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Garty Y, Bussi Y, Levin-Zaidman S, Shimoni E, Kirchhoff H, Charuvi D, Nevo R, Reich Z. Thylakoid membrane stacking controls electron transport mode during the dark-to-light transition by adjusting the distances between PSI and PSII. Nat Plants 2024; 10:512-524. [PMID: 38396112 DOI: 10.1038/s41477-024-01628-9] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [What about the content of this article? (0)] [Affiliation(s)] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 04/27/2023] [Accepted: 01/23/2024] [Indexed: 02/25/2024]
Abstract
The balance between linear electron transport (LET) and cyclic electron transport (CET) plays an essential role in plant adaptation and protection against photo-induced damage. This balance is largely maintained by phosphorylation-driven alterations in the PSII-LHCII assembly and thylakoid membrane stacking. During the dark-to-light transition, plants shift this balance from CET, which prevails to prevent overreduction of the electron transport chain and consequent photo-induced damage, towards LET, which enables efficient CO2 assimilation and biomass production. Using freeze-fracture cryo-scanning electron microscopy and transmission electron microscopy of Arabidopsis leaves, we reveal unique membrane regions possessing characteristics of both stacked and unstacked regions of the thylakoid network that form during this transition. A notable consequence of the morphological attributes of these regions, which we refer to as 'stacked thylakoid doublets', is an overall increase in the proximity and connectivity of the two photosystems (PSI and PSII) that drive LET. This, in turn, reduces diffusion distances and barriers for the mobile carriers that transfer electrons between the two PSs, thereby maximizing LET and optimizing the plant's ability to utilize light energy. The mechanics described here for the shift between CET and LET during the dark-to-light transition are probably also used during chromatic adaptation mediated by state transitions.
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Affiliation(s)
- Yuval Garty
- Department of Biomolecular Sciences, Weizmann Institute of Science, Rehovot, Israel
| | - Yuval Bussi
- Department of Biomolecular Sciences, Weizmann Institute of Science, Rehovot, Israel
| | - Smadar Levin-Zaidman
- Department of Chemical Research Support, Weizmann Institute of Science, Rehovot, Israel
| | - Eyal Shimoni
- Department of Chemical Research Support, Weizmann Institute of Science, Rehovot, Israel
| | - Helmut Kirchhoff
- Institute of Biological Chemistry, Washington State University, Pullman, WA, USA
| | - Dana Charuvi
- Institute of Plant Sciences, Agricultural Research Organization, Volcani Institute, Rishon LeZion, Israel
| | - Reinat Nevo
- Department of Biomolecular Sciences, Weizmann Institute of Science, Rehovot, Israel.
| | - Ziv Reich
- Department of Biomolecular Sciences, Weizmann Institute of Science, Rehovot, Israel.
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Wightman R. An Overview of Cryo-Scanning Electron Microscopy Techniques for Plant Imaging. Plants (Basel) 2022; 11:plants11091113. [PMID: 35567113 PMCID: PMC9106016 DOI: 10.3390/plants11091113] [Citation(s) in RCA: 6] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [What about the content of this article? (0)] [Affiliation(s)] [Abstract] [Key Words] [Track Full Text] [Subscribe] [Scholar Register] [Received: 03/25/2022] [Revised: 04/14/2022] [Accepted: 04/14/2022] [Indexed: 05/02/2023]
Abstract
Many research questions require the study of plant morphology, in particular cells and tissues, as close to their native context as possible and without physical deformations from some preparatory chemical reagents or sample drying. Cryo-scanning electron microscopy (cryoSEM) involves rapid freezing and maintenance of the sample at an ultra-low temperature for detailed surface imaging by a scanning electron beam. The data are useful for exploring tissue/cell morphogenesis, plus an additional cryofracture/cryoplaning/milling step gives information on air and water spaces as well as subcellular ultrastructure. This review gives an overview from sample preparation through to imaging and a detailed account of how this has been applied across diverse areas of plant research. Future directions and improvements to the technique are discussed.
