1
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Razavi S, Wong F, Abubaker-Sharif B, Matsubayashi HT, Nakamura H, Nguyen NTH, Robinson DN, Chen B, Iglesias PA, Inoue T. Synthetic control of actin polymerization and symmetry breaking in active protocells. SCIENCE ADVANCES 2024; 10:eadk9731. [PMID: 38865458 PMCID: PMC11168455 DOI: 10.1126/sciadv.adk9731] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 09/20/2023] [Accepted: 05/08/2024] [Indexed: 06/14/2024]
Abstract
Nonlinear biomolecular interactions on membranes drive membrane remodeling crucial for biological processes including chemotaxis, cytokinesis, and endocytosis. The complexity of biomolecular interactions, their redundancy, and the importance of spatiotemporal context in membrane organization impede understanding of the physical principles governing membrane mechanics. Developing a minimal in vitro system that mimics molecular signaling and membrane remodeling while maintaining physiological fidelity poses a major challenge. Inspired by chemotaxis, we reconstructed chemically regulated actin polymerization inside vesicles, guiding membrane self-organization. An external, undirected chemical input induced directed actin polymerization and membrane deformation uncorrelated with upstream biochemical cues, suggesting symmetry breaking. A biophysical model incorporating actin dynamics and membrane mechanics proposes that uneven actin distributions cause nonlinear membrane deformations, consistent with experimental findings. This protocellular system illuminates the interplay between actin dynamics and membrane shape during symmetry breaking, offering insights into chemotaxis and other cell biological processes.
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Affiliation(s)
- Shiva Razavi
- Department of Biomedical Engineering, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
| | - Felix Wong
- Institute for Medical Engineering and Science, Department of Biological Engineering, Massachusetts Institute of Technology (MIT), Cambridge, MA 02139, USA
- Infectious Disease and Microbiome Program, Broad Institute of MIT and Harvard, Cambridge, MA 02139, USA
| | - Bedri Abubaker-Sharif
- Department of Biomedical Engineering, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
| | - Hideaki T. Matsubayashi
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
| | - Hideki Nakamura
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
| | - Nhung Thi Hong Nguyen
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
| | - Douglas N. Robinson
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
- Department of Chemical and Biomolecular Engineering, Johns Hopkins University, Baltimore, MD 21218, USA
| | - Baoyu Chen
- Roy J. Carver Department of Biochemistry, Biophysics and Molecular Biology, Iowa State University, Ames, IA 50011, USA
| | - Pablo A. Iglesias
- Department of Biomedical Engineering, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
- Department of Electrical and Computer Engineering, Whiting School of Engineering, Johns Hopkins University, Baltimore, MD 21218, USA
| | - Takanari Inoue
- Department of Biomedical Engineering, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
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2
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Ioannou IA, Brooks NJ, Kuimova MK, Elani Y. Visualizing Actin Packing and the Effects of Actin Attachment on Lipid Membrane Viscosity Using Molecular Rotors. JACS AU 2024; 4:2041-2049. [PMID: 38818078 PMCID: PMC11134356 DOI: 10.1021/jacsau.4c00237] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Figures] [Subscribe] [Scholar Register] [Received: 03/14/2024] [Revised: 04/10/2024] [Accepted: 04/11/2024] [Indexed: 06/01/2024]
Abstract
The actin cytoskeleton and its elaborate interplay with the plasma membrane participate in and control numerous biological processes in eukaryotic cells. Malfunction of actin networks and changes in their dynamics are related to various diseases, from actin myopathies to uncontrolled cell growth and tumorigenesis. Importantly, the biophysical and mechanical properties of actin and its assemblies are deeply intertwined with the biological functions of the cytoskeleton. Novel tools to study actin and its associated biophysical features are, therefore, of prime importance. Here we develop a new approach which exploits fluorescence lifetime imaging microscopy (FLIM) and environmentally sensitive fluorophores termed molecular rotors, acting as quantitative microviscosity sensors, to monitor dynamic viscoelastic properties of both actin structures and lipid membranes. In order to reproduce a minimal actin cortex in synthetic cell models, we encapsulated and attached actin networks to the lipid bilayer of giant unilamellar vesicles (GUVs). Using a cyanine-based molecular rotor, DiSC2(3), we show that different types of actin bundles are characterized by distinct packing, which can be spatially resolved using FLIM. Similarly, we show that a lipid bilayer-localized molecular rotor can monitor the effects of attaching cross-linked actin networks to the lipid membrane, revealing an increase in membrane viscosity upon actin attachment. Our approach bypasses constraints associated with existing methods for actin imaging, many of which lack the capability for direct visualization of biophysical properties. It can therefore contribute to a deeper understanding of the role that actin plays in fundamental biological processes and help elucidate the underlying biophysics of actin-related diseases.
