesis. DNM2 and BIN1 are two causative genes for CNM that encode essential membrane remodelling proteins in endocytosis, dynamin 2 and BIN1, respectively. In this review, we overview the functions of dynamin 2 and BIN1 in T-tubule biogenesis and discuss how their dysfunction in membrane remodelling leads to CNM pathogenesis.
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Affiliation(s)
- Kenshiro Fujise
- Departments of Neuroscience and Cell Biology, Howard Hughes Medical Institute, Yale University School of Medicine, New Haven, CT 06520-8001, USA;
| | - Satoru Noguchi
- National Institute of Neuroscience, National Center of Neurology and Psychiatry (NCNP), Tokyo 187-8502, Japan;
| | - Tetsuya Takeda
- Department of Biochemistry, Faculty of Medicine, Dentistry and Pharmaceutical Sciences, Okayama University, Shikata-cho 2-5-1, Kita-ku, Okayama 700-8558, Japan
- Correspondence: ; Tel.: +81-86-235-7125; Fax: +81-86-235-7126
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11
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Nie J, Tian F, Zheng B, Wang Z, Zheng P. Exploration of Metal-Ligand Coordination Bonds in Proteins by Single-molecule Force Spectroscopy. CHEM LETT 2021. [DOI: 10.1246/cl.210307] [Citation(s) in RCA: 6] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/22/2022]
Affiliation(s)
- Jingyuan Nie
- State Key Laboratory of Coordination Chemistry, Chemistry and Biomedicine Innovation Center (ChemBIC), School of Chemistry and Chemical Engineering, Nanjing University, Nanjing, Jiangsu 210023, P. R. China
| | - Fang Tian
- State Key Laboratory of Coordination Chemistry, Chemistry and Biomedicine Innovation Center (ChemBIC), School of Chemistry and Chemical Engineering, Nanjing University, Nanjing, Jiangsu 210023, P. R. China
| | - Bin Zheng
- State Key Laboratory of Coordination Chemistry, Chemistry and Biomedicine Innovation Center (ChemBIC), School of Chemistry and Chemical Engineering, Nanjing University, Nanjing, Jiangsu 210023, P. R. China
| | - Ziyi Wang
- State Key Laboratory of Coordination Chemistry, Chemistry and Biomedicine Innovation Center (ChemBIC), School of Chemistry and Chemical Engineering, Nanjing University, Nanjing, Jiangsu 210023, P. R. China
| | - Peng Zheng
- State Key Laboratory of Coordination Chemistry, Chemistry and Biomedicine Innovation Center (ChemBIC), School of Chemistry and Chemical Engineering, Nanjing University, Nanjing, Jiangsu 210023, P. R. China
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12
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Tagiltsev G, Haselwandter CA, Scheuring S. Nanodissected elastically loaded clathrin lattices relax to increased curvature. SCIENCE ADVANCES 2021; 7:7/33/eabg9934. [PMID: 34389539 PMCID: PMC8363152 DOI: 10.1126/sciadv.abg9934] [Citation(s) in RCA: 14] [Impact Index Per Article: 4.7] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Figures] [Subscribe] [Scholar Register] [Received: 02/08/2021] [Accepted: 06/24/2021] [Indexed: 06/13/2023]
Abstract
Clathrin-mediated endocytosis (CME) is the major endocytosis pathway for the specific internalization of large compounds, growth factors, and receptors. Formation of internalized vesicles from the flat plasma membrane is accompanied by maturation of cytoplasmic clathrin coats. How clathrin coats mature and the mechanistic role of clathrin coats are still largely unknown. Maturation models proposed clathrin coats to mature at constant radius or constant area, driven by molecular actions or elastic energy. Here, combining high-speed atomic force microscopy (HS-AFM) imaging, HS-AFM nanodissection, and elasticity theory, we show that clathrin lattices deviating from the intrinsic curvature of clathrin form elastically loaded assemblies. Upon nanodissection of the clathrin network, the stored elastic energy in these lattices drives lattice relaxation to accommodate an ideal area-curvature ratio toward the formation of closed clathrin-coated vesicles. Our work supports that the release of elastic energy stored in curvature-frustrated clathrin lattices could play a major role in CME.
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Affiliation(s)
- Grigory Tagiltsev
- Department of Anesthesiology, Weill Cornell Medicine, 1300 York Avenue, New York, NY 10065, USA
- Department of Physiology and Biophysics, Weill Cornell Medicine, 1300 York Avenue, New York, NY 10065, USA
| | - Christoph A Haselwandter
- Department of Physics and Astronomy, University of Southern California, Los Angeles, CA 90089, USA
- Department of Quantitative and Computational Biology, University of Southern California, Los Angeles, CA 90089, USA
| | - Simon Scheuring
- Department of Anesthesiology, Weill Cornell Medicine, 1300 York Avenue, New York, NY 10065, USA.
- Department of Physiology and Biophysics, Weill Cornell Medicine, 1300 York Avenue, New York, NY 10065, USA
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13
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Ganichkin OM, Vancraenenbroeck R, Rosenblum G, Hofmann H, Mikhailov AS, Daumke O, Noel JK. Quantification and demonstration of the collective constriction-by-ratchet mechanism in the dynamin molecular motor. Proc Natl Acad Sci U S A 2021; 118:e2101144118. [PMID: 34244431 PMCID: PMC8285958 DOI: 10.1073/pnas.2101144118] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/27/2022] Open
Abstract
Dynamin oligomerizes into helical filaments on tubular membrane templates and, through constriction, cleaves them in a GTPase-driven way. Structural observations of GTP-dependent cross-bridges between neighboring filament turns have led to the suggestion that dynamin operates as a molecular ratchet motor. However, the proof of such mechanism remains absent. Particularly, it is not known whether a powerful enough stroke is produced and how the motor modules would cooperate in the constriction process. Here, we characterized the dynamin motor modules by single-molecule Förster resonance energy transfer (smFRET) and found strong nucleotide-dependent conformational preferences. Integrating smFRET with molecular dynamics simulations allowed us to estimate the forces generated in a power stroke. Subsequently, the quantitative force data and the measured kinetics of the GTPase cycle were incorporated into a model including both a dynamin filament, with explicit motor cross-bridges, and a realistic deformable membrane template. In our simulations, collective constriction of the membrane by dynamin motor modules, based on the ratchet mechanism, is directly reproduced and analyzed. Functional parallels between the dynamin system and actomyosin in the muscle are seen. Through concerted action of the motors, tight membrane constriction to the hemifission radius can be reached. Our experimental and computational study provides an example of how collective motor action in megadalton molecular assemblies can be approached and explicitly resolved.
