1
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Jansen RLM, de Boer R, de Lange EMF, Koster J, Vlijm R, Waterham HR, van der Klei IJ. Overexpression of PEX14 results in mistargeting to mitochondria, accompanied by organelle fragmentation and clustering in human embryonic kidney cells. BIOCHIMICA ET BIOPHYSICA ACTA. MOLECULAR CELL RESEARCH 2024; 1871:119754. [PMID: 38762172 DOI: 10.1016/j.bbamcr.2024.119754] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Received: 10/18/2023] [Revised: 04/15/2024] [Accepted: 05/07/2024] [Indexed: 05/20/2024]
Abstract
Peroxisome biogenesis disorders are caused by pathogenic variants in genes involved in biogenesis and maintenance of peroxisomes. However, mitochondria are also often affected in these diseases. Peroxisomal membrane proteins, including PEX14, have been found to mislocalise to mitochondria in cells lacking peroxisomes. Recent studies indicated that this mislocalisation contributes to mitochondrial abnormalities in PEX3-deficient patient fibroblasts cells. Here, we studied whether mitochondrial morphology is also affected in PEX3-deficient HEK293 cells and whether PEX14 mislocalises to mitochondria in these cells. Using high-resolution imaging techniques, we show that although endogenous PEX14 mislocalises to mitochondria, mitochondrial morphology was normal in PEX3-KO HEK293 cells. However, we discovered that overexpression of tagged PEX14 in wild-type HEK293 cells resulted in its mitochondrial localisation, accompanied by altered mitochondrial morphology. Our data indicate that overexpression of tagged PEX14 alone directly or indirectly cause mitochondrial abnormalities in cells containing peroxisomes.
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Affiliation(s)
- Renate L M Jansen
- Molecular Cell Biology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, the Netherlands
| | - Rinse de Boer
- Molecular Cell Biology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, the Netherlands
| | - Eline M F de Lange
- Molecular Cell Biology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, the Netherlands; Molecular Biophysics, Zernike Institute for Advanced Materials, University of Groningen, Groningen, the Netherlands
| | - Janet Koster
- Laboratory of Genetic Metabolic Diseases & Amsterdam Gastroenterology, Endocrinology & Metabolism (AGEM), Amsterdam UMC - location AMC, Amsterdam, the Netherlands
| | - Rifka Vlijm
- Molecular Biophysics, Zernike Institute for Advanced Materials, University of Groningen, Groningen, the Netherlands
| | - Hans R Waterham
- Laboratory of Genetic Metabolic Diseases & Amsterdam Gastroenterology, Endocrinology & Metabolism (AGEM), Amsterdam UMC - location AMC, Amsterdam, the Netherlands
| | - Ida J van der Klei
- Molecular Cell Biology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, the Netherlands.
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2
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Sinclair R, Wang M, Jawaid MZ, Longkumer T, Aaron J, Rossetti B, Wait E, McDonald K, Cox D, Heddleston J, Wilkop T, Drakakaki G. Four-dimensional quantitative analysis of cell plate development in Arabidopsis using lattice light sheet microscopy identifies robust transition points between growth phases. JOURNAL OF EXPERIMENTAL BOTANY 2024; 75:2829-2847. [PMID: 38436428 PMCID: PMC11282576 DOI: 10.1093/jxb/erae091] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 09/20/2023] [Accepted: 02/29/2024] [Indexed: 03/05/2024]
Abstract
Cell plate formation during cytokinesis entails multiple stages occurring concurrently and requiring orchestrated vesicle delivery, membrane remodelling, and timely deposition of polysaccharides, such as callose. Understanding such a dynamic process requires dissection in time and space; this has been a major hurdle in studying cytokinesis. Using lattice light sheet microscopy (LLSM), we studied cell plate development in four dimensions, through the behavior of yellow fluorescent protein (YFP)-tagged cytokinesis-specific GTPase RABA2a vesicles. We monitored the entire duration of cell plate development, from its first emergence, with the aid of YFP-RABA2a, in both the presence and absence of cytokinetic callose. By developing a robust cytokinetic vesicle volume analysis pipeline, we identified distinct behavioral patterns, allowing the identification of three easily trackable cell plate developmental phases. Notably, the phase transition between phase I and phase II is striking, indicating a switch from membrane accumulation to the recycling of excess membrane material. We interrogated the role of callose using pharmacological inhibition with LLSM and electron microscopy. Loss of callose inhibited the phase transitions, establishing the critical role and timing of the polysaccharide deposition in cell plate expansion and maturation. This study exemplifies the power of combining LLSM with quantitative analysis to decode and untangle such a complex process.
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Affiliation(s)
- Rosalie Sinclair
- Department of Plant Sciences, University of California Davis, Davis, CA, USA
| | - Minmin Wang
- Department of Plant Sciences, University of California Davis, Davis, CA, USA
| | - Muhammad Zaki Jawaid
- Department of Physics and Astronomy, University of California Davis, Davis, CA, USA
| | | | | | | | - Eric Wait
- Janelia Research Campus, Ashburn, VA, USA
| | - Kent McDonald
- Electron Microscope Laboratory, University of California, Berkeley, CA, USA
| | - Daniel Cox
- Department of Physics and Astronomy, University of California Davis, Davis, CA, USA
| | | | - Thomas Wilkop
- Department of Molecular and Cellular Biology, Light Microscopy Imaging Facility, University of California Davis, Davis, CA, USA
| | - Georgia Drakakaki
- Department of Plant Sciences, University of California Davis, Davis, CA, USA
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3
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Prochera A, Muppirala AN, Kuziel GA, Soualhi S, Shepherd A, Sun L, Issac B, Rosenberg HJ, Karim F, Perez K, Smith KH, Archibald TH, Rakoff-Nahoum S, Hagen SJ, Rao M. Enteric glia regulate Paneth cell secretion and intestinal microbial ecology. BIORXIV : THE PREPRINT SERVER FOR BIOLOGY 2024:2024.04.15.589545. [PMID: 38659931 PMCID: PMC11042301 DOI: 10.1101/2024.04.15.589545] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 04/26/2024]
Abstract
Glial cells of the enteric nervous system (ENS) interact closely with the intestinal epithelium and secrete signals that influence epithelial cell proliferation and barrier formation in vitro. Whether these interactions are important in vivo, however, is unclear because previous studies reached conflicting conclusions [1]. To better define the roles of enteric glia in steady state regulation of the intestinal epithelium, we characterized the glia in closest proximity to epithelial cells and found that the majority express PLP1 in both mice and humans. To test their functions using an unbiased approach, we genetically depleted PLP1+ cells in mice and transcriptionally profiled the small and large intestines. Surprisingly, glial loss had minimal effects on transcriptional programs and the few identified changes varied along the gastrointestinal tract. In the ileum, where enteric glia had been considered most essential for epithelial integrity, glial depletion did not drastically alter epithelial gene expression but caused a modest enrichment in signatures of Paneth cells, a secretory cell type important for innate immunity. In the absence of PLP1+ glia, Paneth cell number was intact, but a subset appeared abnormal with irregular and heterogenous cytoplasmic granules, suggesting a secretory deficit. Consistent with this possibility, ileal explants from glial-depleted mice secreted less functional lysozyme than controls with corresponding effects on fecal microbial composition. Collectively, these data suggest that enteric glia do not exert broad effects on the intestinal epithelium but have an essential role in regulating Paneth cell function and gut microbial ecology.
