1
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Wu H, Fujioka Y, Sakaguchi S, Suzuki Y, Nakano T. Morphological analysis for two types of viral particles in vacuoles of SARS-CoV-2-infected cells. Med Mol Morphol 2024; 57:124-135. [PMID: 38393367 DOI: 10.1007/s00795-024-00381-4] [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: 10/27/2023] [Accepted: 01/17/2024] [Indexed: 02/25/2024]
Abstract
In this study, we analyzed the morphological structure of severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) in human cells. We identified the two types of viral particles present within the vacuoles of infected cells. Using transmission electron microscopy, we observed that SARS-CoV-2 particles exhibited both low- and high-electron-density structures, which was further confirmed through three-dimensional reconstruction using electron tomography. The budding of these particles was exclusively observed within these vacuoles. Intriguingly, viral particles with low-electron-density structures were confined to vacuoles, whereas those with high-electron-density structures were found in vacuoles and on the cell membrane surface of infected cells. Notably, high-electron-density particles found within vacuoles exhibited the same morphology as those outside the infected cells. This observation suggests that the two types of viral particles identified in this study had different maturation status. Our findings provide valuable insights into the molecular details of SARS-CoV-2 particles, contributing to our understanding of the virus.
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Affiliation(s)
- Hong Wu
- Department of Microbiology and Infection Control, Faculty of Medicine, Osaka Medical and Pharmaceutical University, 2-7 Daigaku-machi, Takatsuki, Osaka, 569-8686, Japan.
| | - Yoshihiko Fujioka
- Department of Microbiology and Infection Control, Faculty of Medicine, Osaka Medical and Pharmaceutical University, 2-7 Daigaku-machi, Takatsuki, Osaka, 569-8686, Japan
| | - Shoichi Sakaguchi
- Department of Microbiology and Infection Control, Faculty of Medicine, Osaka Medical and Pharmaceutical University, 2-7 Daigaku-machi, Takatsuki, Osaka, 569-8686, Japan
| | - Youichi Suzuki
- Department of Microbiology and Infection Control, Faculty of Medicine, Osaka Medical and Pharmaceutical University, 2-7 Daigaku-machi, Takatsuki, Osaka, 569-8686, Japan
| | - Takashi Nakano
- Department of Microbiology and Infection Control, Faculty of Medicine, Osaka Medical and Pharmaceutical University, 2-7 Daigaku-machi, Takatsuki, Osaka, 569-8686, Japan
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2
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Yang X, Wang L, Smaluk V, Shaftan T. Optimize Electron Beam Energy toward In Situ Imaging of Thick Frozen Bio-Samples with Nanometer Resolution Using MeV-STEM. NANOMATERIALS (BASEL, SWITZERLAND) 2024; 14:803. [PMID: 38727397 PMCID: PMC11085184 DOI: 10.3390/nano14090803] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Subscribe] [Scholar Register] [Received: 04/02/2024] [Revised: 04/21/2024] [Accepted: 05/03/2024] [Indexed: 05/12/2024]
Abstract
To optimize electron energy for in situ imaging of large biological samples up to 10 μm in thickness with nanoscale resolutions, we implemented an analytical model based on elastic and inelastic characteristic angles. This model has been benchmarked by Monte Carlo simulations and can be used to predict the transverse beam size broadening as a function of electron energy while the probe beam traverses through the sample. As a result, the optimal choice of the electron beam energy can be realized. In addition, the impact of the dose-limited resolution was analysed. While the sample thickness is less than 10 μm, there exists an optimal electron beam energy below 10 MeV regarding a specific sample thickness. However, for samples thicker than 10 μm, the optimal beam energy is 10 MeV or higher depending on the sample thickness, and the ultimate resolution could become worse with the increase in the sample thickness. Moreover, a MeV-STEM column based on a two-stage lens system can be applied to reduce the beam size from one micron at aperture to one nanometre at the sample with the energy tuning range from 3 to 10 MeV. In conjunction with the state-of-the-art ultralow emittance electron source that we recently implemented, the maximum size of an electron beam when it traverses through an up to 10 μm thick bio-sample can be kept less than 10 nm. This is a critical step toward the in situ imaging of large, thick biological samples with nanometer resolution.
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Affiliation(s)
- Xi Yang
- National Synchrotron Light Source II, Brookhaven National Laboratory, Upton, NY 11973, USA; (V.S.); (T.S.)
| | - Liguo Wang
- Laboratory for BioMolecular Structure, Brookhaven National Laboratory, Upton, NY 11973, USA
| | - Victor Smaluk
- National Synchrotron Light Source II, Brookhaven National Laboratory, Upton, NY 11973, USA; (V.S.); (T.S.)
| | - Timur Shaftan
- National Synchrotron Light Source II, Brookhaven National Laboratory, Upton, NY 11973, USA; (V.S.); (T.S.)