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Affiliation(s)
- Ion A. Ioannou
- Department
of Chemistry, Imperial College London, Molecular
Sciences Research Hub, London W12 0BZ, U.K.
- Department
of Chemical Engineering, Imperial College
London, South Kensington, London SW7 2AZ, U.K.
| | - Nickolas J. Brooks
- Department
of Chemistry, Imperial College London, Molecular
Sciences Research Hub, London W12 0BZ, U.K.
| | - Marina K. Kuimova
- Department
of Chemistry, Imperial College London, Molecular
Sciences Research Hub, London W12 0BZ, U.K.
| | - Yuval Elani
- Department
of Chemical Engineering, Imperial College
London, South Kensington, London SW7 2AZ, U.K.
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3
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Gandikota MC, Das S, Cacciuto A. Spontaneous crumpling of active spherical shells. SOFT MATTER 2024; 20:3635-3640. [PMID: 38619604 DOI: 10.1039/d4sm00015c] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 04/16/2024]
Abstract
The existence of a crumpled phase for self-avoiding elastic surfaces was postulated more than three decades ago using simple Flory-like scaling arguments. Despite much effort, its stability in a microscopic environment has been the subject of much debate. In this paper we show how a crumpled phase develops reliably and consistently upon subjecting a thin spherical shell to active fluctuations. We find a master curve describing how the relative volume of a shell changes with the strength of the active forces, that applies for every shell independent of size and elastic constants. Furthermore, we extract a general expression for the onset active force beyond which a shell begins to crumple. Finally, we calculate how the size exponent varies along the crumpling curve.
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Affiliation(s)
- M C Gandikota
- Department of Chemistry, Columbia University, 3000 Broadway, NY 10027, New York, USA.
| | - Shibananda Das
- Department of Chemistry, Columbia University, 3000 Broadway, NY 10027, New York, USA.
- Institute for Theoretical Physics, Georg-August University, Friedrich-Hund-Platz 1, 37077 Göttingen, Germany
| | - A Cacciuto
- Department of Chemistry, Columbia University, 3000 Broadway, NY 10027, New York, USA.
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4
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Razavi S, Wong F, Abubaker-Sharif B, Matsubayashi HT, Nakamura H, Sandoval E, Robinson DN, Chen B, Liu J, Iglesias PA, Inoue T. Synthetic control of actin polymerization and symmetry breaking in active protocells. BIORXIV : THE PREPRINT SERVER FOR BIOLOGY 2023:2023.09.22.559060. [PMID: 37790449 PMCID: PMC10542490 DOI: 10.1101/2023.09.22.559060] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 10/05/2023]
Abstract
Non-linear biomolecular interactions on the membranes drive membrane remodeling that underlies fundamental biological processes including chemotaxis, cytokinesis, and endocytosis. The multitude of biomolecules, the redundancy in their interactions, and the importance of spatiotemporal context in membrane organization hampers understanding the physical principles governing membrane mechanics. A minimal, in vitro system that models the functional interactions between molecular signaling and membrane remodeling, while remaining faithful to cellular physiology and geometry is powerful yet remains unachieved. Here, inspired by the biophysical processes underpinning chemotaxis, we reconstituted externally-controlled actin polymerization inside giant unilamellar vesicles, guiding self-organization on the membrane. We show that applying undirected external chemical inputs to this system results in directed actin polymerization and membrane deformation that are uncorrelated with upstream biochemical cues, indicating symmetry breaking. A biophysical model of the dynamics and mechanics of both actin polymerization and membrane shape suggests that inhomogeneous distributions of actin generate membrane shape deformations in a non-linear fashion, a prediction consistent with experimental measurements and subsequent local perturbations. The active protocellular system demonstrates the interplay between actin dynamics and membrane shape in a symmetry breaking context that is relevant to chemotaxis and a suite of other biological processes.