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Affiliation(s)
- Aleksandra Prochera
- Division of Gastroenterology, Department of Pediatrics, Boston Children's Hospital and Harvard Medical School, 300 Longwood Ave, Boston, MA 02115, USA
| | - Anoohya N Muppirala
- Division of Gastroenterology, Department of Pediatrics, Boston Children's Hospital and Harvard Medical School, 300 Longwood Ave, Boston, MA 02115, USA
| | - Gavin A Kuziel
- Division of Gastroenterology, Department of Pediatrics, Boston Children's Hospital and Harvard Medical School, 300 Longwood Ave, Boston, MA 02115, USA
- Division of Infectious Diseases, Department of Pediatrics, Boston Children's Hospital and Harvard Medical School, 300 Longwood Ave, Boston, MA 02115, USA
- Department of Microbiology, Harvard Medical School, Boston, MA 02115, USA
| | - Salima Soualhi
- Division of Gastroenterology, Department of Pediatrics, Boston Children's Hospital and Harvard Medical School, 300 Longwood Ave, Boston, MA 02115, USA
| | - Amy Shepherd
- Division of Gastroenterology, Department of Pediatrics, Boston Children's Hospital and Harvard Medical School, 300 Longwood Ave, Boston, MA 02115, USA
| | - Liang Sun
- Research Computing, Department of Information Technology, Boston Children's Hospital, 300 Longwood Ave, Boston, MA 02115, USA
| | - Biju Issac
- Research Computing, Department of Information Technology, Boston Children's Hospital, 300 Longwood Ave, Boston, MA 02115, USA
| | - Harry J Rosenberg
- Division of Gastroenterology, Department of Pediatrics, Boston Children's Hospital and Harvard Medical School, 300 Longwood Ave, Boston, MA 02115, USA
- Department of Pathology, Beth Israel Deaconess Medical Center, Boston, MA, USA
| | - Farah Karim
- Institute of Human Nutrition, Columbia University Irving Medical Center, New York, NY, USA
| | - Kristina Perez
- Division of Gastroenterology, Department of Pediatrics, Boston Children's Hospital and Harvard Medical School, 300 Longwood Ave, Boston, MA 02115, USA
| | - Kyle H Smith
- Department of Surgery, Beth Israel Deaconess Medical Center, Boston, MA, USA
| | - Tonora H Archibald
- Department of Pathology, Beth Israel Deaconess Medical Center, Boston, MA, USA
| | - Seth Rakoff-Nahoum
- Division of Gastroenterology, Department of Pediatrics, Boston Children's Hospital and Harvard Medical School, 300 Longwood Ave, Boston, MA 02115, USA
- Division of Infectious Diseases, Department of Pediatrics, Boston Children's Hospital and Harvard Medical School, 300 Longwood Ave, Boston, MA 02115, USA
- Department of Microbiology, Harvard Medical School, Boston, MA 02115, USA
| | - Susan J Hagen
- Department of Surgery, Beth Israel Deaconess Medical Center, Boston, MA, USA
| | - Meenakshi Rao
- Division of Gastroenterology, Department of Pediatrics, Boston Children's Hospital and Harvard Medical School, 300 Longwood Ave, Boston, MA 02115, USA
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4
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Tranfield EM, Fabig G, Kurth T, Müller-Reichert T. How to apply the broad toolbox of correlative light and electron microscopy to address a specific biological question. Methods Cell Biol 2024; 187:1-41. [PMID: 38705621 DOI: 10.1016/bs.mcb.2024.02.030] [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: 05/07/2024]
Abstract
Correlative light and electron microscopy (CLEM) is an approach that combines the strength of multiple imaging techniques to obtain complementary information about a given specimen. The "toolbox" for CLEM is broad, making it sometimes difficult to choose an appropriate approach for a given biological question. In this chapter, we provide experimental details for three CLEM approaches that can help the interested reader in designing a personalized CLEM strategy for obtaining ultrastructural data by using transmission electron microscopy (TEM). First, we describe chemical fixation of cells grown on a solid support (broadest approach). Second, we apply high-pressure freezing/freeze substitution to describe cellular ultrastructure (cryo-immobilization approach). Third, we give a protocol for a ultrastructural labeling by immuno-electron microscopy (immuno-EM approach). In addition, we also describe how to overlay fluorescence and electron microscopy images, an approach that is applicable to each of the reported different CLEM strategies. Here we provide step-by step descriptions prior to discussing possible technical problems and variations of these three general schemes to suit different models or different biological questions. This chapter is written for electron microscopists that are new to CLEM and unsure how to begin. Therefore, our protocols are meant to provide basic information with further references that should help the reader get started with applying a tailored strategy for a specific CLEM experiment.
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Affiliation(s)
- Erin M Tranfield
- Electron Microscopy Facility, Instituto Gulbenkian de Ciência, Oeiras, Portugal
| | - Gunar Fabig
- Experimental Center, Faculty of Medicine Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany
| | - Thomas Kurth
- Core Facility Electron Microscopy and Histology Facility, Technology Platform, Center for Molecular and Cellular Bioengineering, Technische Universität Dresden, Dresden, Germany
| | - Thomas Müller-Reichert
- Experimental Center, Faculty of Medicine Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany.