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3
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Alonso Baez L, Bacete L. Cell wall dynamics: novel tools and research questions. JOURNAL OF EXPERIMENTAL BOTANY 2023; 74:6448-6467. [PMID: 37539735 PMCID: PMC10662238 DOI: 10.1093/jxb/erad310] [Citation(s) in RCA: 1] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 04/16/2023] [Accepted: 08/02/2023] [Indexed: 08/05/2023]
Abstract
Years ago, a classic textbook would define plant cell walls based on passive features. For instance, a sort of plant exoskeleton of invariable polysaccharide composition, and probably painted in green. However, currently, this view has been expanded to consider plant cell walls as active, heterogeneous, and dynamic structures with a high degree of complexity. However, what do we mean when we refer to a cell wall as a dynamic structure? How can we investigate the different implications of this dynamism? While the first question has been the subject of several recent publications, defining the ideal strategies and tools needed to address the second question has proven to be challenging due to the myriad of techniques available. In this review, we will describe the capacities of several methodologies to study cell wall composition, structure, and other aspects developed or optimized in recent years. Keeping in mind cell wall dynamism and plasticity, the advantages of performing long-term non-invasive live-imaging methods will be emphasized. We specifically focus on techniques developed for Arabidopsis thaliana primary cell walls, but the techniques could be applied to both secondary cell walls and other plant species. We believe this toolset will help researchers in expanding knowledge of these dynamic/evolving structures.
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Affiliation(s)
- Luis Alonso Baez
- Institute for Biology, Faculty of Natural Sciences, Norwegian University of Science and Technology, 5 Høgskoleringen, Trondheim, 7491, Norway
| | - Laura Bacete
- Institute for Biology, Faculty of Natural Sciences, Norwegian University of Science and Technology, 5 Høgskoleringen, Trondheim, 7491, Norway
- Umeå Plant Science Centre (UPSC), Department of Plant Physiology, Umeå University, 901 87 Umeå, Sweden
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4
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Sanchez Carrillo IB, Hoffmann PC, Barff T, Beck M, Germain H. Preparing Arabidopsis thaliana root protoplasts for cryo electron tomography. FRONTIERS IN PLANT SCIENCE 2023; 14:1261180. [PMID: 37810374 PMCID: PMC10556516 DOI: 10.3389/fpls.2023.1261180] [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: 07/19/2023] [Accepted: 09/04/2023] [Indexed: 10/10/2023]
Abstract
The use of protoplasts in plant biology has become a convenient tool for the application of transient gene expression. This model system has allowed the study of plant responses to biotic and abiotic stresses, protein location and trafficking, cell wall dynamics, and single-cell transcriptomics, among others. Although well-established protocols for isolating protoplasts from different plant tissues are available, they have never been used for studying plant cells using cryo electron microscopy (cryo-EM) and cryo electron tomography (cryo-ET). Here we describe a workflow to prepare root protoplasts from Arabidopsis thaliana plants for cryo-ET. The process includes protoplast isolation and vitrification on EM grids, and cryo-focused ion beam milling (cryo-FIB), with the aim of tilt series acquisition. The whole workflow, from growing the plants to the acquisition of the tilt series, may take a few months. Our protocol provides a novel application to use plant protoplasts as a tool for cryo-ET.
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Affiliation(s)
| | - Patrick C. Hoffmann
- Department of Molecular Sociology, Max-Planck-Institute for Biophysics, Frankfurt, Germany
| | - Teura Barff
- Department of Chemistry, Biochemistry, and Physics, Université du Québec à Trois-Rivières, Trois-Rivières, QC, Canada
| | - Martin Beck
- Department of Molecular Sociology, Max-Planck-Institute for Biophysics, Frankfurt, Germany
- Institute of Biochemistry, Goethe University Frankfurt, Frankfurt, Germany
| | - Hugo Germain
- Department of Chemistry, Biochemistry, and Physics, Université du Québec à Trois-Rivières, Trois-Rivières, QC, Canada
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5
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Wickramanayake JS, Czymmek KJ. A conventional fixation volume electron microscopy protocol for plants. Methods Cell Biol 2023; 177:83-99. [PMID: 37451777 DOI: 10.1016/bs.mcb.2023.04.008] [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] [Indexed: 07/18/2023]
Abstract
Volume electron microscopy techniques play an important role in plant research from understanding organelles and unicellular forms to developmental studies, environmental effects and microbial interactions with large plant structures, to name a few. Due to large air voids central vacuole, cell wall and waxy cuticle, many plant tissues pose challenges when trying to achieve high quality morphology, metal staining and adequate conductivity for high-resolution volume EM studies. Here, we applied a robust conventional chemical fixation strategy to address the special challenges of plant samples and suitable for, but not limited to, serial block-face and focused ion beam scanning electron microscopy. The chemistry of this protocol was modified from an approach developed for improved and uniform staining of large brain volumes. Briefly, primary fixation was in paraformaldehyde and glutaraldehyde with malachite green followed by secondary fixation with osmium tetroxide, potassium ferrocyanide, thiocarbohydrazide, osmium tetroxide and finally uranyl acetate and lead aspartate staining. Samples were then dehydrated in acetone with a propylene oxide transition and embedded in a hard formulation Quetol 651 resin. The samples were trimmed and mounted with silver epoxy, metal coated and imaged via serial block-face scanning electron microscopy and focal charge compensation for charge suppression. High-contrast plant tobacco and duckweed leaf cellular structures were readily visible including mitochondria, Golgi, endoplasmic reticulum and nuclear envelope membranes, as well as prominent chloroplast thylakoid membranes and individual lamella in grana stacks. This sample preparation protocol serves as a reliable starting point for routine plant volume electron microscopy.