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Affiliation(s)
- Oleg M Ganichkin
- Crystallography, Max Delbrück Center for Molecular Medicine in the Helmholtz Association, 13125 Berlin, Germany
| | - Renee Vancraenenbroeck
- Department of Chemical and Structural Biology, Weizmann Institute of Science, 76100 Rehovot, Israel
| | - Gabriel Rosenblum
- Department of Chemical and Structural Biology, Weizmann Institute of Science, 76100 Rehovot, Israel
| | - Hagen Hofmann
- Department of Chemical and Structural Biology, Weizmann Institute of Science, 76100 Rehovot, Israel
| | - Alexander S Mikhailov
- Department of Physical Chemistry, Fritz Haber Institute of the Max Planck Society, 14195 Berlin, Germany
- Computational Molecular Biophysics, Nano Life Science Institute, Kanazawa University, Kanazawa 920-1192, Japan
| | - Oliver Daumke
- Crystallography, Max Delbrück Center for Molecular Medicine in the Helmholtz Association, 13125 Berlin, Germany;
- Institute for Chemistry and Biochemistry, Free University of Berlin, 14195 Berlin, Germany
| | - Jeffrey K Noel
- Crystallography, Max Delbrück Center for Molecular Medicine in the Helmholtz Association, 13125 Berlin, Germany;
- Department of Physical Chemistry, Fritz Haber Institute of the Max Planck Society, 14195 Berlin, Germany
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14
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Johannes L, Valades-Cruz CA. The final twist in endocytic membrane scission. Nat Cell Biol 2021; 23:812-813. [PMID: 34253895 DOI: 10.1038/s41556-021-00711-z] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/09/2022]
Affiliation(s)
- Ludger Johannes
- Cellular and Chemical Biology unit, Institut Curie, U1143 INSERM, UMR3666 CNRS, PSL Research University, Paris, France.
| | - Cesar Augusto Valades-Cruz
- Cellular and Chemical Biology unit, Institut Curie, U1143 INSERM, UMR3666 CNRS, PSL Research University, Paris, France.,Serpico - STED Team, Inria Centre Rennes-Bretagne Atlantique, Institut Curie, UMR144 CNRS, PSL Research University, Campus Universitaire de Beaulieu, Rennes, France
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15
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Cheng X, Chen K, Dong B, Yang M, Filbrun SL, Myoung Y, Huang TX, Gu Y, Wang G, Fang N. Dynamin-dependent vesicle twist at the final stage of clathrin-mediated endocytosis. Nat Cell Biol 2021; 23:859-869. [PMID: 34253896 PMCID: PMC8355216 DOI: 10.1038/s41556-021-00713-x] [Citation(s) in RCA: 21] [Impact Index Per Article: 7.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/06/2021] [Accepted: 06/09/2021] [Indexed: 12/11/2022]
Abstract
Dynamin plays an important role in clathrin-mediated endocytosis (CME) by cutting the neck of nascent vesicles from the cell membrane. Here through using gold nanorods as cargos to image dynamin action during live CME, we show that near the peak of dynamin accumulation, the cargo-containing vesicles always exhibit abrupt, right-handed rotations that finish in a short time (~0.28 s). The large and quick twist, herein named the super twist, is the result of the coordinated dynamin helix action upon GTP hydrolysis. After the super twist, the rotational freedom of the vesicle drastically increases, accompanied with simultaneous or delayed translational movement, indicating that it detaches from the cell membrane. These observations suggest that dynamin-mediated scission involves a large torque generated by coordinated actions of multiple dynamins in the helix, which is the main driving force for vesicle scission.
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Affiliation(s)
- Xiaodong Cheng
- Department of Chemistry, Georgia State University, Atlanta, GA, USA.,State Key Laboratory of Physical Chemistry of Solid Surfaces, Innovation Laboratory for Sciences and Technologies of Energy Materials of Fujian Province (IKKEM), College of Chemistry and Chemical Engineering, Xiamen University, Xiamen, China
| | - Kuangcai Chen
- Department of Chemistry, Georgia State University, Atlanta, GA, USA
| | - Bin Dong
- Department of Chemistry, Georgia State University, Atlanta, GA, USA
| | - Meek Yang
- Department of Chemistry, Georgia State University, Atlanta, GA, USA
| | - Seth L Filbrun
- Department of Chemistry, Georgia State University, Atlanta, GA, USA
| | - Yong Myoung
- Department of Chemistry, Georgia State University, Atlanta, GA, USA
| | - Teng-Xiang Huang
- Department of Chemistry, Georgia State University, Atlanta, GA, USA
| | - Yan Gu
- The Bristol-Myers Squibb Company, Devens, MA, USA
| | - Gufeng Wang
- Department of Chemistry, Georgia State University, Atlanta, GA, USA.
| | - Ning Fang
- Department of Chemistry, Georgia State University, Atlanta, GA, USA. .,State Key Laboratory of Physical Chemistry of Solid Surfaces, Innovation Laboratory for Sciences and Technologies of Energy Materials of Fujian Province (IKKEM), College of Chemistry and Chemical Engineering, Xiamen University, Xiamen, China.