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5
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Li XM, Jenke H, Strauss S, Bazakos C, Mosca G, Lymbouridou R, Kierzkowski D, Neumann U, Naik P, Huijser P, Laurent S, Smith RS, Runions A, Tsiantis M. Cell-cycle-linked growth reprogramming encodes developmental time into leaf morphogenesis. Curr Biol 2024; 34:541-556.e15. [PMID: 38244542 DOI: 10.1016/j.cub.2023.12.050] [Citation(s) in RCA: 2] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/18/2023] [Revised: 12/14/2023] [Accepted: 12/15/2023] [Indexed: 01/22/2024]
Abstract
How is time encoded into organ growth and morphogenesis? We address this question by investigating heteroblasty, where leaf development and form are modified with progressing plant age. By combining morphometric analyses, fate-mapping through live-imaging, computational analyses, and genetics, we identify age-dependent changes in cell-cycle-associated growth and histogenesis that underpin leaf heteroblasty. We show that in juvenile leaves, cell proliferation competence is rapidly released in a "proliferation burst" coupled with fast growth, whereas in adult leaves, proliferative growth is sustained for longer and at a slower rate. These effects are mediated by the SPL9 transcription factor in response to inputs from both shoot age and individual leaf maturation along the proximodistal axis. SPL9 acts by activating CyclinD3 family genes, which are sufficient to bypass the requirement for SPL9 in the control of leaf shape and in heteroblastic reprogramming of cellular growth. In conclusion, we have identified a mechanism that bridges across cell, tissue, and whole-organism scales by linking cell-cycle-associated growth control to age-dependent changes in organ geometry.
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Affiliation(s)
- Xin-Min Li
- Department of Comparative Development and Genetics, Max Planck Institute for Plant Breeding Research, Carl-von-Linné Weg 10, 50829 Cologne, Germany
| | - Hannah Jenke
- Department of Comparative Development and Genetics, Max Planck Institute for Plant Breeding Research, Carl-von-Linné Weg 10, 50829 Cologne, Germany
| | - Sören Strauss
- Department of Comparative Development and Genetics, Max Planck Institute for Plant Breeding Research, Carl-von-Linné Weg 10, 50829 Cologne, Germany
| | - Christos Bazakos
- Department of Comparative Development and Genetics, Max Planck Institute for Plant Breeding Research, Carl-von-Linné Weg 10, 50829 Cologne, Germany
| | - Gabriella Mosca
- Department of Comparative Development and Genetics, Max Planck Institute for Plant Breeding Research, Carl-von-Linné Weg 10, 50829 Cologne, Germany
| | - Rena Lymbouridou
- Department of Comparative Development and Genetics, Max Planck Institute for Plant Breeding Research, Carl-von-Linné Weg 10, 50829 Cologne, Germany
| | - Daniel Kierzkowski
- Department of Comparative Development and Genetics, Max Planck Institute for Plant Breeding Research, Carl-von-Linné Weg 10, 50829 Cologne, Germany
| | - Ulla Neumann
- Central Microscopy (CeMic), Max Planck Institute for Plant Breeding Research, Carl-von-Linné Weg 10, 50829 Cologne, Germany
| | - Purva Naik
- Department of Comparative Development and Genetics, Max Planck Institute for Plant Breeding Research, Carl-von-Linné Weg 10, 50829 Cologne, Germany
| | - Peter Huijser
- Department of Comparative Development and Genetics, Max Planck Institute for Plant Breeding Research, Carl-von-Linné Weg 10, 50829 Cologne, Germany
| | - Stefan Laurent
- Department of Comparative Development and Genetics, Max Planck Institute for Plant Breeding Research, Carl-von-Linné Weg 10, 50829 Cologne, Germany
| | - Richard S Smith
- Department of Comparative Development and Genetics, Max Planck Institute for Plant Breeding Research, Carl-von-Linné Weg 10, 50829 Cologne, Germany
| | - Adam Runions
- Department of Comparative Development and Genetics, Max Planck Institute for Plant Breeding Research, Carl-von-Linné Weg 10, 50829 Cologne, Germany
| | - Miltos Tsiantis
- Department of Comparative Development and Genetics, Max Planck Institute for Plant Breeding Research, Carl-von-Linné Weg 10, 50829 Cologne, Germany.
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6
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Köckert M, Okafornta CW, Hill C, Ryndyk A, Striese C, Müller-Reichert T, Paliulis L, Fabig G. Ultrastructure of the nebenkern during spermatogenesis in the praying mantid Hierodula membranacea. PLoS One 2023; 18:e0285073. [PMID: 37498864 PMCID: PMC10374135 DOI: 10.1371/journal.pone.0285073] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/13/2023] [Accepted: 07/04/2023] [Indexed: 07/29/2023] Open
Abstract
Spermatogenesis leads to the formation of functional sperm cells. Here we have applied high-pressure freezing in combination with transmission electron microscopy (TEM) to study the ultrastructure of sperm development in subadult males of the praying mantid Hierodula membranacea, a species in which spermatogenesis had not previously been studied. We show the ultrastructure of different stages of sperm development in this species. Thorough examination of TEM data and electron tomographic reconstructions revealed interesting structural features of the nebenkern, an organelle composed of fused mitochondria that has been studied in spermatids of other insect species. We have applied serial-section electron tomography of the nebenkern to demonstrate in three dimensions (3D) that this organelle in H. membranacea is composed of two interwoven mitochondrial derivatives, and that the mitochondrial derivatives are connected by a zipper-like structure at opposing positions. Our approach will enable further ultrastructural analyses of the nebenkern in other organisms.
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Affiliation(s)
- Maria Köckert
- Experimental Center, Faculty of Medicine Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany
| | | | - Charlice Hill
- Experimental Center, Faculty of Medicine Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany
| | - Anne Ryndyk
- Experimental Center, Faculty of Medicine Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany
| | - Cynthia Striese
- Experimental Center, Faculty of Medicine Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany
| | - Thomas Müller-Reichert
- Experimental Center, Faculty of Medicine Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany
| | - Leocadia Paliulis
- Biology Department, Bucknell University, Lewisburg, PA, United States of America
| | - Gunar Fabig
- Experimental Center, Faculty of Medicine Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany
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7
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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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8
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Imaging the Plant Cytoskeleton by High-Pressure Freezing and Electron Tomography. Methods Mol Biol 2023; 2604:89-102. [PMID: 36773227 DOI: 10.1007/978-1-0716-2867-6_7] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 02/12/2023]
Abstract
Electron tomography (ET) imaging of high-pressure frozen/freeze-substituted samples provides a unique opportunity to study structural details of organelles and cytoskeletal arrays in plant cells. In this chapter, we discuss approaches for sample preparation by cryofixation at high pressure, freeze substitution, and resin embedding. We also include pipelines for data collection for electron tomography at ambient temperature, tomogram calculation, and segmentation.