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Affiliation(s)
- Janithri S Wickramanayake
- Donald Danforth Plant Science Center, Saint Louis, MO, United States; Advanced Bioimaging Laboratory, Donald Danforth Plant Science Center, Saint Louis, MO, United States
| | - Kirk J Czymmek
- Donald Danforth Plant Science Center, Saint Louis, MO, United States; Advanced Bioimaging Laboratory, Donald Danforth Plant Science Center, Saint Louis, MO, United States.
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6
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Electron tomography unravels new insights into fiber cell wall nanostructure; exploring 3D macromolecular biopolymeric nano-architecture of spruce fiber secondary walls. Sci Rep 2023; 13:2350. [PMID: 36759530 PMCID: PMC9911387 DOI: 10.1038/s41598-023-29113-x] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/01/2022] [Accepted: 01/31/2023] [Indexed: 02/11/2023] Open
Abstract
Lignocellulose biomass has a tremendous potential as renewable biomaterials for fostering the "bio-based society" and circular bioeconomy paradigm. It requires efficient use and breakdown of fiber cell walls containing mainly cellulose, hemicellulose and lignin biopolymers. Despite their great importance, there is an extensive debate on the true structure of fiber walls and knowledge on the macromolecular nano-organization is limited and remains elusive in 3D. We employed dual-axis electron tomography that allows visualization of previously unseen 3D macromolecular organization/biopolymeric nano-architecture of the secondary S2 layer of Norway spruce fiber wall. Unprecedented 3D nano-structural details with novel insights into cellulose microfibrils (~ 2 nm diameter), macrofibrils, nano-pore network and cell wall chemistry (volume %) across the S2 were explored and quantified including simulation of structure related permeability. Matrix polymer association with cellulose varied between microfibrils and macrofibrils with lignin directly associated with MFs. Simulated bio-nano-mechanical properties revealed stress distribution within the S2 and showed similar properties between the idealized 3D model and the native S2 (actual tomogram). Present work has great potential for significant advancements in lignocellulose research on nano-scale understanding of cell wall assembly/disassembly processes leading to more efficient industrial processes of functionalization, valorization and target modification technologies.
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7
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Zhang L, Liang X, Takáč T, Komis G, Li X, Zhang Y, Ovečka M, Chen Y, Šamaj J. Spatial proteomics of vesicular trafficking: coupling mass spectrometry and imaging approaches in membrane biology. PLANT BIOTECHNOLOGY JOURNAL 2023; 21:250-269. [PMID: 36204821 PMCID: PMC9884029 DOI: 10.1111/pbi.13929] [Citation(s) in RCA: 9] [Impact Index Per Article: 9.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Subscribe] [Scholar Register] [Received: 06/10/2022] [Revised: 08/14/2022] [Accepted: 09/08/2022] [Indexed: 06/16/2023]
Abstract
In plants, membrane compartmentalization requires vesicle trafficking for communication among distinct organelles. Membrane proteins involved in vesicle trafficking are highly dynamic and can respond rapidly to changes in the environment and to cellular signals. Capturing their localization and dynamics is thus essential for understanding the mechanisms underlying vesicular trafficking pathways. Quantitative mass spectrometry and imaging approaches allow a system-wide dissection of the vesicular proteome, the characterization of ligand-receptor pairs and the determination of secretory, endocytic, recycling and vacuolar trafficking pathways. In this review, we highlight major proteomics and imaging methods employed to determine the location, distribution and abundance of proteins within given trafficking routes. We focus in particular on methodologies for the elucidation of vesicle protein dynamics and interactions and their connections to downstream signalling outputs. Finally, we assess their biological applications in exploring different cellular and subcellular processes.
