1
|
Penedo M, Miyazawa K, Okano N, Furusho H, Ichikawa T, Alam MS, Miyata K, Nakamura C, Fukuma T. Visualizing intracellular nanostructures of living cells by nanoendoscopy-AFM. SCIENCE ADVANCES 2021; 7:eabj4990. [PMID: 34936434 PMCID: PMC10954033 DOI: 10.1126/sciadv.abj4990] [Citation(s) in RCA: 15] [Impact Index Per Article: 5.0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 06/07/2021] [Accepted: 11/10/2021] [Indexed: 06/14/2023]
Abstract
Atomic force microscopy (AFM) is the only technique that allows label-free imaging of nanoscale biomolecular dynamics, playing a crucial role in solving biological questions that cannot be addressed by other major bioimaging tools (fluorescence or electron microscopy). However, such imaging is possible only for systems either extracted from cells or reconstructed on solid substrates. Thus, nanodynamics inside living cells largely remain inaccessible with the current nanoimaging techniques. Here, we overcome this limitation by nanoendoscopy-AFM, where a needle-like nanoprobe is inserted into a living cell, presenting actin fiber three-dimensional (3D) maps, and 2D nanodynamics of the membrane inner scaffold, resulting in undetectable changes in cell viability. Unlike previous AFM methods, the nanoprobe directly accesses the target intracellular components, exploiting all the AFM capabilities, such as high-resolution imaging, nanomechanical mapping, and molecular recognition. These features should greatly expand the range of intracellular structures observable in living cells.
Collapse
Affiliation(s)
- Marcos Penedo
- Nano Life Science Institute (WPI-NanoLSI), Kanazawa University, Kanazawa 920-1192, Japan
| | - Keisuke Miyazawa
- Nano Life Science Institute (WPI-NanoLSI), Kanazawa University, Kanazawa 920-1192, Japan
- Division of Electric Engineering and Computer Science, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japan
- Faculty of Frontier Engineering, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japan
| | - Naoko Okano
- Nano Life Science Institute (WPI-NanoLSI), Kanazawa University, Kanazawa 920-1192, Japan
| | - Hirotoshi Furusho
- Nano Life Science Institute (WPI-NanoLSI), Kanazawa University, Kanazawa 920-1192, Japan
| | - Takehiko Ichikawa
- Nano Life Science Institute (WPI-NanoLSI), Kanazawa University, Kanazawa 920-1192, Japan
| | - Mohammad Shahidul Alam
- Division of Nano Life Science, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japan
| | - Kazuki Miyata
- Nano Life Science Institute (WPI-NanoLSI), Kanazawa University, Kanazawa 920-1192, Japan
- Division of Electric Engineering and Computer Science, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japan
- Faculty of Frontier Engineering, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japan
- Division of Nano Life Science, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japan
| | - Chikashi Nakamura
- AIST-INDIA Diverse Assets and Applications International Laboratory (DAILAB), Cellular and Molecular Biotechnology Research Institute (CMB), National Institute of Advanced Industrial Science and Technology (AIST), Tsukuba, Ibaraki 305-8565, Japan
| | - Takeshi Fukuma
- Nano Life Science Institute (WPI-NanoLSI), Kanazawa University, Kanazawa 920-1192, Japan
- Division of Electric Engineering and Computer Science, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japan
- Faculty of Frontier Engineering, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japan
- Division of Nano Life Science, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japan
| |
Collapse
|
2
|
Akisaka T. Platinum replicas of broken-open osteoclasts imaged by transmission electron microscopy. J Oral Biosci 2021; 63:307-318. [PMID: 34628004 DOI: 10.1016/j.job.2021.09.006] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/23/2021] [Revised: 09/02/2021] [Accepted: 09/18/2021] [Indexed: 10/24/2022]
Abstract
BACKGROUND Preserving the cellular structure at the highest possible resolution is a prerequisite for morphological studies to deepen our understanding of cellular functions. A revival of interest in rapid-freezing methods combined with breaking-open techniques has taken place with the development of effective and informative approaches in platinum replica electron microscopy, thus providing new approaches to address unresolved issues in cell biology. HIGHLIGHT The images produced with platinum replicas revealed 3D structures of the cell interior: (1) cell membranes associated with highly organized cytoskeletons, including podosomes or geodomes, (2) heterogeneous clathrin assemblies and membrane skeletons on the inner side of the membrane, and (3) organization of the cytoskeleton after detergent extraction. CONCLUSION In this review, I will focus on the platinum replica method after brokenopen cells have been broken open with mechanical shearing or detergent extraction. Often forgotten nowadays is the use of platinum replicas with stereomicroscopic observations for transmission electron microscopy study; these "old-fashioned" imaging techniques, combined with the breaking-open technique represent a highly informative approach to deepen our understanding of the organization of the cell interior. These are still being pursued to answer outstanding biological questions.