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9
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A cryo-fixation protocol to study the structure of the synaptonemal complex. Chromosome Res 2022; 30:385-400. [PMID: 35486207 DOI: 10.1007/s10577-022-09689-2] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/08/2022] [Revised: 03/08/2022] [Accepted: 03/28/2022] [Indexed: 01/25/2023]
Abstract
Genetic variability in sexually reproducing organisms results from an exchange of genetic material between homologous chromosomes. The genetic exchange mechanism is dependent on the synaptonemal complex (SC), a protein structure localized between the homologous chromosomes. The current structural models of the mammalian SC are based on electron microscopy, superresolution, and expansion microscopy studies using chemical fixatives and sample dehydration of gonads, which are methodologies known to produce structural artifacts. To further analyze the structure of the SC, without chemical fixation, we have adapted a cryo-fixation method for electron microscopy where pachytene cells are isolated from mouse testis by FACS, followed by cryo-fixation, cryo-substitution, and electron tomography. In parallel, we performed conventional chemical fixation and electron tomography on mouse seminiferous tubules to compare the SC structure obtained with the two fixation methods. We found several differences in the structure and organization of the SC in cryo-fixed samples when compared to chemically preserved samples. We found the central region of the SC to be wider and the transverse filaments to be more densely packed in the central region of the SC.
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10
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Baranov MV, Ioannidis M, Balahsioui S, Boersma A, de Boer R, Kumar M, Niwa M, Hirayama T, Zhou Q, Hopkins TM, Grijpstra P, Thutupalli S, Sacanna S, van den Bogaart G. Irregular particle morphology and membrane rupture facilitate ion gradients in the lumen of phagosomes. BIOPHYSICAL REPORTS 2022; 2:100069. [PMID: 36425330 PMCID: PMC9680789 DOI: 10.1016/j.bpr.2022.100069] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Figures] [Subscribe] [Scholar Register] [Received: 04/19/2022] [Accepted: 08/08/2022] [Indexed: 06/16/2023]
Abstract
Localized fluxes, production, and/or degradation coupled to limited diffusion are well known to result in stable spatial concentration gradients of biomolecules in the cell. In this study, we demonstrate that this also holds true for small ions, since we found that the close membrane apposition between the membrane of a phagosome and the surface of the cargo particle it encloses, together with localized membrane rupture, suffice for stable gradients of protons and iron cations within the lumen of the phagosome. Our data show that, in phagosomes containing hexapod-shaped silica colloid particles, the phagosomal membrane is ruptured at the positions of the tips of the rods, but not at other positions. This results in the confined leakage at these positions of protons and iron from the lumen of the phagosome into the cytosol. In contrast, acidification and iron accumulation still occur at the positions of the phagosomes nearer to the cores of the particles. Our study strengthens the concept that coupling metabolic and signaling reaction cascades can be spatially confined by localized limited diffusion.
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Affiliation(s)
- Maksim V. Baranov
- Department of Molecular Immunology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, Netherlands
| | - Melina Ioannidis
- Department of Molecular Immunology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, Netherlands
| | - Sami Balahsioui
- Department of Molecular Immunology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, Netherlands
| | - Auke Boersma
- Department of Molecular Immunology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, Netherlands
| | - Rinse de Boer
- Department of Molecular Immunology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, Netherlands
| | - Manoj Kumar
- Simons Centre for the Study of Living Machines, National Centre for Biological Sciences, Tata Institute of Fundamental Research, Bangalore, India
| | - Masato Niwa
- Laboratory of Pharmaceutical and Medicinal Chemistry, Gifu Pharmaceutical University, 1–25–4, Daigaku-nishi, Gifu 201–1196, Japan
| | - Tasuku Hirayama
- Laboratory of Pharmaceutical and Medicinal Chemistry, Gifu Pharmaceutical University, 1–25–4, Daigaku-nishi, Gifu 201–1196, Japan
| | - Qintian Zhou
- Molecular Design Institute, Department of Chemistry, New York University, New York, NY, United States
| | - Terrence M. Hopkins
- Molecular Design Institute, Department of Chemistry, New York University, New York, NY, United States
| | - Pieter Grijpstra
- Department of Molecular Immunology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, Netherlands
| | - Shashi Thutupalli
- Simons Centre for the Study of Living Machines, National Centre for Biological Sciences, Tata Institute of Fundamental Research, Bangalore, India
- nternational Centre for Theoretical Sciences, Tata Institute of Fundamental Research, Bangalore, India
| | - Stefano Sacanna
- Molecular Design Institute, Department of Chemistry, New York University, New York, NY, United States
| | - Geert van den Bogaart
- Department of Molecular Immunology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, Netherlands
- Department of Medical Biology and Pathology, University Medical Center Groningen, Groningen, Netherlands
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11
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Bélanger S, Berensmann H, Baena V, Duncan K, Meyers BC, Narayan K, Czymmek KJ. A versatile enhanced freeze-substitution protocol for volume electron microscopy. Front Cell Dev Biol 2022; 10:933376. [PMID: 36003147 PMCID: PMC9393620 DOI: 10.3389/fcell.2022.933376] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/30/2022] [Accepted: 07/08/2022] [Indexed: 11/18/2022] Open
Abstract
Volume electron microscopy, a powerful approach to generate large three-dimensional cell and tissue volumes at electron microscopy resolutions, is rapidly becoming a routine tool for understanding fundamental and applied biological questions. One of the enabling factors for its adoption has been the development of conventional fixation protocols with improved heavy metal staining. However, freeze-substitution with organic solvent-based fixation and staining has not realized the same level of benefit. Here, we report a straightforward approach including osmium tetroxide, acetone and up to 3% water substitution fluid (compatible with traditional or fast freeze-substitution protocols), warm-up and transition from organic solvent to aqueous 2% osmium tetroxide. Once fully hydrated, samples were processed in aqueous based potassium ferrocyanide, thiocarbohydrazide, osmium tetroxide, uranyl acetate and lead acetate before resin infiltration and polymerization. We observed a consistent and substantial increase in heavy metal staining across diverse and difficult-to-fix test organisms and tissue types, including plant tissues (Hordeum vulgare), nematode (Caenorhabditis elegans) and yeast (Saccharomyces cerevisiae). Our approach opens new possibilities to combine the benefits of cryo-preservation with enhanced contrast for volume electron microscopy in diverse organisms.