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Affiliation(s)
- Liang Zhang
- State Key Laboratory of Plant Physiology and Biochemistry, College of Biological SciencesChina Agricultural UniversityBeijingChina
- College of Life ScienceHenan Normal UniversityXinxiangChina
| | - Xinlin Liang
- State Key Laboratory of Plant Physiology and Biochemistry, College of Biological SciencesChina Agricultural UniversityBeijingChina
| | - Tomáš Takáč
- Department of Biotechnology, Faculty of SciencePalacky University OlomoucOlomoucCzech Republic
| | - George Komis
- Department of Cell Biology, Centre of the Region Hana for Biotechnological and Agricultural Research, Faculty of SciencePalacky University OlomoucOlomoucCzech Republic
| | - Xiaojuan Li
- College of Biological Sciences and TechnologyBeijing Forestry UniversityBeijingChina
| | - Yuan Zhang
- College of Biological Sciences and TechnologyBeijing Forestry UniversityBeijingChina
| | - Miroslav Ovečka
- Department of Biotechnology, Faculty of SciencePalacky University OlomoucOlomoucCzech Republic
| | - Yanmei Chen
- State Key Laboratory of Plant Physiology and Biochemistry, College of Biological SciencesChina Agricultural UniversityBeijingChina
| | - Jozef Šamaj
- Department of Biotechnology, Faculty of SciencePalacky University OlomoucOlomoucCzech Republic
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8
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Ito Y, Uemura T. Super resolution live imaging: The key for unveiling the true dynamics of membrane traffic around the Golgi apparatus in plant cells. FRONTIERS IN PLANT SCIENCE 2022; 13:1100757. [PMID: 36618665 PMCID: PMC9818705 DOI: 10.3389/fpls.2022.1100757] [Citation(s) in RCA: 2] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Figures] [Subscribe] [Scholar Register] [Received: 11/17/2022] [Accepted: 12/12/2022] [Indexed: 06/17/2023]
Abstract
In contrast to the relatively static image of the plants, the world inside each cell is surprisingly dynamic. Membrane-bounded organelles move actively on the cytoskeletons and exchange materials by vesicles, tubules, or direct contact between each other. In order to understand what is happening during those events, it is essential to visualize the working components in vivo. After the breakthrough made by the application of fluorescent proteins, the development of light microscopy enabled many discoveries in cell biology, including those about the membrane traffic in plant cells. Especially, super-resolution microscopy, which is becoming more and more accessible, is now one of the most powerful techniques. However, although the spatial resolution has improved a lot, there are still some difficulties in terms of the temporal resolution, which is also a crucial parameter for the visualization of the living nature of the intracellular structures. In this review, we will introduce the super resolution microscopy developed especially for live-cell imaging with high temporal resolution, and show some examples that were made by this tool in plant membrane research.
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Affiliation(s)
- Yoko Ito
- Institute for Human Life Science, Ochanomizu University, Tokyo, Japan
| | - Tomohiro Uemura
- Graduate School of Humanities and Sciences, Ochanomizu University, Tokyo, Japan
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9
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Hsiao AS, Huang JY. Bioimaging tools move plant physiology studies forward. FRONTIERS IN PLANT SCIENCE 2022; 13:976627. [PMID: 36204075 PMCID: PMC9530904 DOI: 10.3389/fpls.2022.976627] [Citation(s) in RCA: 2] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Key Words] [Track Full Text] [Figures] [Subscribe] [Scholar Register] [Received: 06/23/2022] [Accepted: 09/05/2022] [Indexed: 06/16/2023]
Affiliation(s)
- An-Shan Hsiao
- Biosciences, College of Life and Environmental Sciences, University of Exeter, Exeter, United Kingdom
| | - Ji-Ying Huang
- Cell Biology Core Lab, Institute of Plant and Microbial Biology, Academia Sinica, Taipei, Taiwan
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10
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Mair A, Bergmann DC. Advances in enzyme-mediated proximity labeling and its potential for plant research. PLANT PHYSIOLOGY 2022; 188:756-768. [PMID: 34662401 PMCID: PMC8825456 DOI: 10.1093/plphys/kiab479] [Citation(s) in RCA: 14] [Impact Index Per Article: 7.0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 08/03/2021] [Accepted: 09/21/2021] [Indexed: 06/12/2023]
Abstract
Cellular processes rely on the intimate interplay of different molecules, including DNA, RNA, proteins, and metabolites. Obtaining and integrating data on their abundance and dynamics at high temporal and spatial resolution are essential for our understanding of plant growth and development. In the past decade, enzymatic proximity labeling (PL) has emerged as a powerful tool to study local protein and nucleotide ensembles, discover protein-protein and protein-nucleotide interactions, and resolve questions about protein localization and membrane topology. An ever-growing number and continuous improvement of enzymes and methods keep broadening the spectrum of possible applications for PL and make it more accessible to different organisms, including plants. While initial PL experiments in plants required high expression levels and long labeling times, recently developed faster enzymes now enable PL of proteins on a cell type-specific level, even with low-abundant baits, and in different plant species. Moreover, expanding the use of PL for additional purposes, such as identification of locus-specific gene regulators or high-resolution electron microscopy may now be in reach. In this review, we give an overview of currently available PL enzymes and their applications in mammalian cell culture and plants. We discuss the challenges and limitations of PL methods and highlight open questions and possible future directions for PL in plants.