Collapse
Affiliation(s)
- Toshitaka Akisaka
- Department of Oral Anatomy and Neurobiology, Graduate School of Dentistry, Osaka University, Suita, Osaka, 565-0871, Japan.
| |
Collapse
|
3
|
Choi MH, Jeong S, Wang Y, Cho SJ, Park SI, Ye X, Baker LA. Characterization of Ligand Adsorption at Individual Gold Nanocubes. LANGMUIR : THE ACS JOURNAL OF SURFACES AND COLLOIDS 2021; 37:7701-7711. [PMID: 34143943 DOI: 10.1021/acs.langmuir.1c00694] [Citation(s) in RCA: 6] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/12/2023]
Abstract
Cetyltrimethylammonium bromide (CTAB) is a widely used surfactant that aids the aqueous synthesis of colloidal nanoparticles. However, the presence of residual CTAB on nanoparticle surfaces can significantly impact nanoparticle applications, such as catalysis and sensing, under hydrated conditions. As such, consideration of the presence and quantity of CTAB on nanoparticle surfaces under hydrated conditions is of significance. Herein, as part of an integrated material characterization framework, we demonstrate the feasibility of in situ atomic force microscopy (AFM) to detect CTAB on the surface of Au nanocubes (Au NCs) under hydrated conditions, which enabled superior characterization compared to conventional spectroscopic methods. In situ force-distance (FD) spectroscopy and Kelvin probe force microscopy (KPFM) measurements support additional characterization of adsorbed CTAB, while correlative in situ AFM and scanning electron microscopy (SEM) measurements were used to evaluate sequential steps of CTAB removal from Au NCs across hydrated and dehydrated environments, respectively. Notably, a substantial quantity of CTAB remained on the Au NC surface after methanol washing, which was detected in AFM measurements but was not detected in infrared spectroscopy measurements. Subsequent electrochemical cleaning was found to be critically important to remove CTAB from the Au NC surface. Correlative measurements were also performed on individual nanoparticles, which further validate the method described here as a powerful tool to determine the extent and degree of CTAB removal from nanoparticle surfaces. This AFM-based method is broadly applicable to characterize the presence and removal of ligands from nanomaterial surfaces under hydrated conditions.
Collapse
Affiliation(s)
- Myung-Hoon Choi
- Department of Chemistry, Indiana University, 800 E. Kirkwood Avenue, Bloomington, Indiana 47405, United States
| | - Soojin Jeong
- Department of Chemistry, Indiana University, 800 E. Kirkwood Avenue, Bloomington, Indiana 47405, United States
| | - Yi Wang
- Department of Chemistry, Indiana University, 800 E. Kirkwood Avenue, Bloomington, Indiana 47405, United States
| | - Sang-Joon Cho
- Park Systems Corporation, KANC 4F, Gwanggyo-ro 109, Suwon 16229, Korea
| | - Sang-Il Park
- Park Systems Corporation, KANC 4F, Gwanggyo-ro 109, Suwon 16229, Korea
| | - Xingchen Ye
- Department of Chemistry, Indiana University, 800 E. Kirkwood Avenue, Bloomington, Indiana 47405, United States
| | - Lane A Baker
- Department of Chemistry, Indiana University, 800 E. Kirkwood Avenue, Bloomington, Indiana 47405, United States
| |
Collapse
|
4
|
Morone N, Usukura E, Narita A, Usukura J. Improved unroofing protocols for cryo-electron microscopy, atomic force microscopy and freeze-etching electron microscopy and the associated mechanisms. Microscopy (Oxf) 2020; 69:350-359. [PMID: 32447402 DOI: 10.1093/jmicro/dfaa028] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/10/2020] [Revised: 05/05/2020] [Accepted: 05/18/2020] [Indexed: 11/13/2022] Open
Abstract
Unroofing, which is the mechanical shearing of a cell to expose the cytoplasmic surface of the cell membrane, is a unique preparation method that allows membrane cytoskeletons to be observed by cryo-electron microscopy, atomic force microscopy, freeze-etching electron microscopy and other methods. Ultrasound and adhesion have been known to mechanically unroof cells. In this study, unroofing using these two means was denoted sonication unroofing and adhesion unroofing, respectively. We clarified the mechanisms by which cell membranes are removed in these unroofing procedures and established efficient protocols for each based on the mechanisms. In sonication unroofing, fine bubbles generated by sonication adhered electrostatically to apical cell surfaces and then removed the apical (dorsal) cell membrane with the assistance of buoyancy and water flow. The cytoplasmic surface of the ventral cell membrane remaining on the grids became observable by this method. In adhesion unroofing, grids charged positively by coating with Alcian blue were pressed onto the cells, thereby tightly adsorbing the dorsal cell membrane. Subsequently, a part of the cell membrane strongly adhered to the grids was peeled from the cells and transferred onto the grids when the grids were lifted. This method thus allowed the visualization of the cytoplasmic surface of the dorsal cell membrane. This paper describes robust, improved protocols for the two unroofing methods in detail. In addition, micro-unroofing (perforation) likely due to nanobubbles is introduced as a new method to make cells transparent to electron beams.