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Affiliation(s)
| | - Heather Berensmann
- Center for Molecular Microscopy, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD, United States
- Cancer Research Technology Program, Frederick National Laboratory for Cancer Research, Frederick, MD, United States
| | - Valentina Baena
- Center for Molecular Microscopy, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD, United States
- Cancer Research Technology Program, Frederick National Laboratory for Cancer Research, Frederick, MD, United States
| | - Keith Duncan
- Donald Danforth Plant Science Center, Saint Louis, MO, United States
| | - Blake C. Meyers
- Donald Danforth Plant Science Center, Saint Louis, MO, United States
- Division of Plant Science and Technology, University of Missouri–Columbia, Columbia, MO, United States
| | - Kedar Narayan
- Center for Molecular Microscopy, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD, United States
- Cancer Research Technology Program, Frederick National Laboratory for Cancer Research, Frederick, MD, 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
- *Correspondence: Kirk J. Czymmek,
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12
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Peddie CJ, Genoud C, Kreshuk A, Meechan K, Micheva KD, Narayan K, Pape C, Parton RG, Schieber NL, Schwab Y, Titze B, Verkade P, Aubrey A, Collinson LM. Volume electron microscopy. NATURE REVIEWS. METHODS PRIMERS 2022; 2:51. [PMID: 37409324 PMCID: PMC7614724 DOI: 10.1038/s43586-022-00131-9] [Citation(s) in RCA: 35] [Impact Index Per Article: 17.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Accepted: 05/10/2022] [Indexed: 07/07/2023]
Abstract
Life exists in three dimensions, but until the turn of the century most electron microscopy methods provided only 2D image data. Recently, electron microscopy techniques capable of delving deep into the structure of cells and tissues have emerged, collectively called volume electron microscopy (vEM). Developments in vEM have been dubbed a quiet revolution as the field evolved from established transmission and scanning electron microscopy techniques, so early publications largely focused on the bioscience applications rather than the underlying technological breakthroughs. However, with an explosion in the uptake of vEM across the biosciences and fast-paced advances in volume, resolution, throughput and ease of use, it is timely to introduce the field to new audiences. In this Primer, we introduce the different vEM imaging modalities, the specialized sample processing and image analysis pipelines that accompany each modality and the types of information revealed in the data. We showcase key applications in the biosciences where vEM has helped make breakthrough discoveries and consider limitations and future directions. We aim to show new users how vEM can support discovery science in their own research fields and inspire broader uptake of the technology, finally allowing its full adoption into mainstream biological imaging.
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Affiliation(s)
- Christopher J. Peddie
- Electron Microscopy Science Technology Platform, The Francis Crick Institute, London, UK
| | - Christel Genoud
- Electron Microscopy Facility, Faculty of Biology and Medicine, University of Lausanne, Lausanne, Switzerland
| | - Anna Kreshuk
- Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Kimberly Meechan
- Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
- Present address: Faculty of Biosciences, Heidelberg University, Heidelberg, Germany
| | - Kristina D. Micheva
- Department of Molecular and Cellular Physiology, Stanford University, Palo Alto, CA, USA
| | - Kedar Narayan
- Center for Molecular Microscopy, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA
- Cancer Research Technology Program, Frederick National Laboratory for Cancer Research, Frederick, MD, USA
| | - Constantin Pape
- Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Robert G. Parton
- The Institute for Molecular Bioscience, The University of Queensland, Brisbane, Queensland, Australia
- Centre for Microscopy and Microanalysis, The University of Queensland, Brisbane, Queensland, Australia
| | - Nicole L. Schieber
- Centre for Microscopy and Microanalysis, The University of Queensland, Brisbane, Queensland, Australia
| | - Yannick Schwab
- Cell Biology and Biophysics Unit/ Electron Microscopy Core Facility, European Molecular Biology Laboratory, Heidelberg, Germany
| | | | - Paul Verkade
- School of Biochemistry, University of Bristol, Bristol, UK
| | - Aubrey Aubrey
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA, USA
| | - Lucy M. Collinson
- Electron Microscopy Science Technology Platform, The Francis Crick Institute, London, UK
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13
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Yamaguchi M, Takahashi-Nakaguchi A, Uematsu K, Naito K, Sato-Okamoto M, Ishiwata K, Naraoka S, Chibana H. Electron Microscopy of Mouse Tissues by Sandwich Freezing and Freeze-Substitution. CYTOLOGIA 2022. [DOI: 10.1508/cytologia.87.149] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.5] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/28/2022]
Affiliation(s)
| | | | | | - Kumiko Naito
- Department of Diabetes, Metabolism and Endocrinology, Chiba University Hospital
| | | | - Kazuki Ishiwata
- Department of Endocrinology, Hematology and Gerontology, Chiba University Hospital
| | - Sakino Naraoka
- Graduate School of Science and Engineering, Advanced Science and Engineering Biology, Chiba University
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14
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Reagan BC, Dunlap JR, Burch-Smith TM. Focused Ion Beam-Scanning Electron Microscopy for Investigating Plasmodesmal Densities. METHODS IN MOLECULAR BIOLOGY (CLIFTON, N.J.) 2022; 2457:109-123. [PMID: 35349135 DOI: 10.1007/978-1-0716-2132-5_6] [Citation(s) in RCA: 2] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Subscribe] [Scholar Register] [Indexed: 12/15/2022]
Abstract
Plasmodesmata (PD) facilitate the exchange of nutrients and signaling molecules between neighboring plant cells, and they are therefore essential for proper growth and development. PD have been studied extensively in efforts to elucidate the ultrastructure of individual PD nanopores and the distribution of PD in a variety of cell walls. These studies often involved the use of serial ultrathin sections and manual quantification of PD by transmission electron microscopy (TEM). In recent years, a variety of techniques that offer more amenable approaches for quantifying PD distribution have been reported. Here, we describe the quantification of PD densities using the serial scanning electron microscopy technique called focused ion beam-scanning electron microscopy (FIB-SEM). For this, resin-embedded samples prepared by standard TEM methods undergo successive rounds of imaging by SEM interspersed with milling of the sample surface by a focused beam of gallium ions to reveal a new surface. In this way, the details of the sample are sequentially revealed and imaged. Over the course of a few hours, repetitive milling and imaging facilitates the automated collection of nanometer-resolution data of several μm of sample depth. FIB-SEM can be targeted to interrogate specific cell walls and cell wall junctions, and the subsequent three-dimensional renderings of the data can be used to visualize the ultrastructural details of the sample. PD densities can then be rapidly quantified by calculating the number of PD per μm2 of cell wall observed in the renderings.