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Affiliation(s)
- Shiva Razavi
- Department of Biomedical Engineering, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
| | - Felix Wong
- Institute for Medical Engineering & Science, Department of Biological Engineering, Massachusetts Institute of Technology (MIT), Cambridge, MA 02139, USA
- Infectious Disease and Microbiome Program, Broad Institute of MIT and Harvard, Cambridge, MA 02139, USA
| | - Bedri Abubaker-Sharif
- Department of Biomedical Engineering, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
| | - Hideaki T. Matsubayashi
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
| | - Hideki Nakamura
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
| | - Eduardo Sandoval
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
| | - Douglas N. Robinson
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
- Department of Chemical and Biomolecular Engineering, Johns Hopkins University, Baltimore, MD 21218, USA
| | - Baoyu Chen
- Roy J. Carver Department of Biochemistry, Biophysics and Molecular Biology, Iowa State University, Ames, IA 50011, USA
| | - Jian Liu
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
| | - Pablo A. Iglesias
- Department of Biomedical Engineering, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
- Department of Electrical and Computer Engineering, Whiting School of Engineering, Johns Hopkins University, Baltimore, MD 21218, USA
| | - Takanari Inoue
- Department of Biomedical Engineering, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
- Department of Cell Biology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA
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5
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Lopes dos Santos R, Campillo C. Studying actin-induced cell shape changes using Giant Unilamellar Vesicles and reconstituted actin networks. Biochem Soc Trans 2022; 50:1527-1539. [PMID: 36111807 PMCID: PMC9704537 DOI: 10.1042/bst20220900] [Citation(s) in RCA: 2] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/08/2022] [Revised: 08/12/2022] [Accepted: 08/22/2022] [Indexed: 10/05/2023]
Abstract
Cell shape changes that are fuelled by the dynamics of the actomyosin cytoskeleton control cellular processes such as motility and division. However, the mechanisms of interplay between cell membranes and actomyosin are complicated to decipher in the complex environment of the cytoplasm. Using biomimetic systems offers an alternative approach to studying cell shape changes in assays with controlled biochemical composition. Biomimetic systems allow quantitative experiments that can help to build physical models describing the processes of cell shape changes. This article reviews works in which actin networks are reconstructed inside or outside cell-sized Giant Unilamellar Vesicles (GUVs), which are models of cell membranes. We show how various actin networks affect the shape and mechanics of GUVs and how some cell shape changes can be reproduced in vitro using these minimal systems.
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Affiliation(s)
- Rogério Lopes dos Santos
- LAMBE, Université d'Evry Val d'Essonne, CNRS, CEA, Université Paris-Saclay, 91025 Evry-Courcouronnes, France
| | - Clément Campillo
- LAMBE, Université d'Evry Val d'Essonne, CNRS, CEA, Université Paris-Saclay, 91025 Evry-Courcouronnes, France
- Institut Universitaire de France (IUF), Paris, France
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6
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Niggel V, Hsu CP, Isa L. Dynamically shaping the surface of silica colloids. SOFT MATTER 2022; 18:7794-7803. [PMID: 36193704 DOI: 10.1039/d2sm00842d] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/16/2023]
Abstract
Surface roughness is an important design parameter to influence the processing of particle-based materials. Current methods to synthesize rough particles present some limitations, e.g. low yield, relative methodological complexity, requirements of multiple steps, or poor roughness control. Here, we thoroughly investigate a facile synthesis route where two silanes, tetraethyl orthosilicate (TEOS) and vinyltrimethoxysilane (VTMS), are added in one pot to form silica particles with controlled corrugated surfaces. We first show that the morphology of these particles can be defined by regulating the amount and ratio of the two silane precursors and by adjusting the concentration of ammonia during synthesis. We characterize the surface topography of the particles using atomic force microscopy and show a direct correlation between surface roughness and the synthesis conditions. Furthermore, we carry out an in situ observation of the evolution of surface morphology and propose a mechanism for surface structuring that hinges on the formation of silane droplets, followed by the preferential hydrolysis/condensation reaction of VTMS starting from the droplet surface and evolving towards the center. The exchange of liquid from the droplets through the VTMS shell leads to stress accumulation and wrinkling/buckling of the particles. Moreover, we explicitly show that osmotic imbalances between the inside and the outside of the droplets regulate their shrinking. We therefore demonstrate that exchanging solvents has a comparable impact to adjusting silane and ammonia content in defining the particle shape and that this synthesis route is highly dynamical. Finally, we demonstrate that it is possible to incorporate fluorescent dyes during synthesis to enable future studies on the impact of surface roughness on dynamic processes, including shear, via direct high-resolution imaging. Our findings show that the mechanism for wrinkling and buckling in colloidal silica particles follows a general scheme found in a broad range of systems, from liposomes and polymeric capsules to Pickering emulsion droplets.