Collapse
Affiliation(s)
- Nobuhiro Morone
- Medical Research Council Toxicology Unit, University of Cambridge, Lancaster Road, Leicester LE1 9HN, UK
| | - Eiji Usukura
- Institute of Materials and Systems for Sustainability, Nagoya University, Chikusa-ku, Nagoya 464-8601, Japan
| | - Akihiro Narita
- Structural Biology Research Centre, Graduate School of Science, Nagoya University, Chikusa-ku,Nagoya,464-8601, Japan
| | - Jiro Usukura
- Institute of Materials and Systems for Sustainability, Nagoya University, Chikusa-ku, Nagoya 464-8601, Japan
| |
Collapse
|
5
|
Liang W, Shi H, Yang X, Wang J, Yang W, Zhang H, Liu L. Recent advances in AFM-based biological characterization and applications at multiple levels. SOFT MATTER 2020; 16:8962-8984. [PMID: 32996549 DOI: 10.1039/d0sm01106a] [Citation(s) in RCA: 22] [Impact Index Per Article: 5.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/11/2023]
Abstract
Atomic force microscopy (AFM) has found a wide range of bio-applications in the past few decades due to its ability to measure biological samples in natural environments at a high spatial resolution. AFM has become a key platform in biomedical, bioengineering and drug research fields, enabling mechanical and morphological characterization of live biological systems. Hence, we provide a comprehensive review on recent advances in the use of AFM for characterizing the biomechanical properties of multi-scale biological samples, ranging from molecule, cell to tissue levels. First, we present the fundamental principles of AFM and two AFM-based models for the characterization of biomechanical properties of biological samples, covering key AFM devices and AFM bioimaging as well as theoretical models for characterizing the elasticity and viscosity of biomaterials. Then, we elaborate on a series of new experimental findings through analysis of biomechanics. Finally, we discuss the future directions and challenges. It is envisioned that the AFM technique will enable many remarkable discoveries, and will have far-reaching impacts on bio-related studies and applications in the future.
Collapse
Affiliation(s)
- Wenfeng Liang
- School of Mechanical Engineering, Shenyang Jianzhu University, Shenyang, 110168, China.
| | - Haohao Shi
- School of Mechanical Engineering, Shenyang Jianzhu University, Shenyang, 110168, China.
| | - Xieliu Yang
- School of Mechanical Engineering, Shenyang Jianzhu University, Shenyang, 110168, China.
| | - Junhai Wang
- School of Mechanical Engineering, Shenyang Jianzhu University, Shenyang, 110168, China.
| | - Wenguang Yang
- School of Electromechanical and Automotive Engineering, Yantai University, Yantai 264005, China
| | - Hemin Zhang
- Department of Neurology, The People's Hospital of Liaoning Province, Shenyang 110016, China.
| | - Lianqing Liu
- State Key Laboratory of Robotics, Shenyang Institute of Automation, Chinese Academy of Sciences, Shenyang 110016, China.
| |
Collapse
|
6
|
Svitkina TM. Actin Cell Cortex: Structure and Molecular Organization. Trends Cell Biol 2020; 30:556-565. [PMID: 32278656 PMCID: PMC7566779 DOI: 10.1016/j.tcb.2020.03.005] [Citation(s) in RCA: 99] [Impact Index Per Article: 24.8] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/21/2020] [Revised: 03/11/2020] [Accepted: 03/19/2020] [Indexed: 12/20/2022]
Abstract
The actin cytoskeleton consists of structurally and biochemically different actin filament arrays. Among them, the actin cortex is thought to have key roles in cell mechanics, but remains a poorly characterized part of the actin cytoskeleton. The cell cortex is typically defined as a thin layer of actin meshwork that uniformly underlies the plasma membrane of the entire cell. However, this definition applies only to specific cases. In general, the cortex structure and subcellular distribution vary significantly across cell types and physiological states of the cell. In this review, I focus on our current knowledge of the structure and molecular composition of the cell cortex.
Collapse
Affiliation(s)
- Tatyana M Svitkina
- Department of Biology, School of Arts and Sciences, University of Pennsylvania, Philadelphia, PA, USA.
| |
Collapse
|
7
|
Liu L, Wei Y, Liu J, Wang K, Zhang J, Zhang P, Zhou Y, Li B. Spatial high resolution of actin filament organization by PeakForce atomic force microscopy. Cell Prolif 2019; 53:e12670. [PMID: 31568631 PMCID: PMC6985672 DOI: 10.1111/cpr.12670] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/17/2019] [Revised: 05/08/2019] [Accepted: 05/16/2019] [Indexed: 12/12/2022] Open
Abstract
Objectives To investigate the heterogeneous feature of actin filaments (ACFs) associated with the cellular membrane in HeLa and HCT‐116 cells at the nanoscale level. Materials and Methods Fluorescence microscopy coupled with atomic force microscopy (AFM) was used to identify and characterize ACFs of cells. The distribution of ACFs was detected by Fluor‐488‐phalloidin–labelled actin. The morphology of the ACFs was probed by AFM images. The spatial correlation of the microvilli and ACFs was explored with different forces of AFM loading on cells. Results Intricate but ordered structures of the actin cytoskeletons associated with cellular membrane were characterized and revealed. Two different layers of ACFs with distinct structural organizations were directly observed in HCT‐116 and HeLa cells. Bundle‐shaped ACFs protruding the cellular membrane forming the microvilli, and the network ACFs underneath the cellular membrane were resolved with high resolution under near‐physiological conditions. Approximately 14 nm lateral resolution was achieved when imaging single ACF beneath the cellular membrane. On the basis of the observed spatial distribution of the ultrastructure of the ACF organization, a model for this organization of ACFs was proposed. Conclusions We revealed the two layers of the ACF organization in Hela and HCT‐116 cells. The resolved heterogeneous structures at the nanoscale level provide a spatial view of the ACFs, which would contribute to the understanding of the essential biological functions of the actin cytoskeleton.