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Affiliation(s)
- Brandon C Reagan
- Department of Biochemistry & Cellular and Molecular Biology, University of Tennessee, Knoxville, TN, USA.,Department of Plant Biology, Michigan State University, East Lansing, MI, USA
| | - John R Dunlap
- The Joint Institute for Advanced Materials, University of Tennessee, Knoxville, TN, USA
| | - Tessa M Burch-Smith
- Department of Biochemistry & Cellular and Molecular Biology, University of Tennessee, Knoxville, TN, USA. .,Donald Danforth Plant Science Center, Saint Louis, MO, USA.
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15
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Shankar R, Lettman MM, Whisler W, Frankel EB, Audhya A. The ESCRT machinery directs quality control over inner nuclear membrane architecture. Cell Rep 2022; 38:110263. [PMID: 35045304 PMCID: PMC8801257 DOI: 10.1016/j.celrep.2021.110263] [Citation(s) in RCA: 11] [Impact Index Per Article: 5.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/16/2021] [Revised: 11/21/2021] [Accepted: 12/21/2021] [Indexed: 11/20/2022] Open
Abstract
The late-acting endosomal sorting complex required for transport (ESCRT) machinery has been implicated in facilitating the resealing of the nuclear envelope (NE) after mitosis, enabling compartmentalization of the genome away from the cytoplasm. Here, we leverage the stereotypic first division of the C. elegans embryo to identify additional functions of the ESCRT machinery in maintaining the structure of the inner nuclear membrane. Specifically, impaired ESCRT function results in a defect in the pruning of inner nuclear membrane invaginations, which arise normally during NE reformation and expansion. Additionally, in combination with a hypomorphic mutation that interferes with assembly of the underlying nuclear lamina, inhibition of ESCRT function significantly perturbs NE architecture and increases chromosome segregation defects, resulting in penetrant embryonic lethality. Our findings highlight links between ESCRT-mediated inner nuclear membrane remodeling, maintenance of nuclear envelope morphology, and the preservation of the genome during early development. In this study, Shankar et al. demonstrate that defects in ESCRT machinery functions impair pruning of inner nuclear membrane invaginations that form normally after mitotic exit as the nuclear envelope undergoes expansion. These findings highlight a critical role for the ESCRT machinery in the maintenance of inner nuclear membrane morphology.
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Affiliation(s)
- Raakhee Shankar
- Department of Biomolecular Chemistry, University of Wisconsin School of Medicine and Public Health, Madison, WI 53705, USA
| | - Molly M Lettman
- Department of Biomolecular Chemistry, University of Wisconsin School of Medicine and Public Health, Madison, WI 53705, USA
| | - William Whisler
- Department of Biomolecular Chemistry, University of Wisconsin School of Medicine and Public Health, Madison, WI 53705, USA
| | - Elisa B Frankel
- Department of Biomolecular Chemistry, University of Wisconsin School of Medicine and Public Health, Madison, WI 53705, USA
| | - Anjon Audhya
- Department of Biomolecular Chemistry, University of Wisconsin School of Medicine and Public Health, Madison, WI 53705, USA.
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16
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Richardson AC, Fišerová J, Goldberg MW. NPC Structure in Model Organisms: Transmission Electron Microscopy and Immunogold Labeling Using High-Pressure Freezing/Freeze Substitution of Yeast, Worms, and Plants. Methods Mol Biol 2022; 2502:439-459. [PMID: 35412255 DOI: 10.1007/978-1-0716-2337-4_28] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 06/14/2023]
Abstract
The nuclear pore complex (NPC) is a large elaborate structure embedded within the nuclear envelope, and intimately linked to the cytoskeleton, nucleoskeleton, and chromatin. Many different cargoes pass through its central channel and along the membrane at its periphery. The NPC is dismantled and reassembly, fully or partially, every cell cycle. In post-mitotic cells it consists of a combination of hyper-stable and highly dynamic proteins. Because of its size, dynamics, heterogeneity and integration, it is not possible to understand its structure and molecular function by any one, or even several, methods. For decades, and to this day, thin section transmission electron microscopy (TEM) has been a central tool for understanding the NPC, its associations, dynamics and role in transport as it can uniquely answer questions concerning fine structural detail within a cellular context. Using immunogold labeling specific components can also be identified within the ultrastructural context. Model organisms such as Saccharomyces cerevisiae are also central to NPC studies but have not been used extensively in structural work. This is because the cell wall presents difficulties with structural preservation and processing for TEM. In recent years, high-pressure freezing and freeze substitution have overcome these problems, as well as opened up methods to combine immunogold labeling with detailed structural analysis. Other model organisms such as the worm Caenorhabditis elegans and the plant Arabidopsis thaliana have been underused for similar reasons, but with similar solutions, which we present here. There are also many advantages to using these methods, adapted for use in mammalian systems, due to the instant nature of the initial fixation, to capture rapid processes such as nuclear transport, and preservation of dynamic membranes.