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Affiliation(s)
- Andrea Mair
- Howard Hughes Medical Institute and Department of Biology, Stanford University, Stanford, California 94305, USA
| | - Dominique C Bergmann
- Howard Hughes Medical Institute and Department of Biology, Stanford University, Stanford, California 94305, USA
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11
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Three-dimensional reconstruction by electron tomography for the application to ultrastructural analysis of SARS-CoV-2 particles. Med Mol Morphol 2021; 55:60-67. [PMID: 34825978 PMCID: PMC8617558 DOI: 10.1007/s00795-021-00309-2] [Citation(s) in RCA: 7] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/17/2021] [Accepted: 11/11/2021] [Indexed: 12/02/2022]
Abstract
SARS-CoV-2 is the cause of COVID-19. The three-dimensional morphology of viral particles existing and multiplying in infected cells has not been established by electron tomography, which is different from cryo-electron tomography using frozen samples. In this study, we establish the morphological structure of SARS-CoV-2 particles by three-dimensional reconstruction of images obtained by electron tomography and transmission electron microscopy of biological samples embedded in epoxy resin. The characteristic roots of spike structures were found to be arranged at the surface of a virion covered with an envelope. A high-electron-density structure that appears to be a nucleocapsid was observed inside the envelope of the virion on three-dimensional images reconstructed by electron tomography. The SARS-CoV-2 particles that budded in the vacuoles in the cytoplasm were morphologically identical to those found outside the cells, suggesting that mature and infectious SARS-CoV-2 particles were already produced in the vacuoles. Here, we show the three-dimensional morphological structure of SARS-CoV-2 particles reconstructed by electron tomography. To control infection, inhibition of viral release from vacuoles would be a new target in the development of prophylactic agents against SARS-CoV-2.
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12
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Mazur R, Mostowska A, Kowalewska Ł. How to Measure Grana - Ultrastructural Features of Thylakoid Membranes of Plant Chloroplasts. FRONTIERS IN PLANT SCIENCE 2021; 12:756009. [PMID: 34691132 PMCID: PMC8527009 DOI: 10.3389/fpls.2021.756009] [Citation(s) in RCA: 6] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 08/09/2021] [Accepted: 09/09/2021] [Indexed: 06/11/2023]
Abstract
Granum is a basic structural unit of the thylakoid membrane network of plant chloroplasts. It is composed of multiple flattened membranes forming a stacked arrangement of a cylindrical shape. Grana membranes are composed of lipids and tightly packed pigment-protein complexes whose primary role is the catalysis of photosynthetic light reactions. These membranes are highly dynamic structures capable of adapting to changing environmental conditions by fine-tuning photochemical efficiency, manifested by the structural reorganization of grana stacks. Due to a nanometer length scale of the structural granum features, the application of high-resolution electron microscopic techniques is essential for a detailed analysis of the granum architecture. This mini-review overviews recent approaches to quantitative grana structure analyses from electron microscopy data, highlighting the basic manual measurements and semi-automated workflows. We outline and define structural parameters used by different authors, for instance, granum height and diameter, thylakoid thickness, end-membrane length, Stacking Repeat Distance, and Granum Lateral Irregularity. This article also presents insights into efficient and effective measurements of grana stacks visualized on 2D micrographs. The information on how to correctly interpret obtained data, taking into account the 3D nature of grana stacks projected onto 2D space of electron micrograph, is also given. Grana ultrastructural observations reveal key features of this intriguing membrane arrangement, broadening our knowledge of the thylakoid network's remarkable plasticity.
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Affiliation(s)
- Radosław Mazur
- Department of Metabolic Regulation, Institute of Biochemistry, Faculty of Biology, University of Warsaw, Warsaw, Poland
| | - Agnieszka Mostowska
- Department of Plant Anatomy and Cytology, Institute of Plant Experimental Biology and Biotechnology, Faculty of Biology, University of Warsaw, Warsaw, Poland
| | - Łucja Kowalewska
- Department of Plant Anatomy and Cytology, Institute of Plant Experimental Biology and Biotechnology, Faculty of Biology, University of Warsaw, Warsaw, Poland
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13
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Liu Z, Gao J, Cui Y, Klumpe S, Xiang Y, Erdmann PS, Jiang L. Membrane imaging in the plant endomembrane system. PLANT PHYSIOLOGY 2021; 185:562-576. [PMID: 33793889 PMCID: PMC8133680 DOI: 10.1093/plphys/kiaa040] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 06/01/2020] [Accepted: 08/11/2020] [Indexed: 05/10/2023]
Abstract
Recent studies on membrane imaging in the plant endomembrane system by 2-D/3-D CLSM and TEM provide future perspectives of whole-cell ET and cryo-FIB-aided cryo-ET analysis.