Collapse
Affiliation(s)
- Lin Liu
- Division of Physical Biology & Bioimaging Centre, Shanghai Synchrotron Radiation Facility, CAS Key Laboratory of Interfacial Physics and Technology, Shanghai Institute of Applied Physics, Chinese Academy of Sciences, Shanghai, China.,University of Chinese Academy of Sciences, Beijing, China
| | - Yuhui Wei
- Division of Physical Biology & Bioimaging Centre, Shanghai Synchrotron Radiation Facility, CAS Key Laboratory of Interfacial Physics and Technology, Shanghai Institute of Applied Physics, Chinese Academy of Sciences, Shanghai, China
| | - Jingyuan Liu
- Fourth Military Medical University, Xi'an, China
| | - Kaizhe Wang
- Division of Physical Biology & Bioimaging Centre, Shanghai Synchrotron Radiation Facility, CAS Key Laboratory of Interfacial Physics and Technology, Shanghai Institute of Applied Physics, Chinese Academy of Sciences, Shanghai, China.,University of Chinese Academy of Sciences, Beijing, China
| | - Jinjin Zhang
- Division of Physical Biology & Bioimaging Centre, Shanghai Synchrotron Radiation Facility, CAS Key Laboratory of Interfacial Physics and Technology, Shanghai Institute of Applied Physics, Chinese Academy of Sciences, Shanghai, China
| | - Ping Zhang
- Division of Physical Biology & Bioimaging Centre, Shanghai Synchrotron Radiation Facility, CAS Key Laboratory of Interfacial Physics and Technology, Shanghai Institute of Applied Physics, Chinese Academy of Sciences, Shanghai, China.,University of Chinese Academy of Sciences, Beijing, China
| | - Yi Zhou
- School of Basic Medicine, Chengdu University of Traditional Chinese Medicine, Chengdu, China
| | - Bin Li
- Division of Physical Biology & Bioimaging Centre, Shanghai Synchrotron Radiation Facility, CAS Key Laboratory of Interfacial Physics and Technology, Shanghai Institute of Applied Physics, Chinese Academy of Sciences, Shanghai, China
| |
Collapse
|
8
|
Janel S, Popoff M, Barois N, Werkmeister E, Divoux S, Perez F, Lafont F. Stiffness tomography of eukaryotic intracellular compartments by atomic force microscopy. NANOSCALE 2019; 11:10320-10328. [PMID: 31106790 DOI: 10.1039/c8nr08955h] [Citation(s) in RCA: 12] [Impact Index Per Article: 2.4] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/09/2023]
Abstract
Precise localization and biophysical characterization of cellular structures is a key to the understanding of biological processes happening both inside the cell and at the cell surface. Atomic force microscopy is a powerful tool to study the cell surface - topography, elasticity, viscosity, interactions - and also the viscoelastic behavior of the underlying cytoplasm, cytoskeleton or the nucleus. Here, we demonstrate the ability of atomic force microscopy to also map and characterize organelles and microorganisms inside cells, at the nanoscale, by combining stiffness tomography with super-resolution fluorescence and electron microscopy. By using this correlative approach, we could both identify and characterize intracellular compartments. The validation of this approach was performed by monitoring the stiffening effect according to the metabolic status of the mitochondria in living cells in real-time.
Collapse
Affiliation(s)
- Sébastien Janel
- Center for Infection and Immunity of Lille, CNRS UMR 8204, INSERM U1019, CHU Lille, Institut Pasteur de Lille, Univ Lille, F-59000 Lille, France.
| | | | | | | | | | | | | |
Collapse
|
9
|
Puljung M, Vedovato N, Usher S, Ashcroft F. Activation mechanism of ATP-sensitive K + channels explored with real-time nucleotide binding. eLife 2019; 8:41103. [PMID: 30789344 PMCID: PMC6400584 DOI: 10.7554/elife.41103] [Citation(s) in RCA: 23] [Impact Index Per Article: 4.6] [Reference Citation Analysis] [Abstract] [Key Words] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/14/2018] [Accepted: 02/14/2019] [Indexed: 01/15/2023] Open
Abstract
The response of ATP-sensitive K+ channels (KATP) to cellular metabolism is coordinated by three classes of nucleotide binding site (NBS). We used a novel approach involving labeling of intact channels in a native, membrane environment with a non-canonical fluorescent amino acid and measurement (using FRET with fluorescent nucleotides) of steady-state and time-resolved nucleotide binding to dissect the role of NBS2 of the accessory SUR1 subunit of KATP in channel gating. Binding to NBS2 was Mg2+-independent, but Mg2+ was required to trigger a conformational change in SUR1. Mutation of a lysine (K1384A) in NBS2 that coordinates bound nucleotides increased the EC50 for trinitrophenyl-ADP binding to NBS2, but only in the presence of Mg2+, indicating that this mutation disrupts the ligand-induced conformational change. Comparison of nucleotide-binding with ionic currents suggests a model in which each nucleotide binding event to NBS2 of SUR1 is independent and promotes KATP activation by the same amount.
Collapse
Affiliation(s)
- Michael Puljung
- Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom
| | - Natascia Vedovato
- Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom
| | - Samuel Usher
- Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom
| | - Frances Ashcroft
- Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom
| |
Collapse
|
10
|
Abstract
Atomic force microscopy (AFM) is a very versatile tool for studying biological samples at nanometer-scale resolution. The cell membrane plays a key role in compartmentalization, nutrient transportation, and signal transduction, while the structural feature of both sides of the membrane remains elusive. Here we describe our methods for the preparation of the cell membrane from the red blood cells and nucleated cells. High-resolution AFM topographs reveal substructural details of both sides of the cell membrane. The structure composition of cell membrane can be directly observed by time-lapse AFM and the positional information of membrane proteins can be located by molecular recognition.