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Affiliation(s)
- Raymond Wightman
- The Sainsbury Laboratory, University of Cambridge, Bateman Street, Cambridge CB2 1LR, UK
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Li M, Mukhopadhyay R, Svoboda V, Oung HMO, Mullendore DL, Kirchhoff H. Measuring the dynamic response of the thylakoid architecture in plant leaves by electron microscopy. Plant Direct 2020; 4:e00280. [PMID: 33195966 PMCID: PMC7644818 DOI: 10.1002/pld3.280] [Citation(s) in RCA: 14] [Impact Index Per Article: 3.5] [Reference Citation Analysis] [What about the content of this article? (0)] [Affiliation(s)] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 08/05/2020] [Revised: 09/15/2020] [Accepted: 09/29/2020] [Indexed: 05/19/2023]
Abstract
UNLABELLED The performance of the photosynthesis machinery in plants, including light harvesting, electron transport, and protein repair, is controlled by structural changes in the thylakoid membrane system inside the chloroplasts. In particular, the structure of the stacked grana area of thylakoid membranes is highly dynamic, changing in response to different environmental cues such as light intensity. For example, the aqueous thylakoid lumen enclosed by thylakoid membranes in grana has been documented to swell in the presence of light. However, light-induced alteration of the stromal gap in the stacked grana (partition gap) and of the unstacked stroma lamellae has not been well characterized. Light-induced changes in the entire thylakoid membrane system, including the lumen in both stacked and unstacked domains as well as the partition gap, are presented here, and the functional implications are discussed. This structural analysis was made possible by development of a robust semi-automated image analysis method combined with optimized plant tissue fixation techniques for transmission electron microscopy generating quantitative structural results for the analysis of thylakoid ultrastructure. SIGNIFICANCE STATEMENT A methodical pipeline ranging from optimized leaf tissue preparation for electron microscopy to quantitative image analysis was established. This methodical development was employed to study details of light-induced changes in the plant thylakoid ultrastructure. It was found that the lumen of the entire thylakoid system (stacked and unstacked domains) undergoes light-induced swelling, whereas adjacent membranes on the stroma side in stacked grana thylakoid approach each other.
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Affiliation(s)
- Meng Li
- Institute of Biological ChemistryWashington State UniversityPullmanWAUSA
- Present address:
School of OceanographyUniversity of WashingtonSeattleWAUSA
| | - Roma Mukhopadhyay
- Institute of Biological ChemistryWashington State UniversityPullmanWAUSA
| | - Václav Svoboda
- Institute of Biological ChemistryWashington State UniversityPullmanWAUSA
| | | | | | - Helmut Kirchhoff
- Institute of Biological ChemistryWashington State UniversityPullmanWAUSA
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Wietrzynski W, Schaffer M, Tegunov D, Albert S, Kanazawa A, Plitzko JM, Baumeister W, Engel BD. Charting the native architecture of Chlamydomonas thylakoid membranes with single-molecule precision. eLife 2020; 9:53740. [PMID: 32297859 PMCID: PMC7164959 DOI: 10.7554/elife.53740] [Citation(s) in RCA: 52] [Impact Index Per Article: 13.0] [Reference Citation Analysis] [What about the content of this article? (0)] [Affiliation(s)] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/22/2019] [Accepted: 04/03/2020] [Indexed: 12/18/2022] Open
Abstract
Thylakoid membranes scaffold an assortment of large protein complexes that work together to harness the energy of light. It has been a longstanding challenge to visualize how the intricate thylakoid network organizes these protein complexes to finely tune the photosynthetic reactions. Previously, we used in situ cryo-electron tomography to reveal the native architecture of thylakoid membranes (Engel et al., 2015). Here, we leverage technical advances to resolve the individual protein complexes within these membranes. Combined with a new method to visualize membrane surface topology, we map the molecular landscapes of thylakoid membranes inside green algae cells. Our tomograms provide insights into the molecular forces that drive thylakoid stacking and reveal that photosystems I and II are strictly segregated at the borders between appressed and non-appressed membrane domains. This new approach to charting thylakoid topology lays the foundation for dissecting photosynthetic regulation at the level of single protein complexes within the cell.