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Affiliation(s)
| | - Jindřiška Fišerová
- Department of Biology of the Cell Nucleus, Institute of Molecular Genetics AS CR, Prague, Czech Republic
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17
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Sankoh AF, Burch-Smith TM. Approaches for investigating plasmodesmata and effective communication. CURRENT OPINION IN PLANT BIOLOGY 2021; 64:102143. [PMID: 34826658 DOI: 10.1016/j.pbi.2021.102143] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Received: 06/23/2021] [Revised: 10/13/2021] [Accepted: 10/19/2021] [Indexed: 06/13/2023]
Abstract
Plasmodesmata (PD) are integral plant cell wall components that provide routes for intercellular communication, signaling, and resource sharing. They are therefore essential for plant growth and survival. Much effort has been put forth to understand how PD are generated and their structure is refined for function and to determine how they regulate intercellular trafficking. This review provides an overview of some of the approaches that have been used to study PD structure and function, highlighting those that may be more widely adopted to address questions of PD cell biology and function. Extending our focus on the importance of communication, we address how effective communication strategies can increase diversity and accessibility in the research laboratory, focusing on challenges faced by our deaf/hard-of-hearing colleagues, and highlight successful approaches to including them in the research laboratory.
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Affiliation(s)
- Amie F Sankoh
- Department of Biochemistry & Cellular and Molecular Biology, University of Tennessee, Knoxville, TN 37996, United States
| | - Tessa M Burch-Smith
- Department of Biochemistry & Cellular and Molecular Biology, University of Tennessee, Knoxville, TN 37996, United States.
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18
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Mohr W, Lehnen N, Ahmerkamp S, Marchant HK, Graf JS, Tschitschko B, Yilmaz P, Littmann S, Gruber-Vodicka H, Leisch N, Weber M, Lott C, Schubert CJ, Milucka J, Kuypers MMM. Terrestrial-type nitrogen-fixing symbiosis between seagrass and a marine bacterium. Nature 2021; 600:105-109. [PMID: 34732889 PMCID: PMC8636270 DOI: 10.1038/s41586-021-04063-4] [Citation(s) in RCA: 37] [Impact Index Per Article: 12.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/29/2021] [Accepted: 09/22/2021] [Indexed: 01/23/2023]
Abstract
Symbiotic N2-fixing microorganisms have a crucial role in the assimilation of nitrogen by eukaryotes in nitrogen-limited environments1-3. Particularly among land plants, N2-fixing symbionts occur in a variety of distantly related plant lineages and often involve an intimate association between host and symbiont2,4. Descriptions of such intimate symbioses are lacking for seagrasses, which evolved around 100 million years ago from terrestrial flowering plants that migrated back to the sea5. Here we describe an N2-fixing symbiont, 'Candidatus Celerinatantimonas neptuna', that lives inside seagrass root tissue, where it provides ammonia and amino acids to its host in exchange for sugars. As such, this symbiosis is reminiscent of terrestrial N2-fixing plant symbioses. The symbiosis between Ca. C. neptuna and its host Posidonia oceanica enables highly productive seagrass meadows to thrive in the nitrogen-limited Mediterranean Sea. Relatives of Ca. C. neptuna occur worldwide in coastal ecosystems, in which they may form similar symbioses with other seagrasses and saltmarsh plants. Just like N2-fixing microorganisms might have aided the colonization of nitrogen-poor soils by early land plants6, the ancestors of Ca. C. neptuna and its relatives probably enabled flowering plants to invade nitrogen-poor marine habitats, where they formed extremely efficient blue carbon ecosystems7.
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Affiliation(s)
- Wiebke Mohr
- Max Planck Institute for Marine Microbiology, Bremen, Germany.
| | - Nadine Lehnen
- Max Planck Institute for Marine Microbiology, Bremen, Germany
| | | | | | - Jon S Graf
- Max Planck Institute for Marine Microbiology, Bremen, Germany
| | | | - Pelin Yilmaz
- Max Planck Institute for Marine Microbiology, Bremen, Germany
- Data Science Research Group, Institute for Artificial Intelligence in Medicine, University Hospital Essen, Essen, Germany
| | - Sten Littmann
- Max Planck Institute for Marine Microbiology, Bremen, Germany
| | | | - Nikolaus Leisch
- Max Planck Institute for Marine Microbiology, Bremen, Germany
| | | | | | - Carsten J Schubert
- Swiss Federal Institute of Aquatic Science and Technology (Eawag), Department of Surface Waters-Research and Management, Kastanienbaum, Switzerland
| | - Jana Milucka
- Max Planck Institute for Marine Microbiology, Bremen, Germany
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19
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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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20
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Wood BM, Baena V, Huang H, Jorgens DM, Terasaki M, Kornberg TB. Cytonemes with complex geometries and composition extend into invaginations of target cells. J Cell Biol 2021; 220:211896. [PMID: 33734293 PMCID: PMC7980254 DOI: 10.1083/jcb.202101116] [Citation(s) in RCA: 11] [Impact Index Per Article: 3.7] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/19/2021] [Revised: 02/13/2021] [Accepted: 02/23/2021] [Indexed: 12/16/2022] Open
Abstract
Cytonemes are specialized filopodia that mediate paracrine signaling in Drosophila and other animals. Studies using fluorescence confocal microscopy (CM) established their general paths, cell targets, and essential roles in signaling. To investigate details unresolvable by CM, we used high-pressure freezing and EM to visualize cytoneme structures, paths, contents, and contacts. We observed cytonemes previously seen by CM in the Drosophila wing imaginal disc system, including disc, tracheal air sac primordium (ASP), and myoblast cytonemes, and identified cytonemes extending into invaginations of target cells, and cytonemes connecting ASP cells and connecting myoblasts. Diameters of cytoneme shafts vary between repeating wide (206 ± 51.8 nm) and thin (55.9 ± 16.2 nm) segments. Actin, ribosomes, and membranous compartments are present throughout; rough ER and mitochondria are in wider proximal sections. These results reveal novel structural features of filopodia and provide a basis for understanding cytoneme cell biology and function.