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16
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Ramirez-Diaz DA, Merino-Salomón A, Meyer F, Heymann M, Rivas G, Bramkamp M, Schwille P. FtsZ induces membrane deformations via torsional stress upon GTP hydrolysis. Nat Commun 2021; 12:3310. [PMID: 34083531 PMCID: PMC8175707 DOI: 10.1038/s41467-021-23387-3] [Citation(s) in RCA: 20] [Impact Index Per Article: 6.7] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/11/2020] [Accepted: 04/27/2021] [Indexed: 01/28/2023] Open
Abstract
FtsZ is a key component in bacterial cell division, being the primary protein of the presumably contractile Z ring. In vivo and in vitro, it shows two distinctive features that could so far, however, not be mechanistically linked: self-organization into directionally treadmilling vortices on solid supported membranes, and shape deformation of flexible liposomes. In cells, circumferential treadmilling of FtsZ was shown to recruit septum-building enzymes, but an active force production remains elusive. To gain mechanistic understanding of FtsZ dependent membrane deformations and constriction, we design an in vitro assay based on soft lipid tubes pulled from FtsZ decorated giant lipid vesicles (GUVs) by optical tweezers. FtsZ filaments actively transform these tubes into spring-like structures, where GTPase activity promotes spring compression. Operating the optical tweezers in lateral vibration mode and assigning spring constants to FtsZ coated tubes, the directional forces that FtsZ-YFP-mts rings exert upon GTP hydrolysis can be estimated to be in the pN range. They are sufficient to induce membrane budding with constricting necks on both, giant vesicles and E.coli cells devoid of their cell walls. We hypothesize that these forces result from torsional stress in a GTPase activity dependent manner.
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Affiliation(s)
- Diego A Ramirez-Diaz
- Department of Cellular and Molecular Biophysics, Max Planck Institute of Biochemistry, Martinsried, Germany
- Graduate School for Quantitative Biosciences (QBM), Ludwig-Maximillians-University, Munich, Germany
| | - Adrián Merino-Salomón
- Department of Cellular and Molecular Biophysics, Max Planck Institute of Biochemistry, Martinsried, Germany
- International Max Planck Research School for Molecular Life Sciences (IMPRS-LS), Munich, Germany
| | - Fabian Meyer
- Institute of General Microbiology, Christian-Albrechts-Unversity, Kiel, Germany
| | - Michael Heymann
- Department of Cellular and Molecular Biophysics, Max Planck Institute of Biochemistry, Martinsried, Germany
- Institute of Biomaterials and Biomolecular Systems, University of Stuttgart, Stuttgart, Germany
| | - Germán Rivas
- Centro de Investigaciones Biológicas Margarita Salas, Consejo Superior de Investigaciones Cientificas (CSIC), Madrid, Spain
| | - Marc Bramkamp
- Institute of General Microbiology, Christian-Albrechts-Unversity, Kiel, Germany
| | - Petra Schwille
- Department of Cellular and Molecular Biophysics, Max Planck Institute of Biochemistry, Martinsried, Germany.
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17
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Mass photometry enables label-free tracking and mass measurement of single proteins on lipid bilayers. Nat Methods 2021; 18:1247-1252. [PMID: 34608319 PMCID: PMC8490153 DOI: 10.1038/s41592-021-01261-w] [Citation(s) in RCA: 43] [Impact Index Per Article: 14.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/15/2021] [Accepted: 08/04/2021] [Indexed: 02/08/2023]
Abstract
The quantification of membrane-associated biomolecular interactions is crucial to our understanding of various cellular processes. State-of-the-art single-molecule approaches rely largely on the addition of fluorescent labels, which complicates the quantification of the involved stoichiometries and dynamics because of low temporal resolution and the inherent limitations associated with labeling efficiency, photoblinking and photobleaching. Here, we demonstrate dynamic mass photometry, a method for label-free imaging, tracking and mass measurement of individual membrane-associated proteins diffusing on supported lipid bilayers. Application of this method to the membrane remodeling GTPase, dynamin-1, reveals heterogeneous mixtures of dimer-based oligomers, oligomer-dependent mobilities, membrane affinities and (dis)association of individual complexes. These capabilities, together with assay-based advances for studying integral membrane proteins, will enable the elucidation of biomolecular mechanisms in and on lipid bilayers.
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18
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Rode S, Elgeti J, Gompper G. Chiral-filament self-assembly on curved manifolds. SOFT MATTER 2020; 16:10548-10557. [PMID: 33078824 DOI: 10.1039/d0sm01339k] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/11/2023]
Abstract
Rod-like and banana-shaped proteins, like BAR-domain proteins and MreB proteins, adsorb on membranes and regulate the membrane curvature. The formation of large filamentous complexes of these proteins plays an important role in cellular processes like membrane trafficking, cytokinesis and cell motion. We propose a simplified model to investigate such curvature-dependent self-assembly processes. Anisotropic building blocks, modeled as trimer molecules, which have a preferred binding site, interact via pair-wise Lennard-Jones potentials. When several trimers assemble, they form an elastic ribbon with an intrinsic curvature and twist, controlled by bending and torsional rigidity. For trimer self-assembly on the curved surface of a cylindrical membrane, this leads to a preferred spatial orientation of the ribbon. We show that these interactions can lead to the formation of helices with several windings around the cylinder. The emerging helix angle and pitch depend on the rigidities and the intrinsic curvature and twist values. In particular, a well-defined and controllable helix angle emerges in the case of equal bending and torsional rigidity. The dynamics of filament growth is characterized by three regimes, in which filament length increases with the power laws tz in time, with z≃ 3/4, z = 1/2, and z = 0 for short, intermediate, and long times, respectively. A comparison with the solutions of the Smoluchowski aggregation equation allows the identification of the underlying mechanism in the short-time regime as a crossover from size-independent to diffusion-limited aggregation. Thus, helical structures, as often observed in biology, can arise by self-assembly of anisotropic and chiral proteins.