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Affiliation(s)
- Vincent Niggel
- Laboratory for Soft Materials and Interfaces, Department of Materials, ETH Zurich, CH-8093, Zurich, Switzerland.
| | - Chiao-Peng Hsu
- Laboratory for Soft Materials and Interfaces, Department of Materials, ETH Zurich, CH-8093, Zurich, Switzerland.
| | - Lucio Isa
- Laboratory for Soft Materials and Interfaces, Department of Materials, ETH Zurich, CH-8093, Zurich, Switzerland.
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7
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Alam SB, Soligno G, Yang J, Bustillo KC, Ercius P, Zheng H, Whitelam S, Chan EM. Dynamics of Polymer Nanocapsule Buckling and Collapse Revealed by In Situ Liquid-Phase TEM. LANGMUIR : THE ACS JOURNAL OF SURFACES AND COLLOIDS 2022; 38:7168-7178. [PMID: 35621188 DOI: 10.1021/acs.langmuir.2c00432] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/15/2023]
Abstract
Nanocapsules are hollow nanoscale shells that have applications in drug delivery, batteries, self-healing materials, and as model systems for naturally occurring shell geometries. In many applications, nanocapsules are designed to release their cargo as they buckle and collapse, but the details of this transient buckling process have not been directly observed. Here, we use in situ liquid-phase transmission electron microscopy to record the electron-irradiation-induced buckling in spherical 60-187 nm polymer capsules with ∼3.5 nm walls. We observe in real time the release of aqueous cargo from these nanocapsules and their buckling into morphologies with single or multiple indentations. The in situ buckling of nanoscale capsules is compared to ex situ measurements of collapsed and micrometer-sized capsules and to Monte Carlo (MC) simulations. The shape and dynamics of the collapsing nanocapsules are consistent with MC simulations, which reveal that the excessive wrinkling of nanocapsules with ultrathin walls results from their large Föppl-von Kármán numbers around 105. Our experiments suggest design rules for nanocapsules with the desired buckling response based on parameters such as capsule radius, wall thickness, and collapse rate.
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Affiliation(s)
- Sardar B Alam
- The Molecular Foundry, Lawrence Berkeley National Laboratory, Berkeley, California 94720, United States
- Materials Sciences Division, Lawrence Berkeley National Laboratory, Berkeley, California 94720, United States
| | - Giuseppe Soligno
- The Molecular Foundry, Lawrence Berkeley National Laboratory, Berkeley, California 94720, United States
- Debye Institute for Nanomaterials Science, Utrecht University, Utrecht 3584 CC, The Netherlands
| | - Jiwoong Yang
- Materials Sciences Division, Lawrence Berkeley National Laboratory, Berkeley, California 94720, United States
| | - Karen C Bustillo
- The Molecular Foundry, Lawrence Berkeley National Laboratory, Berkeley, California 94720, United States
| | - Peter Ercius
- The Molecular Foundry, Lawrence Berkeley National Laboratory, Berkeley, California 94720, United States
| | - Haimei Zheng
- Materials Sciences Division, Lawrence Berkeley National Laboratory, Berkeley, California 94720, United States
- Department of Materials Science and Engineering, University of California, Berkeley, Berkeley, California 94720, United States
| | - Stephen Whitelam
- The Molecular Foundry, Lawrence Berkeley National Laboratory, Berkeley, California 94720, United States
| | - Emory M Chan
- The Molecular Foundry, Lawrence Berkeley National Laboratory, Berkeley, California 94720, United States
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8
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Preparation of a deformable nanocapsule by living radical polymerization in a liposome. Polym J 2022. [DOI: 10.1038/s41428-022-00632-0] [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]
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9
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Binysh J, Wilks TR, Souslov A. Active elastocapillarity in soft solids with negative surface tension. SCIENCE ADVANCES 2022; 8:eabk3079. [PMID: 35275714 PMCID: PMC8916726 DOI: 10.1126/sciadv.abk3079] [Citation(s) in RCA: 6] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Figures] [Subscribe] [Scholar Register] [Received: 07/05/2021] [Accepted: 01/20/2022] [Indexed: 06/14/2023]
Abstract
Active solids consume energy to allow for actuation, shape change, and wave propagation not possible in equilibrium. Whereas active interfaces have been realized across many experimental systems, control of three-dimensional (3D) bulk materials remains a challenge. Here, we develop continuum theory and microscopic simulations that describe a 3D soft solid whose boundary experiences active surface stresses. The competition between active boundary and elastic bulk yields a broad range of previously unexplored phenomena, which are demonstrations of so-called active elastocapillarity. In contrast to thin shells and vesicles, we discover that bulk 3D elasticity controls snap-through transitions between different anisotropic shapes. These transitions meet at a critical point, allowing a universal classification via Landau theory. In addition, the active surface modifies elastic wave propagation to allow zero, or even negative, group velocities. These phenomena offer robust principles for programming shape change and functionality into active solids, from robotic metamaterials down to shape-shifting nanoparticles.