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Affiliation(s)
- Zhiqi Liu
- State Key Laboratory of Agrobiotechnology, School of Life Sciences, Centre for Cell and Developmental Biology, The Chinese University of Hong Kong, Shatin, Hong Kong
| | - Jiayang Gao
- State Key Laboratory of Agrobiotechnology, School of Life Sciences, Centre for Cell and Developmental Biology, The Chinese University of Hong Kong, Shatin, Hong Kong
| | - Yong Cui
- State Key Laboratory of Agrobiotechnology, School of Life Sciences, Centre for Cell and Developmental Biology, The Chinese University of Hong Kong, Shatin, Hong Kong
| | - Sven Klumpe
- Max Planck Institute of Biochemistry, Am Klopferspitz 18, 82152 Martinsried, Germany
| | - Yun Xiang
- MOE Key Laboratory of Cell Activities and Stress Adaptations, School of Life Sciences, Lanzhou University, Lanzhou 730000, China
| | - Philipp S Erdmann
- Max Planck Institute of Biochemistry, Am Klopferspitz 18, 82152 Martinsried, Germany
| | - Liwen Jiang
- State Key Laboratory of Agrobiotechnology, School of Life Sciences, Centre for Cell and Developmental Biology, The Chinese University of Hong Kong, Shatin, Hong Kong
- CUHK Shenzhen Research Institute, The Chinese University of Hong Kong, Shenzhen 518057, China
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14
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Kozieł E, Otulak-Kozieł K, Bujarski JJ. Plant Cell Wall as a Key Player During Resistant and Susceptible Plant-Virus Interactions. Front Microbiol 2021; 12:656809. [PMID: 33776985 PMCID: PMC7994255 DOI: 10.3389/fmicb.2021.656809] [Citation(s) in RCA: 22] [Impact Index Per Article: 7.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/21/2021] [Accepted: 02/19/2021] [Indexed: 01/06/2023] Open
Abstract
The cell wall is a complex and integral part of the plant cell. As a structural element it sustains the shape of the cell and mediates contact among internal and external factors. We have been aware of its involvement in both abiotic (like drought or frost) and biotic stresses (like bacteria or fungi) for some time. In contrast to bacterial and fungal pathogens, viruses are not mechanical destructors of host cell walls, but relatively little is known about remodeling of the plant cell wall in response to viral biotic stress. New research results indicate that the cell wall represents a crucial active component during the plant’s response to different viral infections. Apparently, cell wall genes and proteins play key roles during interaction, having a direct influence on the rebuilding of the cell wall architecture. The plant cell wall is involved in both susceptibility as well as resistance reactions. In this review we summarize important progress made in research on plant virus impact on cell wall remodeling. Analyses of essential defensive wall associated proteins in susceptible and resistant responses demonstrate that the components of cell wall metabolism can affect the spread of the virus as well as activate the apoplast- and symplast-based defense mechanisms, thus contributing to the complex network of the plant immune system. Although the cell wall reorganization during the plant-virus interaction remains a challenging task, the use of novel tools and methods to investigate its composition and structure will greatly contribute to our knowledge in the field.
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Affiliation(s)
- Edmund Kozieł
- Institute of Biology, Department of Botany, Warsaw University of Life Sciences - SGGW, Warsaw, Poland
| | - Katarzyna Otulak-Kozieł
- Institute of Biology, Department of Botany, Warsaw University of Life Sciences - SGGW, Warsaw, Poland
| | - Józef Julian Bujarski
- Department of Biological Sciences, Northern Illinois University, DeKalb, IL, United States
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15
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Zhang X, Man Y, Zhuang X, Shen J, Zhang Y, Cui Y, Yu M, Xing J, Wang G, Lian N, Hu Z, Ma L, Shen W, Yang S, Xu H, Bian J, Jing Y, Li X, Li R, Mao T, Jiao Y, Sodmergen, Ren H, Lin J. Plant multiscale networks: charting plant connectivity by multi-level analysis and imaging techniques. SCIENCE CHINA-LIFE SCIENCES 2021; 64:1392-1422. [PMID: 33974222 DOI: 10.1007/s11427-020-1910-1] [Citation(s) in RCA: 18] [Impact Index Per Article: 6.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Subscribe] [Scholar Register] [Received: 12/15/2020] [Accepted: 03/04/2021] [Indexed: 12/21/2022]
Abstract
In multicellular and even single-celled organisms, individual components are interconnected at multiscale levels to produce enormously complex biological networks that help these systems maintain homeostasis for development and environmental adaptation. Systems biology studies initially adopted network analysis to explore how relationships between individual components give rise to complex biological processes. Network analysis has been applied to dissect the complex connectivity of mammalian brains across different scales in time and space in The Human Brain Project. In plant science, network analysis has similarly been applied to study the connectivity of plant components at the molecular, subcellular, cellular, organic, and organism levels. Analysis of these multiscale networks contributes to our understanding of how genotype determines phenotype. In this review, we summarized the theoretical framework of plant multiscale networks and introduced studies investigating plant networks by various experimental and computational modalities. We next discussed the currently available analytic methodologies and multi-level imaging techniques used to map multiscale networks in plants. Finally, we highlighted some of the technical challenges and key questions remaining to be addressed in this emerging field.