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Affiliation(s)
- Sebastian Rode
- Theoretical Physics of Living Matter, Institute of Biological Information Processing and Institute for Advanced Simulation, Forschungszentrum Jülich, 52425 Jülich, Germany.
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19
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Casuso I, Redondo-Morata L, Rico F. Biological physics by high-speed atomic force microscopy. PHILOSOPHICAL TRANSACTIONS. SERIES A, MATHEMATICAL, PHYSICAL, AND ENGINEERING SCIENCES 2020; 378:20190604. [PMID: 33100165 PMCID: PMC7661283 DOI: 10.1098/rsta.2019.0604] [Citation(s) in RCA: 17] [Impact Index Per Article: 4.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 05/08/2023]
Abstract
While many fields have contributed to biological physics, nanotechnology offers a new scale of observation. High-speed atomic force microscopy (HS-AFM) provides nanometre structural information and dynamics with subsecond resolution of biological systems. Moreover, HS-AFM allows us to measure piconewton forces within microseconds giving access to unexplored, fast biophysical processes. Thus, HS-AFM provides a tool to nourish biological physics through the observation of emergent physical phenomena in biological systems. In this review, we present an overview of the contribution of HS-AFM, both in imaging and force spectroscopy modes, to the field of biological physics. We focus on examples in which HS-AFM observations on membrane remodelling, molecular motors or the unfolding of proteins have stimulated the development of novel theories or the emergence of new concepts. We finally provide expected applications and developments of HS-AFM that we believe will continue contributing to our understanding of nature, by serving to the dialogue between biology and physics. This article is part of a discussion meeting issue 'Dynamic in situ microscopy relating structure and function'.
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Affiliation(s)
- Ignacio Casuso
- Aix-Marseile University, Inserm, CNRS, LAI, 163 Av. de Luminy, 13009 Marseille, France
| | - Lorena Redondo-Morata
- Center for Infection and Immunity of Lille, INSERM U1019, CNRS UMR 8204, 59000 Lille, France
| | - Felix Rico
- Aix-Marseile University, Inserm, CNRS, LAI, 163 Av. de Luminy, 13009 Marseille, France
- e-mail:
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20
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Jiao F, Cannon KS, Lin YC, Gladfelter AS, Scheuring S. The hierarchical assembly of septins revealed by high-speed AFM. Nat Commun 2020; 11:5062. [PMID: 33033254 PMCID: PMC7545167 DOI: 10.1038/s41467-020-18778-x] [Citation(s) in RCA: 18] [Impact Index Per Article: 4.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/24/2020] [Accepted: 09/15/2020] [Indexed: 12/15/2022] Open
Abstract
Septins are GTP-binding proteins involved in diverse cellular processes including division and membrane remodeling. Septins form linear, palindromic heteromeric complexes that can assemble in filaments and higher-order structures. Structural studies revealed various septin architectures, but questions concerning assembly-dynamics and -pathways persist. Here we used high-speed atomic force microscopy (HS-AFM) and kinetic modeling which allowed us to determine that septin filament assembly was a diffusion-driven process, while formation of higher-order structures was complex and involved self-templating. Slightly acidic pH and increased monovalent ion concentrations favor filament-assembly, -alignment and -pairing. Filament-alignment and -pairing further favored diffusion-driven assembly. Pairing is mediated by the septin N-termini face, and may occur symmetrically or staggered, likely important for the formation of higher-order structures of different shapes. Multilayered structures are templated by the morphology of the underlying layers. The septin C-termini face, namely the C-terminal extension of Cdc12, may be involved in membrane binding.
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Affiliation(s)
- Fang Jiao
- Department of Anesthesiology, Weill Cornell Medicine, New York, NY, 10065, USA
- Department of Physiology and Biophysics, Weill Cornell Medicine, New York, NY, 10065, USA
| | - Kevin S Cannon
- Department of Biology, University of North Carolina and Chapel Hill, Chapel Hill, NC, 27599, USA
| | - Yi-Chih Lin
- Department of Anesthesiology, Weill Cornell Medicine, New York, NY, 10065, USA
- Department of Physiology and Biophysics, Weill Cornell Medicine, New York, NY, 10065, USA
| | - Amy S Gladfelter
- Department of Biology, University of North Carolina and Chapel Hill, Chapel Hill, NC, 27599, USA
- Marine Biological Laboratory, Woods Hole, MA, 02543, USA
| | - Simon Scheuring
- Department of Anesthesiology, Weill Cornell Medicine, New York, NY, 10065, USA.
- Department of Physiology and Biophysics, Weill Cornell Medicine, New York, NY, 10065, USA.
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21
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Bashkirov PV, Kuzmin PI, Chekashkina K, Arrasate P, Vera Lillo J, Shnyrova AV, Frolov VA. Reconstitution and real-time quantification of membrane remodeling by single proteins and protein complexes. Nat Protoc 2020; 15:2443-2469. [PMID: 32591769 PMCID: PMC10839814 DOI: 10.1038/s41596-020-0337-1] [Citation(s) in RCA: 10] [Impact Index Per Article: 2.5] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/14/2020] [Accepted: 04/14/2020] [Indexed: 02/08/2023]
Abstract
Cellular membrane processes, from signal transduction to membrane fusion and fission, depend on acute membrane deformations produced by small and short-lived protein complexes working in conditions far from equilibrium. Real-time monitoring and quantitative assessment of such deformations are challenging; hence, mechanistic analyses of the protein action are commonly based on ensemble averaging, which masks important mechanistic details of the action. In this protocol, we describe how to reconstruct and quantify membrane remodeling by individual proteins and small protein complexes in vitro, using an ultra-short (80- to 400-nm) lipid nanotube (usNT) template. We use the luminal conductance of the usNT as the real-time reporter of the protein interaction(s) with the usNT. We explain how to make and calibrate the usNT template to achieve subnanometer precision in the geometrical assessment of the molecular footprints on the nanotube membrane. We next demonstrate how membrane deformations driven by purified proteins implicated in cellular membrane remodeling can be analyzed at a single-molecule level. The preparation of one usNT takes ~1 h, and the shortest procedure yielding the basic geometrical parameters of a small protein complex takes 10 h.