Collapse
Affiliation(s)
- Mingjun Cai
- Changchun Institute of Applied Chemistry, Chinese Academy of Sciences, Changchun, Jilin, People's Republic of China
| | - Hongda Wang
- Changchun Institute of Applied Chemistry, Chinese Academy of Sciences, Changchun, Jilin, People's Republic of China.
| |
Collapse
|
11
|
Galvanetto N. Single-cell unroofing: probing topology and nanomechanics of native membranes. BIOCHIMICA ET BIOPHYSICA ACTA-BIOMEMBRANES 2018; 1860:2532-2538. [DOI: 10.1016/j.bbamem.2018.09.019] [Citation(s) in RCA: 5] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Track Full Text] [Subscribe] [Scholar Register] [Received: 06/22/2018] [Revised: 09/19/2018] [Accepted: 09/26/2018] [Indexed: 10/28/2022]
|
12
|
Basoli F, Giannitelli SM, Gori M, Mozetic P, Bonfanti A, Trombetta M, Rainer A. Biomechanical Characterization at the Cell Scale: Present and Prospects. Front Physiol 2018; 9:1449. [PMID: 30498449 PMCID: PMC6249385 DOI: 10.3389/fphys.2018.01449] [Citation(s) in RCA: 41] [Impact Index Per Article: 6.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/01/2018] [Accepted: 09/24/2018] [Indexed: 12/12/2022] Open
Abstract
The rapidly growing field of mechanobiology demands for robust and reproducible characterization of cell mechanical properties. Recent achievements in understanding the mechanical regulation of cell fate largely rely on technological platforms capable of probing the mechanical response of living cells and their physico–chemical interaction with the microenvironment. Besides the established family of atomic force microscopy (AFM) based methods, other approaches include optical, magnetic, and acoustic tweezers, as well as sensing substrates that take advantage of biomaterials chemistry and microfabrication techniques. In this review, we introduce the available methods with an emphasis on the most recent advances, and we discuss the challenges associated with their implementation.
Collapse
Affiliation(s)
- Francesco Basoli
- Department of Engineering, Università Campus Bio-Medico di Roma, Rome, Italy
| | | | - Manuele Gori
- Department of Engineering, Università Campus Bio-Medico di Roma, Rome, Italy
| | - Pamela Mozetic
- Center for Translational Medicine, International Clinical Research Center, St. Anne's University Hospital, Brno, Czechia
| | - Alessandra Bonfanti
- Department of Engineering, University of Cambridge, Cambridge, United Kingdom
| | - Marcella Trombetta
- Department of Engineering, Università Campus Bio-Medico di Roma, Rome, Italy
| | - Alberto Rainer
- Department of Engineering, Università Campus Bio-Medico di Roma, Rome, Italy.,Institute for Photonics and Nanotechnologies, National Research Council, Rome, Italy
| |
Collapse
|
13
|
A Cryosectioning Technique for the Observation of Intracellular Structures and Immunocytochemistry of Tissues in Atomic Force Microscopy (AFM). Sci Rep 2017; 7:6462. [PMID: 28743939 PMCID: PMC5526917 DOI: 10.1038/s41598-017-06942-1] [Citation(s) in RCA: 7] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/23/2017] [Accepted: 07/03/2017] [Indexed: 02/07/2023] Open
Abstract
The use of cryosectioning facilitates the morphological analysis and immunocytochemistry of cells in tissues in atomic force microscopy (AFM). The cantilever can access all parts of a tissue sample in cryosections after the embedding medium (sucrose) has been replaced with phosphate-buffered saline (PBS), and this approach has enabled the production of a type of high-resolution image. The images resembled those obtained from freeze-etching replica electron microscopy (EM) rather than from thin-section EM. The AFM images showed disks stacked and enveloped by the cell membrane in rod photoreceptor outer segments (ROS) at EM resolution. In addition, ciliary necklaces on the surface of connecting cilium, three-dimensional architecture of synaptic ribbons, and the surface of the post-synaptic membrane facing the active site were revealed, which were not apparent using thin-section EM. AFM could depict the molecular binding of anti-opsin antibodies conjugated to a secondary fluorescent antibody bound to the disk membrane. The specific localization of the anti-opsin binding sites was verified through correlation with immunofluorescence signals in AFM combined with confocal fluorescence microscope. To prove reproducibility in other tissues besides retina, cryosectioning-AFM was also applied to elucidate molecular organization of sarcomere in a rabbit psoas muscle.
Collapse
|
14
|
Usukura E, Narita A, Yagi A, Ito S, Usukura J. An Unroofing Method to Observe the Cytoskeleton Directly at Molecular Resolution Using Atomic Force Microscopy. Sci Rep 2016; 6:27472. [PMID: 27273367 PMCID: PMC4895337 DOI: 10.1038/srep27472] [Citation(s) in RCA: 25] [Impact Index Per Article: 3.1] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/22/2016] [Accepted: 05/19/2016] [Indexed: 11/09/2022] Open
Abstract
An improved unroofing method enabled the cantilever of an atomic force microscope (AFM) to reach directly into a cell to visualize the intracellular cytoskeletal actin filaments, microtubules, clathrin coats, and caveolae in phosphate-buffered saline (PBS) at a higher resolution than conventional electron microscopy. All of the actin filaments clearly exhibited a short periodicity of approximately 5-6 nm, which was derived from globular actins linked to each other to form filaments, as well as a long helical periodicity. The polarity of the actin filaments appeared to be determined by the shape of the periodic striations. Microtubules were identified based on their thickness. Clathrin coats and caveolae were observed on the cytoplasmic surface of cell membranes. The area containing clathrin molecules and their terminal domains was directly visualized. Characteristic ridge structures located at the surface of the caveolae were observed at high resolution, similar to those observed with electron microscopy (EM). Overall, unroofing allowed intracellular AFM imaging in a liquid environment with a level of quality equivalent or superior to that of EM. Thus, AFMs are anticipated to provide cutting-edge findings in cell biology and histology.