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Affiliation(s)
- Jack Binysh
- Department of Physics, University of Bath, Claverton Down, Bath BA2 7AY, UK
| | - Thomas R. Wilks
- School of Chemistry, University of Birmingham, Edgbaston, Birmingham B15 2TT, UK
- Exact Sciences Innovation, Sherard Building, Edmund Halley Road, Oxford OX4 4DQ, UK
| | - Anton Souslov
- Department of Physics, University of Bath, Claverton Down, Bath BA2 7AY, UK
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10
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Cheung BCH, Hodgson L, Segall JE, Wu M. Spatial and temporal dynamics of RhoA activities of single breast tumor cells in a 3D environment revealed by a machine learning-assisted FRET technique. Exp Cell Res 2022; 410:112939. [PMID: 34813733 PMCID: PMC8714707 DOI: 10.1016/j.yexcr.2021.112939] [Citation(s) in RCA: 2] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/10/2021] [Revised: 11/15/2021] [Accepted: 11/16/2021] [Indexed: 01/17/2023]
Abstract
One of the hallmarks of cancer cells is their exceptional ability to migrate within the extracellular matrix (ECM) for gaining access to the circulatory system, a critical step of cancer metastasis. RhoA, a small GTPase, is known to be a key molecular switch that toggles between actomyosin contractility and lamellipodial protrusion during cell migration. Current understanding of RhoA activity in cell migration has been largely derived from studies of cells plated on a two-dimensional (2D) substrate using a FRET biosensor. There has been increasing evidence that cells behave differently in a more physiologically relevant three-dimensional (3D) environment. However, studies of RhoA activities in 3D have been hindered by low signal-to-noise ratio in fluorescence imaging. In this paper, we present a a machine learning-assisted FRET technique to follow the spatiotemporal dynamics of RhoA activities of single breast tumor cells (MDA-MB-231) migrating in a 3D as well as a 2D environment. We found that RhoA activity is more polarized along the long axis of the cell for single cells migrating on 2D fibronectin-coated glass versus those embedded in 3D collagen matrices. In particular, RhoA activities of cells in 2D exhibit a distinct front-to-back and back-to-front movement during migration in contrast to those in 3D. Finally, regardless of dimensionality, RhoA polarization is found to be moderately correlated with cell shape.