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Affiliation(s)
- Pavel V Bashkirov
- Federal Research and Clinical Centre of Physical-Chemical Medicine, Moscow, Russia.
- Department of Molecular and Biological Physics, Moscow Institute of Physics and Technology, Dolgoprudnyy, Russia.
| | - Peter I Kuzmin
- A. N. Frumkin Institute of Physical Chemistry and Electrochemistry, Russian Academy of Sciences, Moscow, Russia
| | - Ksenia Chekashkina
- Federal Research and Clinical Centre of Physical-Chemical Medicine, Moscow, Russia
| | - Pedro Arrasate
- Biofisika Institute (CSIC, UPV/EHU), University of the Basque Country, Leioa, Spain
- Department of Biochemistry and Molecular Biology, University of the Basque Country, Leioa, Spain
| | - Javier Vera Lillo
- Biofisika Institute (CSIC, UPV/EHU), University of the Basque Country, Leioa, Spain
- Department of Biochemistry and Molecular Biology, University of the Basque Country, Leioa, Spain
| | - Anna V Shnyrova
- Biofisika Institute (CSIC, UPV/EHU), University of the Basque Country, Leioa, Spain
- Department of Biochemistry and Molecular Biology, University of the Basque Country, Leioa, Spain
| | - Vadim A Frolov
- Biofisika Institute (CSIC, UPV/EHU), University of the Basque Country, Leioa, Spain.
- Department of Biochemistry and Molecular Biology, University of the Basque Country, Leioa, Spain.
- Ikerbasque, Basque Foundation for Science, Bilbao, Spain.
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22
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Maity S, Ottelé J, Santiago GM, Frederix PWJM, Kroon P, Markovitch O, Stuart MCA, Marrink SJ, Otto S, Roos WH. Caught in the Act: Mechanistic Insight into Supramolecular Polymerization-Driven Self-Replication from Real-Time Visualization. J Am Chem Soc 2020; 142:13709-13717. [PMID: 32786814 PMCID: PMC7426903 DOI: 10.1021/jacs.0c02635] [Citation(s) in RCA: 35] [Impact Index Per Article: 8.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/13/2022]
Abstract
![]()
Self-assembly features
prominently in fields ranging from materials
science to biophysical chemistry. Assembly pathways, often passing
through transient intermediates, can control the outcome of assembly
processes. Yet, the mechanisms of self-assembly remain largely obscure
due to a lack of experimental tools for probing these pathways at
the molecular level. Here, the self-assembly of self-replicators into
fibers is visualized in real-time by high-speed atomic force microscopy
(HS-AFM). Fiber growth requires the conversion of precursor molecules
into six-membered macrocycles, which constitute the fibers. HS-AFM
experiments, supported by molecular dynamics simulations, revealed
that aggregates of precursor molecules accumulate at the sides of
the fibers, which then diffuse to the fiber ends where growth takes
place. This mechanism of precursor reservoir formation, followed by
one-dimensional diffusion, which guides the precursor molecules to
the sites of growth, reduces the entropic penalty associated with
colocalizing precursors and growth sites and constitutes a new mechanism
for supramolecular polymerization.
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Affiliation(s)
- Sourav Maity
- Molecular Biophysics, Zernike Institute for Advanced Materials, University of Groningen, Nijenborgh 4, Groningen 9747 AG, The Netherlands
| | - Jim Ottelé
- Centre for Systems Chemistry, Stratingh Institute, University of Groningen, Nijenborgh 4, Groningen 9747 AG, The Netherlands
| | - Guillermo Monreal Santiago
- Centre for Systems Chemistry, Stratingh Institute, University of Groningen, Nijenborgh 4, Groningen 9747 AG, The Netherlands
| | - Pim W J M Frederix
- Groningen Biomolecular Sciences and Biotechnology Institute & Zernike Institute for Advanced Materials, University of Groningen, Nijenborgh 7, Groningen 9747 AG, The Netherlands
| | - Peter Kroon
- Groningen Biomolecular Sciences and Biotechnology Institute & Zernike Institute for Advanced Materials, University of Groningen, Nijenborgh 7, Groningen 9747 AG, The Netherlands
| | - Omer Markovitch
- Centre for Systems Chemistry, Stratingh Institute, University of Groningen, Nijenborgh 4, Groningen 9747 AG, The Netherlands.,Origins Center, Nijenborgh 7, Groningen 9747 AG, The Netherlands
| | - Marc C A Stuart
- Centre for Systems Chemistry, Stratingh Institute, University of Groningen, Nijenborgh 4, Groningen 9747 AG, The Netherlands
| | - Siewert J Marrink
- Groningen Biomolecular Sciences and Biotechnology Institute & Zernike Institute for Advanced Materials, University of Groningen, Nijenborgh 7, Groningen 9747 AG, The Netherlands
| | - Sijbren Otto
- Centre for Systems Chemistry, Stratingh Institute, University of Groningen, Nijenborgh 4, Groningen 9747 AG, The Netherlands
| | - Wouter H Roos
- Molecular Biophysics, Zernike Institute for Advanced Materials, University of Groningen, Nijenborgh 4, Groningen 9747 AG, The Netherlands
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23
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Polymer-like Model to Study the Dynamics of Dynamin Filaments on Deformable Membrane Tubes. Biophys J 2019; 117:1870-1891. [PMID: 31672269 DOI: 10.1016/j.bpj.2019.09.042] [Citation(s) in RCA: 8] [Impact Index Per Article: 1.6] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/18/2019] [Revised: 09/08/2019] [Accepted: 09/24/2019] [Indexed: 01/30/2023] Open
Abstract