Collapse
Affiliation(s)
- Eiji Usukura
- Structural Biology Research Centre, Graduate School of Science, Nagoya University, Nagoya, 464-8603 Japan
| | - Akihiro Narita
- Structural Biology Research Centre, Graduate School of Science, Nagoya University, Nagoya, 464-8603 Japan
| | - Akira Yagi
- Olympus Corporation, Hachioji, Tokyo, 192-8512 Japan
| | - Shuichi Ito
- Olympus Corporation, Hachioji, Tokyo, 192-8512 Japan
| | - Jiro Usukura
- Structural Biology Research Centre, Graduate School of Science, Nagoya University, Nagoya, 464-8603 Japan
| |
Collapse
|
15
|
Narita A, Usukura E, Yagi A, Tateyama K, Akizuki S, Kikumoto M, Matsumoto T, Maéda Y, Ito S, Usukura J. Direct observation of the actin filament by tip-scan atomic force microscopy. Microscopy (Oxf) 2016; 65:370-7. [PMID: 27242058 PMCID: PMC5895109 DOI: 10.1093/jmicro/dfw017] [Citation(s) in RCA: 6] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/23/2015] [Accepted: 04/28/2016] [Indexed: 11/13/2022] Open
Abstract
Actin filaments, the actin–myosin complex and the actin–tropomyosin complex were observed by a tip-scan atomic force microscope (AFM), which was recently developed by Olympus as the AFM part of a correlative microscope. This newly developed AFM uses cantilevers of similar size as stage-scan AFMs to improve substantially the spatial and temporal resolution. Such an approach has previously never been possible by a tip-scan system, in which a cantilever moves in the x, y and z directions. We evaluated the performance of this developed tip-scan AFM by observing the molecular structure of actin filaments and the actin–tropomyosin complex. In the image of the actin filament, the molecular interval of the actin subunits (∼5.5 nm) was clearly observed as stripes. From the shape of the stripes, the polarity of the actin filament was directly determined and the results were consistent with the polarity determined by myosin binding. In the image of the actin–tropomyosin complex, each tropomyosin molecule (∼2 nm in diameter) on the actin filament was directly observed without averaging images of different molecules. Each tropomyosin molecule on the actin filament has never been directly observed by AFM or electron microscopy. Thus, our developed tip-scan AFM offers significant potential in observing purified proteins and cellular structures at nanometer resolution. Current results represent an important step in the development of a new correlative microscope to observe nm-order structures at an acceptable frame rate (∼10 s/frame) by AFM at the position indicated by the fluorescent dye observed under a light microscope.
Collapse
Affiliation(s)
- Akihiro Narita
- Structural Biology Research Center, Graduate School of Science, Nagoya University, Nagoya 464-8602, Japan JST PRESTO, 4-1-8 Honcho, Kawaguchi, Saitama, 332-0012, Japan
| | - Eiji Usukura
- Structural Biology Research Center, Graduate School of Science, Nagoya University, Nagoya 464-8602, Japan
| | - Akira Yagi
- R&D Group, Olympus Corporation, 2-3 Kuboyama-cho, Hachioji, Tokyo 192-8512, Japan
| | - Kiyohiko Tateyama
- R&D Group, Olympus Corporation, 2-3 Kuboyama-cho, Hachioji, Tokyo 192-8512, Japan
| | - Shogo Akizuki
- Structural Biology Research Center, Graduate School of Science, Nagoya University, Nagoya 464-8602, Japan
| | - Mahito Kikumoto
- Structural Biology Research Center, Graduate School of Science, Nagoya University, Nagoya 464-8602, Japan
| | - Tomoharu Matsumoto
- Structural Biology Research Center, Graduate School of Science, Nagoya University, Nagoya 464-8602, Japan
| | - Yuichiro Maéda
- Structural Biology Research Center, Graduate School of Science, Nagoya University, Nagoya 464-8602, Japan
| | - Shuichi Ito
- R&D Group, Olympus Corporation, 2-3 Kuboyama-cho, Hachioji, Tokyo 192-8512, Japan
| | - Jiro Usukura
- Structural Biology Research Center, Graduate School of Science, Nagoya University, Nagoya 464-8602, Japan
| |
Collapse
|
16
|
Kronlage C, Schäfer-Herte M, Böning D, Oberleithner H, Fels J. Feeling for Filaments: Quantification of the Cortical Actin Web in Live Vascular Endothelium. Biophys J 2016; 109:687-98. [PMID: 26287621 PMCID: PMC4547164 DOI: 10.1016/j.bpj.2015.06.066] [Citation(s) in RCA: 39] [Impact Index Per Article: 4.9] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/18/2014] [Revised: 06/07/2015] [Accepted: 06/24/2015] [Indexed: 12/27/2022] Open
Abstract
Contact-mode atomic force microscopy (AFM) has been shown to reveal cortical actin structures. Using live endothelial cells, we visualized cortical actin dynamics simultaneously by AFM and confocal fluorescence microscopy. We present a method that quantifies dynamic changes in the mechanical ultrastructure of the cortical actin web. We argue that the commonly used, so-called error signal imaging in AFM allows a qualitative, but not quantitative, analysis of cortical actin dynamics. The approach we used comprises fast force-curve-based topography imaging and subsequent image processing that enhances local height differences. Dynamic changes in the organization of the cytoskeleton network can be observed and quantified by surface roughness calculations and automated morphometrics. Upon treatment with low concentrations of the actin-destabilizing agent cytochalasin D, the cortical cytoskeleton network is thinned out and the average mesh size increases. In contrast, jasplakinolide, a drug that enhances actin polymerization, consolidates the cytoskeleton network and reduces the average mesh area. In conclusion, cortical actin dynamics can be quantified in live cells. To our knowledge, this opens a new pathway for conducting quantitative structure-function analyses of the endothelial actin web just beneath the apical plasma membrane.