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Affiliation(s)
- Wojciech Wietrzynski
- Helmholtz Pioneer Campus, Helmholtz Zentrum München, Neuherberg, Germany.,Department of Molecular Structural Biology, Max Planck Institute of Biochemistry, Martinsried, Germany
| | - Miroslava Schaffer
- Department of Molecular Structural Biology, Max Planck Institute of Biochemistry, Martinsried, Germany
| | - Dimitry Tegunov
- Department of Molecular Biology, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany
| | - Sahradha Albert
- Department of Molecular Structural Biology, Max Planck Institute of Biochemistry, Martinsried, Germany
| | - Atsuko Kanazawa
- MSU-DOE Plant Research Lab, Michigan State University, East Lansing, United States
| | - Jürgen M Plitzko
- Department of Molecular Structural Biology, Max Planck Institute of Biochemistry, Martinsried, Germany
| | - Wolfgang Baumeister
- Department of Molecular Structural Biology, Max Planck Institute of Biochemistry, Martinsried, Germany
| | - Benjamin D Engel
- Helmholtz Pioneer Campus, Helmholtz Zentrum München, Neuherberg, Germany.,Department of Molecular Structural Biology, Max Planck Institute of Biochemistry, Martinsried, Germany
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Noble JM, Roberts LM, Vidavsky N, Chiou AE, Fischbach C, Paszek MJ, Estroff LA, Kourkoutis LF. Direct comparison of optical and electron microscopy methods for structural characterization of extracellular vesicles. J Struct Biol 2020; 210:107474. [PMID: 32032755 PMCID: PMC7067680 DOI: 10.1016/j.jsb.2020.107474] [Citation(s) in RCA: 27] [Impact Index Per Article: 6.8] [Reference Citation Analysis] [What about the content of this article? (0)] [Affiliation(s)] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/06/2019] [Revised: 12/06/2019] [Accepted: 01/29/2020] [Indexed: 12/28/2022]
Abstract
As interest in the role of extracellular vesicles in cell-to-cell communication has increased, so has the use of microscopy and analytical techniques to assess their formation, release, and morphology. In this study, we evaluate scanning electron microscopy (SEM) and cryo-SEM for characterizing the formation and shedding of vesicles from human breast cell lines, parental and hyaluronan synthase 3-(HAS3)-overexpressing MCF10A cells, grown directly on transmission electron microscopy (TEM) grids. While cells imaged with conventional and cryo-SEM exhibit distinct morphologies due to the sample preparation process for each technique, tubular structures protruding from the cell surfaces were observed with both approaches. For HAS3-MCF10A cells, vesicles were present along the length of membrane protrusions. Once completely shed from the cells, extracellular vesicles were characterized using nanoparticle tracking analysis (NTA) and cryo-TEM. The size distributions obtained by each technique were different not only in the range of vesicles analyzed, but also in the relative proportion of smaller-to-larger vesicles. These differences are attributed to the presence of biological debris in the media, which is difficult to differentiate from vesicles in NTA. Furthermore, we demonstrate that cryo-TEM can be used to distinguish between vesicles based on their respective surface structures, thereby providing a path to differentiating vesicle subpopulations and identifying their size distributions. Our study emphasizes the necessity of pairing several techniques to characterize extracellular vesicles.
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Affiliation(s)
- Jade M Noble
- Robert Frederick Smith School of Chemical and Biomolecular Engineering, Cornell University, Ithaca, NY, USA
| | - LaDeidra Monét Roberts
- Nancy E. and Peter C. Meinig School of Biomedical Engineering, Cornell University, Ithaca, NY, USA
| | - Netta Vidavsky
- Department of Materials Science and Engineering, Cornell University, Ithaca, NY, USA
| | - Aaron E Chiou
- Nancy E. and Peter C. Meinig School of Biomedical Engineering, Cornell University, Ithaca, NY, USA
| | - Claudia Fischbach
- Nancy E. and Peter C. Meinig School of Biomedical Engineering, Cornell University, Ithaca, NY, USA; Kavli Institute at Cornell for Nanoscale Science, Cornell University, Ithaca, NY, USA
| | - Matthew J Paszek
- Robert Frederick Smith School of Chemical and Biomolecular Engineering, Cornell University, Ithaca, NY, USA; Kavli Institute at Cornell for Nanoscale Science, Cornell University, Ithaca, NY, USA.
| | - Lara A Estroff
- Department of Materials Science and Engineering, Cornell University, Ithaca, NY, USA; Kavli Institute at Cornell for Nanoscale Science, Cornell University, Ithaca, NY, USA.
| | - Lena F Kourkoutis
- School of Applied and Engineering Physics, Cornell University, Ithaca, NY, USA; Kavli Institute at Cornell for Nanoscale Science, Cornell University, Ithaca, NY, USA.
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