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Affiliation(s)
- Xi Zhang
- Beijing Advanced Innovation Center for Tree Breeding by Molecular Design, Beijing Forestry University, Beijing, 100083, China.,College of Biological Sciences and Biotechnology, Beijing Forestry University, Beijing, 100083, China
| | - Yi Man
- Beijing Advanced Innovation Center for Tree Breeding by Molecular Design, Beijing Forestry University, Beijing, 100083, China.,College of Biological Sciences and Biotechnology, Beijing Forestry University, Beijing, 100083, China
| | - Xiaohong Zhuang
- School of Life Sciences, Centre for Cell & Developmental Biology and State Key Laboratory of Agrobiotechnology, The Chinese University of Hong Kong, Hong Kong, 999077, China
| | - Jinbo Shen
- State Key Laboratory of Subtropical Silviculture, Zhejiang A&F University, Hangzhou, 311300, China
| | - Yi Zhang
- Key Laboratory of Cell Proliferation and Regulation Biology, Ministry of Education, College of Life Science, Beijing Normal University, Beijing, 100875, China
| | - Yaning Cui
- Beijing Advanced Innovation Center for Tree Breeding by Molecular Design, Beijing Forestry University, Beijing, 100083, China.,College of Biological Sciences and Biotechnology, Beijing Forestry University, Beijing, 100083, China
| | - Meng Yu
- Beijing Advanced Innovation Center for Tree Breeding by Molecular Design, Beijing Forestry University, Beijing, 100083, China.,College of Biological Sciences and Biotechnology, Beijing Forestry University, Beijing, 100083, China
| | - Jingjing Xing
- Key Laboratory of Plant Stress Biology, School of Life Sciences, Henan University, Kaifeng, 457004, China
| | - Guangchao Wang
- College of Biological Sciences and Biotechnology, Beijing Forestry University, Beijing, 100083, China
| | - Na Lian
- Beijing Advanced Innovation Center for Tree Breeding by Molecular Design, Beijing Forestry University, Beijing, 100083, China.,College of Biological Sciences and Biotechnology, Beijing Forestry University, Beijing, 100083, China
| | - Zijian Hu
- College of Biological Sciences and Biotechnology, Beijing Forestry University, Beijing, 100083, China
| | - Lingyu Ma
- College of Biological Sciences and Biotechnology, Beijing Forestry University, Beijing, 100083, China
| | - Weiwei Shen
- College of Biological Sciences and Biotechnology, Beijing Forestry University, Beijing, 100083, China
| | - Shunyao Yang
- College of Biological Sciences and Biotechnology, Beijing Forestry University, Beijing, 100083, China
| | - Huimin Xu
- College of Biological Sciences, China Agricultural University, Beijing, 100193, China
| | - Jiahui Bian
- College of Biological Sciences and Biotechnology, Beijing Forestry University, Beijing, 100083, China
| | - Yanping Jing
- College of Biological Sciences and Biotechnology, Beijing Forestry University, Beijing, 100083, China
| | - Xiaojuan Li
- Beijing Advanced Innovation Center for Tree Breeding by Molecular Design, Beijing Forestry University, Beijing, 100083, China.,College of Biological Sciences and Biotechnology, Beijing Forestry University, Beijing, 100083, China
| | - Ruili Li
- Beijing Advanced Innovation Center for Tree Breeding by Molecular Design, Beijing Forestry University, Beijing, 100083, China.,College of Biological Sciences and Biotechnology, Beijing Forestry University, Beijing, 100083, China
| | - Tonglin Mao
- State Key Laboratory of Plant Physiology and Biochemistry, Department of Plant Sciences, College of Biological Sciences, China Agricultural University, Beijing, 100193, China
| | - Yuling Jiao
- State Key Laboratory of Plant Genomics, Institute of Genetics and Developmental Biology, Chinese Academy of Sciences, and National Center for Plant Gene Research, Beijing, 100101, China
| | - Sodmergen
- Key Laboratory of Ministry of Education for Cell Proliferation and Differentiation, College of Life Sciences, Peking University, Beijing, 100871, China
| | - Haiyun Ren
- Key Laboratory of Cell Proliferation and Regulation Biology, Ministry of Education, College of Life Science, Beijing Normal University, Beijing, 100875, China
| | - Jinxing Lin
- Beijing Advanced Innovation Center for Tree Breeding by Molecular Design, Beijing Forestry University, Beijing, 100083, China. .,College of Biological Sciences and Biotechnology, Beijing Forestry University, Beijing, 100083, China.
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16
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Beyond X-rays: an overview of emerging structural biology methods. Emerg Top Life Sci 2021; 5:221-230. [DOI: 10.1042/etls20200272] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/29/2020] [Revised: 12/27/2020] [Accepted: 01/20/2021] [Indexed: 11/17/2022]
Abstract
Structural biologists rely on X-ray crystallography as the main technique for determining the three-dimensional structures of macromolecules; however, in recent years, new methods that go beyond X-ray-based technologies are broadening the selection of tools to understand molecular structure and function. Simultaneously, national facilities are developing programming tools and maintaining personnel to aid novice structural biologists in de novo structure determination. The combination of X-ray free electron lasers (XFELs) and serial femtosecond crystallography (SFX) now enable time-resolved structure determination that allows for capture of dynamic processes, such as reaction mechanism and conformational flexibility. XFEL and SFX, along with microcrystal electron diffraction (MicroED), help side-step the need for large crystals for structural studies. Moreover, advances in cryogenic electron microscopy (cryo-EM) as a tool for structure determination is revolutionizing how difficult to crystallize macromolecules and/or complexes can be visualized at the atomic scale. This review aims to provide a broad overview of these new methods and to guide readers to more in-depth literature of these methods.