Collapse
Affiliation(s)
| | - Marco Schäfer-Herte
- Institute of Cell Dynamics and Imaging, University of Münster, Münster, Germany
| | - Daniel Böning
- Institute of Medical Physics and Biophysics, University of Münster, Münster, Germany
| | | | - Johannes Fels
- Institute of Physiology II, University of Münster, Münster, Germany; Institute of Cell Dynamics and Imaging, University of Münster, Münster, Germany.
| |
Collapse
|
17
|
Sato F, Asakawa H, Fukuma T, Terada S. Semi-in situ atomic force microscopy imaging of intracellular neurofilaments under physiological conditions through the 'sandwich' method. Microscopy (Oxf) 2016; 65:316-24. [PMID: 26960670 DOI: 10.1093/jmicro/dfw006] [Citation(s) in RCA: 7] [Impact Index Per Article: 0.9] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/16/2015] [Accepted: 02/07/2016] [Indexed: 11/14/2022] Open
Abstract
Neurofilaments are intermediate filament proteins specific for neurons and characterized by formation of biochemically stable, obligate heteropolymers in vivo While purified or reassembled neurofilaments have been subjected to morphological analyses by electron microscopy and atomic force microscopy, there has been a need for direct imaging of cytoplasmic genuine intermediate filaments with minimal risk of artefactualization. In this study, we applied the modified 'cells on glass sandwich' method to exteriorize intracellular neurofilaments, reducing the risk of causing artefacts through sample preparation. SW13vim(-) cells were double transduced with neurofilament medium polypeptide (NF-M) and alpha-internexin (α-inx). Cultured cells were covered with a cationized coverslip after prestabilization with tannic acid to form a sandwich and then split into two. After confirming that neurofilaments could be deposited on ventral plasma membranes exposed via unroofing, we performed atomic force microscopy imaging semi-in situ in aqueous solution. The observed thin filaments, considered to retain native structures of the neurofilaments, exhibited an approximate periodicity of 50-60 nm along their length. Their structural property appeared to reflect the morphology formed by their constituents, i.e. NF-M and α-inx. The success of semi-in situ atomic force microscopy of exposed bona fide assembled neurofilaments through separating the sandwich suggests that it can be an effective and alternative method for investigating cytoplasmic intermediate filaments under physiological conditions by atomic force microscopy.
Collapse
Affiliation(s)
- Fumiya Sato
- Department of Neuroanatomy and Cellular Neurobiology, Graduate School of Medical and Dental Sciences, Tokyo Medical and Dental University, 1-5-45 Yushima, Bunkyo-ku, Tokyo 113-8519, Japan Center for Brain Integration Research, Tokyo Medical and Dental University, 1-5-45 Yushima, Bunkyo-ku, Tokyo 113-8519, Japan Japan Society for the Promotion of Science (DC1), Kojimachi Business Center Building, 5-3-1 Kojimahchi, Chiyoda-ku, Tokyo 102-0083, Japan
| | - Hitoshi Asakawa
- Super-resolution AFM R&D Division, Bio-AFM Frontier Research Center, College of Science and Engineering, Kanazawa University, Kakuma-machi, Kanazawa, Ishikawa 920-1192, Japan
| | - Takeshi Fukuma
- Super-resolution AFM R&D Division, Bio-AFM Frontier Research Center, College of Science and Engineering, Kanazawa University, Kakuma-machi, Kanazawa, Ishikawa 920-1192, Japan Division of Electrical Engineering, Kanazawa University, Kakuma-machi, Kanazawa, Ishikawa 920-1192, Japan
| | - Sumio Terada
- Department of Neuroanatomy and Cellular Neurobiology, Graduate School of Medical and Dental Sciences, Tokyo Medical and Dental University, 1-5-45 Yushima, Bunkyo-ku, Tokyo 113-8519, Japan Center for Brain Integration Research, Tokyo Medical and Dental University, 1-5-45 Yushima, Bunkyo-ku, Tokyo 113-8519, Japan
| |
Collapse
|
18
|
Shan Y, Wang H. The structure and function of cell membranes examined by atomic force microscopy and single-molecule force spectroscopy. Chem Soc Rev 2016; 44:3617-38. [PMID: 25893228 DOI: 10.1039/c4cs00508b] [Citation(s) in RCA: 102] [Impact Index Per Article: 12.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/17/2022]
Abstract
The cell membrane is one of the most complicated biological complexes, and long-term fierce debates regarding the cell membrane persist because of technical hurdles. With the rapid development of nanotechnology and single-molecule techniques, our understanding of cell membranes has substantially increased. Atomic force microscopy (AFM) has provided several unprecedented advances (e.g., high resolution, three-dimensional and in situ measurements) in the study of cell membranes and has been used to systematically dissect the membrane structure in situ from both sides of membranes; as a result, novel models of cell membranes have recently been proposed. This review summarizes the new progress regarding membrane structure using in situ AFM and single-molecule force spectroscopy (SMFS), which may shed light on the study of the structure and functions of cell membranes.