Peripheral membrane proteins with intrinsic curvature can act both as sensors of membrane curvature and shape modulators of the underlying membranes. A well-studied example of such proteins is the mechanochemical GTPase dynamin, which assembles into helical filaments around membrane tubes and catalyzes their scission in a GTPase-dependent manner. It is known that the dynamin coat alone, without GTP, can constrict membrane tubes to radii of ∼10 nm, indicating that the intrinsic shape and elasticity of dynamin filaments should play an important role in membrane remodeling. However, molecular and dynamic understanding of the process is lacking. Here, we develop a dynamical polymer-chain model for a helical elastic filament bound on a deformable membrane tube of conserved mass, accounting for thermal fluctuations in the filament and lipid flows in the membrane. The model is based on the locally cylindrical helix approximation for dynamin. We obtain the elastic parameters of the dynamin filament by molecular dynamics simulations of its tetrameric building block and also from coarse-grained structure-based simulations of a 17-dimer filament. The results show that the stiffness of dynamin is comparable to that of the membrane. We determine equilibrium shapes of the filament and the membrane and find that mostly the pitch of the filament, not its radius, is sensitive to variations in membrane tension and stiffness. The close correspondence between experimental estimates of the inner tube radius and those predicted by the model suggests that dynamin's "stalk" region is responsible for its GTP-independent membrane-shaping ability. The model paves the way for future mesoscopic modeling of dynamin with explicit motor function.
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24
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Bahrami A, Bahrami AH. Vesicle constriction by rings of Janus nanoparticles and aggregates of curved proteins. NANOTECHNOLOGY 2019; 30:345101. [PMID: 31048566 DOI: 10.1088/1361-6528/ab1ed5] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/09/2023]
Abstract
Membrane constriction and associated scission by proteins and nano structures are crucial to many processes in cellular and synthetic biology. We report mechanical constriction of vesicles by rings of adsorbed Janus nanoparticles that represent synthetic nano structures and mimic contractile proteins, and by aggregates of curved crescents that mimic scaffold proteins. Membrane energetics from Monte Carlo simulations and simulated annealing of the elastic membrane model confirms spontaneous vesicle constriction by aggregates of sufficiently-curved crescents of various lengths and by rings of Janus nanoparticles with a variety of ring lengths, particle sizes, and particle area fractions. We show that shorter rings of smaller particles with higher area fractions reinforce the constriction by increasing the energetic drive towards the constricted vesicle with smaller constriction radius. We demonstrate that vesicle constriction by crescent aggregates strongly depends on the crescent curvature. In contrast to aggregates of sufficiently-curved crescents that are capable of inducing full vesicle constriction, those of near flat crescents with negligible curvature leave the vesicle unconstricted. Our results offer promising perspectives for designing membrane-constricting nano structures such as nanoparticle aggregates and clusters of synthetic curved proteins such as DNA origami scaffolds with applications in synthetic biology. Our findings reveal the significant contribution of highly-curved F-BAR domains to cell division and explain how contractile protein rings such as dynamin GTPase, actomyosin rings, and endosomal sorting complexes required for transport constrict the membrane.
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Affiliation(s)
- Arash Bahrami
- School of Mechanical Engineering, College of Engineering, University of Tehran, North Kargar St., 14399-57131 Tehran, Iran
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25
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Abstract
Dynamin proteins assemble into characteristic helical structures around necks of clathrin-coated membrane buds. Hydrolysis of dynamin-bound GTP results in both fission of the membrane neck and partial disruption of the dynamin oligomer. Imaging by atomic force microscopy reveals that, on GTP hydrolysis, dynamin oligomers undergo a dynamic remodeling and lose their distinctive helical shape. While breakup of the dynamin helix is a critical stage in clathrin-mediated endocytosis, the mechanism for this remodeling of the oligomer has not been resolved. In this paper, we formulate an analytical, elasticity-based model for the reshaping and disassembly of the dynamin scaffold. We predict that the shape of the oligomer is modulated by the orientation of dynamin's pleckstrin homology (PH) domain relative to the underlying membrane. Our results indicate that tilt of the PH domain drives deformation and fragmentation of the oligomer, in agreement with experimental observations. This model motivated the introduction of the tilted helix: a curve that maintains a fixed angle between its normal and the normal of the embedding surface. Our findings highlight the importance of tilt as a key regulator of size and morphology of membrane-bound oligomers.
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26
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Visualizing Biological Membrane Organization and Dynamics. J Mol Biol 2019; 431:1889-1919. [DOI: 10.1016/j.jmb.2019.02.018] [Citation(s) in RCA: 13] [Impact Index Per Article: 2.6] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/26/2018] [Revised: 02/02/2019] [Accepted: 02/13/2019] [Indexed: 11/22/2022]
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27
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Yewdall NA, Mason AF, van Hest JCM. The hallmarks of living systems: towards creating artificial cells. Interface Focus 2018; 8:20180023. [PMID: 30443324 PMCID: PMC6227776 DOI: 10.1098/rsfs.2018.0023] [Citation(s) in RCA: 88] [Impact Index Per Article: 14.7] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Accepted: 06/29/2018] [Indexed: 01/01/2023] Open
Abstract
Despite the astonishing diversity and complexity of living systems, they all share five common hallmarks: compartmentalization, growth and division, information processing, energy transduction and adaptability. In this review, we give not only examples of how cells satisfy these requirements for life and the ways in which it is possible to emulate these characteristics in engineered platforms, but also the gaps that remain to be bridged. The bottom-up synthesis of life-like systems continues to be driven forward by the advent of new technologies, by the discovery of biological phenomena through their transplantation to experimentally simpler constructs and by providing insights into one of the oldest questions posed by mankind, the origin of life on Earth.