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Affiliation(s)
- Brian CH Cheung
- Department of Biological and Environmental Engineering, Cornell University, Ithaca, NY, USA
| | - Louis Hodgson
- Department of Anatomy and Structural Biology, Albert Einstein College of Medicine, Bronx, NY, USA,Gruss-Lipper Biophotonics Center, Albert Einstein College of Medicine, Bronx, NY, USA
| | - Jeffrey E Segall
- Department of Anatomy and Structural Biology, Albert Einstein College of Medicine, Bronx, NY, USA
| | - Mingming Wu
- Department of Biological and Environmental Engineering, Cornell University, Ithaca, NY, USA,Corresponding author:
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11
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Ganar KA, Honaker LW, Deshpande S. Shaping synthetic cells through cytoskeleton-condensate-membrane interactions. Curr Opin Colloid Interface Sci 2021. [DOI: 10.1016/j.cocis.2021.101459] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/24/2022]
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12
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Franquelim HG, Dietz H, Schwille P. Reversible membrane deformations by straight DNA origami filaments. SOFT MATTER 2021; 17:276-287. [PMID: 32406895 DOI: 10.1039/d0sm00150c] [Citation(s) in RCA: 29] [Impact Index Per Article: 9.7] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 05/24/2023]
Abstract
Membrane-active cytoskeletal elements, such as FtsZ, septin or actin, form filamentous polymers able to induce and stabilize curvature on cellular membranes. In order to emulate the characteristic dynamic self-assembly properties of cytoskeletal subunits in vitro, biomimetic synthetic scaffolds were here developed using DNA origami. In contrast to our earlier work with pre-curved scaffolds, we specifically assessed the potential of origami mimicking straight filaments, such as actin and microtubules, by origami presenting cholesteryl anchors for membrane binding and additional blunt end stacking interactions for controllable polymerization into linear filaments. By assessing the interaction of our DNA nanostructures with model membranes using fluorescence microscopy, we show that filaments can be formed, upon increasing MgCl2 in solution, for structures displaying blunt ends; and can subsequently depolymerize, by decreasing the concentration of MgCl2. Distinctive spike-like membrane protrusions were generated on giant unilamellar vesicles at high membrane-bound filament densities, and the presence of such deformations was reversible and shown to correlate with the MgCl2-triggered polymerization of DNA origami subunits into filamentous aggregates. In the end, our approach reveals the formation of membrane-bound filaments as a minimal requirement for membrane shaping by straight cytoskeletal-like objects.
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Affiliation(s)
| | - Hendrik Dietz
- Technical University of Munich, Garching Near Munich, Germany
| | - Petra Schwille
- Max Planck Institute of Biochemistry, Martinsried near Munich, Germany.
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13
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Bagatolli LA, Stock RP, Olsen LF. Coupled Response of Membrane Hydration with Oscillating Metabolism in Live Cells: An Alternative Way to Modulate Structural Aspects of Biological Membranes? Biomolecules 2019; 9:E687. [PMID: 31684090 PMCID: PMC6921054 DOI: 10.3390/biom9110687] [Citation(s) in RCA: 10] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/01/2019] [Revised: 10/29/2019] [Accepted: 10/30/2019] [Indexed: 12/12/2022] Open
Abstract
We propose that active metabolic processes may regulate structural changes in biological membranes via the physical state of cell water. This proposition is based on recent results obtained from our group in yeast cells displaying glycolytic oscillations, where we demonstrated that there is a tight coupling between the oscillatory behavior of glycolytic metabolites (ATP, NADH) and the extent of the dipolar relaxation of intracellular water, which oscillates synchronously. The mechanism we suggest involves the active participation of a polarized intracellular water network whose degree of polarization is dynamically modulated by temporal ATP fluctuations caused by metabolism with intervention of a functional cytoskeleton, as conceived in the long overlooked association-induction hypothesis (AIH) of Gilbert Ling. Our results show that the polarized state of intracellular water can be propagated from the cytosol to regions containing membranes. Since changes in the extent of the polarization of water impinge on its chemical activity, we hypothesize that metabolism dynamically controls the local structure of cellular membranes via lyotropic effects. This hypothesis offers an alternative way to interpret membrane related phenomena (e.g., changes in local curvature pertinent to endo/exocytosis or dynamical changes in membranous organelle structure, among others) by integrating relevant but mostly overlooked physicochemical characteristics of the cellular milieu.
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Affiliation(s)
- Luis A Bagatolli
- Instituto de Investigación Médica Mercedes y Martín Ferreyra-INIMEC (CONICET)-Universidad Nacional de Córdoba, Friuli 2434, Córdoba 5016, Argentina.
- Departamento de Química Biológica Ranwel Caputto, Facultad de Ciencias Químicas, Universidad Nacional de Córdoba, Córdoba 5000, Argentina.
| | - Roberto P Stock
- MEMPHYS-International and Interdisciplinary Research Network, 5230 Odense, Denmark.
| | - Lars F Olsen
- University of Southern Denmark, Institute for Biochemistry and Molecular Biology, Campusvej 55, 5230 Odense, Denmark.
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