Collapse
Affiliation(s)
- Yuping Shan
- State Key Laboratory of Electroanalytical Chemistry, Changchun Institute of Applied Chemistry, Chinese Academy of Sciences, Changchun, Jilin 130022, P. R. China.
| | | |
Collapse
|
19
|
ROR1 sustains caveolae and survival signalling as a scaffold of cavin-1 and caveolin-1. Nat Commun 2016; 7:10060. [PMID: 26725982 PMCID: PMC4777216 DOI: 10.1038/ncomms10060] [Citation(s) in RCA: 66] [Impact Index Per Article: 8.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/20/2015] [Accepted: 10/28/2015] [Indexed: 01/03/2023] Open
Abstract
The receptor tyrosine kinase-like orphan receptor 1 (ROR1) sustains prosurvival signalling directly downstream of the lineage-survival oncogene NKX2-1/TTF-1 in lung adenocarcinoma. Here we report an unanticipated function of this receptor tyrosine kinase (RTK) as a scaffold of cavin-1 and caveolin-1 (CAV1), two essential structural components of caveolae. This kinase-independent function of ROR1 facilitates the interactions of cavin-1 and CAV1 at the plasma membrane, thereby preventing the lysosomal degradation of CAV1. Caveolae structures and prosurvival signalling towards AKT through multiple RTKs are consequently sustained. These findings provide mechanistic insight into how ROR1 inhibition can overcome EGFR-tyrosine kinase inhibitor (TKI) resistance due to bypass signalling via diverse RTKs such as MET and IGF-IR, which is currently a major clinical obstacle. Considering its onco-embryonic expression, inhibition of the scaffold function of ROR1 in patients with lung adenocarcinoma is an attractive approach for improved treatment of this devastating cancer.
Collapse
|
20
|
Imaging of the subsurface structures of “unroofed” Plasmodium falciparum-infected erythrocytes. Exp Parasitol 2015; 153:174-9. [DOI: 10.1016/j.exppara.2015.03.021] [Citation(s) in RCA: 7] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/14/2014] [Revised: 03/05/2015] [Accepted: 03/20/2015] [Indexed: 01/24/2023]
|
21
|
Mohan J, Morén B, Larsson E, Holst MR, Lundmark R. Cavin3 interacts with cavin1 and caveolin1 to increase surface dynamics of caveolae. J Cell Sci 2015; 128:979-91. [PMID: 25588833 DOI: 10.1242/jcs.161463] [Citation(s) in RCA: 37] [Impact Index Per Article: 4.1] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/20/2022] Open
Abstract
Caveolae are invaginations of the cell surface thought to regulate membrane tension, signalling, adhesion and lipid homeostasis owing to their dynamic behaviour ranging from stable surface association to dynamic rounds of fission and fusion with the plasma membrane. The caveolae coat is generated by oligomerisation of the membrane protein caveolin and the family of cavin proteins. Here, we show that cavin3 (also known as PRKCDBP) is targeted to caveolae by cavin1 (also known as PTRF) where it interacts with the scaffolding domain of caveolin1 and promote caveolae dynamics. We found that the N-terminal region of cavin3 binds a trimer of the cavin1 N-terminus in competition with a homologous cavin2 (also known as SDPR) region, showing that the cavins form distinct subcomplexes through their N-terminal regions. Our data shows that cavin3 is enriched at deeply invaginated caveolae and that loss of cavin3 in cells results in an increase of stable caveolae and a decrease of caveolae that are only present at the membrane for a short time. We propose that cavin3 is recruited to the caveolae coat by cavin1 to interact with caveolin1 and regulate the duration time of caveolae at the plasma membrane.
Collapse
Affiliation(s)
- Jagan Mohan
- Medical Biochemistry and Biophysics, Laboratory for Molecular Infection Medicine Sweden, Umeå University, 901 87, Umeå, Sweden
| | - Björn Morén
- Medical Biochemistry and Biophysics, Laboratory for Molecular Infection Medicine Sweden, Umeå University, 901 87, Umeå, Sweden
| | - Elin Larsson
- Medical Biochemistry and Biophysics, Laboratory for Molecular Infection Medicine Sweden, Umeå University, 901 87, Umeå, Sweden
| | - Mikkel R Holst
- Integrative Medical Biology, Umeå University, 901 87, Umeå, Sweden
| | - Richard Lundmark
- Medical Biochemistry and Biophysics, Laboratory for Molecular Infection Medicine Sweden, Umeå University, 901 87, Umeå, Sweden Integrative Medical Biology, Umeå University, 901 87, Umeå, Sweden
| |
Collapse
|