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Affiliation(s)
| | | | - Jan C. M. van Hest
- Eindhoven University of Technology, PO Box 513 (STO 3.31), Eindhoven, MB, The Netherlands
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28
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Miyagi A, Scheuring S. A novel phase-shift-based amplitude detector for a high-speed atomic force microscope. THE REVIEW OF SCIENTIFIC INSTRUMENTS 2018; 89:083704. [PMID: 30184715 DOI: 10.1063/1.5038095] [Citation(s) in RCA: 18] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Received: 05/01/2018] [Accepted: 07/29/2018] [Indexed: 06/08/2023]
Abstract
In any atomic force microscope operated in amplitude modulation mode, aka "tapping mode" or "oscillating mode," the most crucial operation is the detection of the cantilever oscillation amplitude. Indeed, it is the change in the cantilever oscillation amplitude that drives the feedback loop, and thus, the accuracy and speed of amplitude detection are of utmost importance for improved atomic force microscopy operation. This becomes even more crucial for the operation of a high-speed atomic force microscope (HS-AFM), where feedback operation on a single or a low number of cantilever oscillation cycles between 500 kHz and 1000 kHz oscillation frequency is desired. So far, the amplitude detection was performed by Fourier analysis of each oscillation, resulting in a single output amplitude value at the end of each oscillation cycle, i.e., 360° phase delay. Here, we present a novel analog amplitude detection circuit with theoretic continuous amplitude detection at 90° phase delay. In factual operation, when exposed to an abrupt amplitude change, our novel amplitude detector circuit reacted with a phase delay of ∼138° compared with the phase delay of ∼682° achieved by the Fourier analysis method. Integrated to a HS-AFM, the novel amplitude detector should allow faster image acquisition with lower invasiveness due to the faster and more accurate detection of cantilever oscillation amplitude change.
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Affiliation(s)
- Atsushi Miyagi
- Department of Anesthesiology, Weill Cornell Medicine, 1300 York Avenue, New York, New York 10065, USA
| | - Simon Scheuring
- Department of Anesthesiology, Weill Cornell Medicine, 1300 York Avenue, New York, New York 10065, USA
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29
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Yoshida A, Sakai N, Uekusa Y, Imaoka Y, Itagaki Y, Suzuki Y, Yoshimura SH. Morphological changes of plasma membrane and protein assembly during clathrin-mediated endocytosis. PLoS Biol 2018; 16:e2004786. [PMID: 29723197 PMCID: PMC5953504 DOI: 10.1371/journal.pbio.2004786] [Citation(s) in RCA: 40] [Impact Index Per Article: 6.7] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/11/2017] [Revised: 05/15/2018] [Accepted: 03/29/2018] [Indexed: 12/21/2022] Open
Abstract
Clathrin-mediated endocytosis (CME) proceeds through a series of morphological changes of the plasma membrane induced by a number of protein components. Although the spatiotemporal assembly of these proteins has been elucidated by fluorescence-based techniques, the protein-induced morphological changes of the plasma membrane have not been fully clarified in living cells. Here, we visualize membrane morphology together with protein localizations during CME by utilizing high-speed atomic force microscopy (HS-AFM) combined with a confocal laser scanning unit. The plasma membrane starts to invaginate approximately 30 s after clathrin starts to assemble, and the aperture diameter increases as clathrin accumulates. Actin rapidly accumulates around the pit and induces a small membrane swelling, which, within 30 s, rapidly covers the pit irreversibly. Inhibition of actin turnover abolishes the swelling and induces a reversible open–close motion of the pit, indicating that actin dynamics are necessary for efficient and irreversible pit closure at the end of CME. Cells communicate with their environments via the plasma membrane and various membrane proteins. Clathrin-mediated endocytosis (CME) plays a central role in such communication and proceeds with a series of multiprotein assembly, deformation of the plasma membrane, and production of a membrane vesicle that delivers extracellular signaling molecules into the cytoplasm. In this study, we utilized our home-built correlative imaging system comprising high-speed atomic force microscopy (HS-AFM) and confocal fluorescence microscopy to simultaneously image morphological changes of the plasma membrane and protein localization during CME in a living cell. The results revealed a tight correlation between the size of the pit and the amount of clathrin assembled. Actin dynamics play multiple roles in the assembly, maturation, and closing phases of the process, and affects membrane morphology, suggesting a close relationship between endocytosis and dynamic events at the cell cortex. Knock down of dynamin also affected the closing motion of the pit and showed functional correlation with actin.