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17
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Czymmek K, Sawant A, Goodman K, Pennington J, Pedersen P, Hoon M, Otegui MS. Imaging Plant Cells by High-Pressure Freezing and Serial block-face scanning electron microscopy. Methods Mol Biol 2020; 2177:69-81. [PMID: 32632806 DOI: 10.1007/978-1-0716-0767-1_7] [Citation(s) in RCA: 7] [Impact Index Per Article: 1.8] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 06/11/2023]
Abstract
This chapter describes methods to enhanced contrast of plant material processed by high-pressure freezing and freeze substitution for improved visualization by serial block-face scanning electron microscopy (SBEM). The contrast enhancing steps are based on a protocol involving the sequential incubation of samples in heavy metals and sodium thiocarbohydrazide (OTO staining). We also describe the pipeline for imaging plant tissues in a commercial SBEM system (Gatan 3View®) and routines for the image analysis and three-dimensional reconstructions using open-source and commercial software packages.
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Affiliation(s)
- Kirk Czymmek
- Advanced Bioimaging Laboratory, Donald Danforth Plant Science Center, St. Louis, MO, USA
| | - Abhilash Sawant
- Department of Ophthalmology and Visual Sciences, University of Wisconsin-Madison, Madison, WI, USA
- Department of Neuroscience, University of Wisconsin-Madison, Madison, WI, USA
| | - Kaija Goodman
- Department of Botany, University of Wisconsin-Madison, Madison, WI, USA
- Laboratory of Cell and Molecular Biology, University of Wisconsin-Madison, Madison, WI, USA
| | - Janice Pennington
- Laboratory of Cell and Molecular Biology, University of Wisconsin-Madison, Madison, WI, USA
| | - Pal Pedersen
- Carl Zeiss Microscopy, LLC, White Plains, NY, USA
| | - Mrinalini Hoon
- Department of Ophthalmology and Visual Sciences, University of Wisconsin-Madison, Madison, WI, USA
- McPherson Eye Research Institute, University of Wisconsin-Madison, Madison, WI, USA
| | - Marisa S Otegui
- Department of Botany, University of Wisconsin-Madison, Madison, WI, USA.
- Laboratory of Cell and Molecular Biology, University of Wisconsin-Madison, Madison, WI, USA.
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18
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Zhu D, Zhang M, Gao C, Shen J. Protein trafficking in plant cells: Tools and markers. SCIENCE CHINA-LIFE SCIENCES 2019; 63:343-363. [DOI: 10.1007/s11427-019-9598-3] [Citation(s) in RCA: 12] [Impact Index Per Article: 2.4] [Reference Citation Analysis] [Track Full Text] [Subscribe] [Scholar Register] [Received: 05/26/2019] [Accepted: 07/22/2019] [Indexed: 12/26/2022]
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19
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Wang P, Liang Z, Kang BH. Electron tomography of plant organelles and the outlook for correlative microscopic approaches. THE NEW PHYTOLOGIST 2019; 223:1756-1761. [PMID: 31058325 DOI: 10.1111/nph.15882] [Citation(s) in RCA: 11] [Impact Index Per Article: 2.2] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Received: 02/24/2019] [Accepted: 04/11/2019] [Indexed: 05/17/2023]
Abstract
Structural analyses of organelles and localization of proteins in their confines are essential to investigate inner workings of eukaryotic cells. Electron tomography (ET) is a three-dimensional electron microscopy method with which we can extract sliced views of organelles from any direction and quantify their structural parameters at nanometer-level resolution. This advanced electron microscopy tool is suited for characterization of convoluted membrane compartments and of cellular constituents of dimensions smaller than 100 nm. ET studies of plant cells fixed by rapid freezing have expanded our understanding of the biogenesis and functions of plant organelles. Here we describe how the molecular imaging capacity of correlative light and electron microscopy can be integrated with ET in studies of plant organelles.
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Affiliation(s)
- Pengfei Wang
- Centre for Cell and Developmental Biology, State Key Laboratory for Agrobiotechnology, School of Life Sciences, The Chinese University of Hong Kong, Shatin, New Territories, Hong Kong, China
| | - Zizhen Liang
- Centre for Cell and Developmental Biology, State Key Laboratory for Agrobiotechnology, School of Life Sciences, The Chinese University of Hong Kong, Shatin, New Territories, Hong Kong, China
| | - Byung-Ho Kang
- Centre for Cell and Developmental Biology, State Key Laboratory for Agrobiotechnology, School of Life Sciences, The Chinese University of Hong Kong, Shatin, New Territories, Hong Kong, China
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