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Affiliation(s)
- Aiko Yoshida
- Graduate School of Biostudies, Kyoto University, Kyoto, Japan
| | | | | | - Yuka Imaoka
- R&D Group, Olympus Corporation, Hachioji, Japan
| | | | - Yuki Suzuki
- Frontier Research Institute for Interdisciplinary Sciences, Tohoku University, Sendai, Japan
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30
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Increasing Diversity of Biological Membrane Fission Mechanisms. Trends Cell Biol 2018; 28:274-286. [DOI: 10.1016/j.tcb.2017.12.001] [Citation(s) in RCA: 34] [Impact Index Per Article: 5.7] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/02/2017] [Revised: 12/06/2017] [Accepted: 12/12/2017] [Indexed: 12/19/2022]
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31
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Takeda T, Kozai T, Yang H, Ishikuro D, Seyama K, Kumagai Y, Abe T, Yamada H, Uchihashi T, Ando T, Takei K. Dynamic clustering of dynamin-amphiphysin helices regulates membrane constriction and fission coupled with GTP hydrolysis. eLife 2018; 7:30246. [PMID: 29357276 PMCID: PMC5780043 DOI: 10.7554/elife.30246] [Citation(s) in RCA: 29] [Impact Index Per Article: 4.8] [Reference Citation Analysis] [Abstract] [Key Words] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/08/2017] [Accepted: 12/18/2017] [Indexed: 01/16/2023] Open
Abstract
Dynamin is a mechanochemical GTPase essential for membrane fission during clathrin-mediated endocytosis. Dynamin forms helical complexes at the neck of clathrin-coated pits and their structural changes coupled with GTP hydrolysis drive membrane fission. Dynamin and its binding protein amphiphysin cooperatively regulate membrane remodeling during the fission, but its precise mechanism remains elusive. In this study, we analyzed structural changes of dynamin-amphiphysin complexes during the membrane fission using electron microscopy (EM) and high-speed atomic force microscopy (HS-AFM). Interestingly, HS-AFM analyses show that the dynamin-amphiphysin helices are rearranged to form clusters upon GTP hydrolysis and membrane constriction occurs at protein-uncoated regions flanking the clusters. We also show a novel function of amphiphysin in size control of the clusters to enhance biogenesis of endocytic vesicles. Our approaches using combination of EM and HS-AFM clearly demonstrate new mechanistic insights into the dynamics of dynamin-amphiphysin complexes during membrane fission. The nerve cells that make up a nervous system connect at junctions known as synapses. When a nerve impulse reaches the end of the cell, membrane-bound packages called vesicles fuse with the surface membrane and release their contents to the outside. The contents, namely chemicals called neurotransmitters, then travels across the synapse, relaying the signal to the next cell. Nerve cells can fire many times per second. The membrane from fused vesicles must be retrieved from the surface membrane and recycled to make new vesicles, ready to transmit more signals across the synapse. Many proteins at these sites are involved in folding the fused membrane back into the cell, constricting the opening, and eventually pinching off the new vesicles – a process known as endocytosis. Two proteins named dynamin and amphiphysin cooperate in this process, but their precise mechanism remained elusive. Dynamin is a protein that acts like a motor; it breaks down a molecule called GTP to release energy. Previous studies have seen that dynamin-amphiphysin complexes join end to end to form long helical structures. Takeda et al. have now looked at how the structure of the helices changes during endocytosis. This revealed that the dynamin-amphiphysin helices rearrange to form clusters when the GTP is broken down. Further analysis showed that the folded membrane becomes constricted at regions that are not coated with the clusters of dynamin-amphiphysin helices. Takeda et al. also discovered that amphiphysin controls the size of the clusters to help make the new vesicles more uniform. The gene for dynamin is altered in a number of disorders affecting the nervous system and muscles, including epileptic encephalopathy, Charcot-Marie-Tooth disease and congenital myopathy. Moreover, a neurological disorder characterized by muscle stiffness (known as Stiff-person syndrome) occurs when an individual’s immune system mistakenly attacks the amphiphysin protein. As such, these new findings will not only help scientists to better understand the process of endocytosis, but they will also give new insight into a number of human diseases.
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Affiliation(s)
- Tetsuya Takeda
- Graduate School of Medicine, Dentistry and Pharmaceutical Sciences, Okayama University, Okayama, Japan
| | - Toshiya Kozai
- Department of Physics, College of Science and Engineering, Kanazawa University, Kanazawa, Japan
| | - Huiran Yang
- Graduate School of Medicine, Dentistry and Pharmaceutical Sciences, Okayama University, Okayama, Japan
| | - Daiki Ishikuro
- Department of Physics, College of Science and Engineering, Kanazawa University, Kanazawa, Japan
| | - Kaho Seyama
- Graduate School of Medicine, Dentistry and Pharmaceutical Sciences, Okayama University, Okayama, Japan
| | - Yusuke Kumagai
- Department of Physics, College of Science and Engineering, Kanazawa University, Kanazawa, Japan
| | - Tadashi Abe
- Graduate School of Medicine, Dentistry and Pharmaceutical Sciences, Okayama University, Okayama, Japan
| | - Hiroshi Yamada
- Graduate School of Medicine, Dentistry and Pharmaceutical Sciences, Okayama University, Okayama, Japan
| | - Takayuki Uchihashi
- CREST, JST, Saitama, Japan.,Department of Physics, School of Science, Nagoya University, Nagoya, Japan
| | - Toshio Ando
- CREST, JST, Saitama, Japan.,Bio-AFM Frontier Research Center, College of Science and Engineering, Kanazawa University, Kanazawa, Japan
| | - Kohji Takei
- Graduate School of Medicine, Dentistry and Pharmaceutical Sciences, Okayama University, Okayama, Japan.,CREST, JST, Saitama, Japan
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32
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Optimum Substrates for Imaging Biological Molecules with High-Speed Atomic Force Microscopy. Methods Mol Biol 2018; 1814:159-179. [PMID: 29956232 DOI: 10.1007/978-1-4939-8591-3_10] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.2] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/26/2022]
Abstract
Recent progresses in high-speed atomic force microscopy (HS-AFM) have enabled us to directly visualize dynamic processes of various proteins in liquid conditions. One of the key factors leading to successful HS-AFM observations is the selection of an appropriate substrate depending on molecules to be observed. For the HS-AFM imaging, a target molecule must be absorbed on a substrate by controlling its orientation without impairing the dynamics or physiological function of the molecule. In this chapter, we describe protocols for preparation of substrates that have been used for HS-AFM and then introduce observation examples on dynamic processes of biological molecules.
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