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Affiliation(s)
- Brent M Wood
- Cardiovascular Research Institute, University of California, San Francisco, San Francisco, CA
| | - Valentina Baena
- Department of Cell Biology, University of Connecticut Health Center, Farmington, CT
| | - Hai Huang
- Cardiovascular Research Institute, University of California, San Francisco, San Francisco, CA
| | - Danielle M Jorgens
- Electron Microscope Laboratory, University of California, Berkeley, Berkeley, CA
| | - Mark Terasaki
- Department of Cell Biology, University of Connecticut Health Center, Farmington, CT
| | - Thomas B Kornberg
- Cardiovascular Research Institute, University of California, San Francisco, San Francisco, CA
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21
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Preparation of Cultured Cells Using High-Pressure Freezing and Freeze Substitution for Subsequent 2D or 3D Visualization in the Transmission Electron Microscope. Methods Mol Biol 2021. [PMID: 32833218 DOI: 10.1007/978-1-0716-0900-2_19] [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] [Indexed: 08/03/2024]
Abstract
Transmission electron microscopy (TEM) is an invaluable technique used for imaging the ultrastructure of samples, and it is particularly useful when determining virus-host interactions at a cellular level. The environment inside a TEM is not favorable for biological material (high vacuum and high energy electrons). Also biological samples have little or no intrinsic electron contrast and rarely do they naturally exist in very thin sheets, as is required for optimum resolution in the TEM. To prepare these samples for imaging in the TEM therefore requires extensive processing which can alter the ultrastructure of the material. Here we describe a method which aims to minimize preparation artifacts by freezing the samples at high pressure to instantaneously preserve ultrastructural detail, then rapidly substituting the ice with resin to provide a firm matrix which can be cut into thin sections for imaging. Thicker sections of this material can also be imaged and reconstructed into 3D volumes using electron tomography.
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22
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Ökmen B, Schwammbach D, Bakkeren G, Neumann U, Doehlemann G. The Ustilago hordei-Barley Interaction Is a Versatile System for Characterization of Fungal Effectors. J Fungi (Basel) 2021; 7:86. [PMID: 33513785 PMCID: PMC7912019 DOI: 10.3390/jof7020086] [Citation(s) in RCA: 6] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/08/2020] [Revised: 01/19/2021] [Accepted: 01/22/2021] [Indexed: 12/02/2022] Open
Abstract
Obligate biotrophic fungal pathogens, such as Blumeria graminis and Puccinia graminis, are amongst the most devastating plant pathogens, causing dramatic yield losses in many economically important crops worldwide. However, a lack of reliable tools for the efficient genetic transformation has hampered studies into the molecular basis of their virulence or pathogenicity. In this study, we present the Ustilago hordei-barley pathosystem as a model to characterize effectors from different plant pathogenic fungi. We generate U. hordei solopathogenic strains, which form infectious filaments without the presence of a compatible mating partner. Solopathogenic strains are suitable for heterologous expression system for fungal virulence factors. A highly efficient Crispr/Cas9 gene editing system is made available for U. hordei. In addition, U. hordei infection structures during barley colonization are analyzed using transmission electron microscopy, showing that U. hordei forms intracellular infection structures sharing high similarity to haustoria formed by obligate rust and powdery mildew fungi. Thus, U. hordei has high potential as a fungal expression platform for functional studies of heterologous effector proteins in barley.
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Affiliation(s)
- Bilal Ökmen
- BioCenter, Institute for Plant Sciences, University of Cologne, Zülpicher Straße 47a, 50674 Cologne, Germany
| | - Daniela Schwammbach
- Max Planck Institute for Terrestrial Microbiology, Karl von Frisch Straße, 35043 Marburg, Germany;
| | - Guus Bakkeren
- Summerland Research and Development Centre, Agriculture and Agri-Food Canada, Summerland, BC V0H 1Z0, Canada;
| | - Ulla Neumann
- Central Microscopy, Max Planck Institute for Plant Breeding Research, Carl-von-Linné-Weg 10, 50829 Cologne, Germany;
| | - Gunther Doehlemann
- BioCenter, Institute for Plant Sciences, University of Cologne, Zülpicher Straße 47a, 50674 Cologne, Germany
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23
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Current Microscopy Strategies to Image Fungal Vesicles: From the Intracellular Trafficking and Secretion to the Inner Structure of Isolated Vesicles. Curr Top Microbiol Immunol 2021; 432:139-159. [DOI: 10.1007/978-3-030-83391-6_11] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 10/19/2022]
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24
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Zielińska A, Savietto A, de Sousa Borges A, Martinez D, Berbon M, Roelofsen JR, Hartman AM, de Boer R, Van der Klei IJ, Hirsch AKH, Habenstein B, Bramkamp M, Scheffers DJ. Flotillin-mediated membrane fluidity controls peptidoglycan synthesis and MreB movement. eLife 2020; 9:e57179. [PMID: 32662773 PMCID: PMC7360373 DOI: 10.7554/elife.57179] [Citation(s) in RCA: 24] [Impact Index Per Article: 6.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/24/2020] [Accepted: 06/12/2020] [Indexed: 01/07/2023] Open
Abstract
The bacterial plasma membrane is an important cellular compartment. In recent years it has become obvious that protein complexes and lipids are not uniformly distributed within membranes. Current hypotheses suggest that flotillin proteins are required for the formation of complexes of membrane proteins including cell-wall synthetic proteins. We show here that bacterial flotillins are important factors for membrane fluidity homeostasis. Loss of flotillins leads to a decrease in membrane fluidity that in turn leads to alterations in MreB dynamics and, as a consequence, in peptidoglycan synthesis. These alterations are reverted when membrane fluidity is restored by a chemical fluidizer. In vitro, the addition of a flotillin increases membrane fluidity of liposomes. Our data support a model in which flotillins are required for direct control of membrane fluidity rather than for the formation of protein complexes via direct protein-protein interactions.
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Affiliation(s)
- Aleksandra Zielińska
- Molecular Microbiology, Groningen Biomolecular Sciences and Biotechnology Institute, University of GroningenGroningenNetherlands
| | - Abigail Savietto
- Biozentrum, Ludwig-Maximilians-Universität MünchenMünchenGermany
- Institute for General Microbiology, Christian-Albrechts-UniversityKielGermany
| | - Anabela de Sousa Borges
- Molecular Microbiology, Groningen Biomolecular Sciences and Biotechnology Institute, University of GroningenGroningenNetherlands
| | - Denis Martinez
- Institute of Chemistry & Biology of Membranes & Nanoobjects (UMR5248 CBMN), IECB, CNRS, Université Bordeaux, Institut Polytechnique BordeauxPessacFrance
| | - Melanie Berbon
- Institute of Chemistry & Biology of Membranes & Nanoobjects (UMR5248 CBMN), IECB, CNRS, Université Bordeaux, Institut Polytechnique BordeauxPessacFrance
| | - Joël R Roelofsen
- Molecular Microbiology, Groningen Biomolecular Sciences and Biotechnology Institute, University of GroningenGroningenNetherlands
| | - Alwin M Hartman
- Department of Drug Design and Optimization (DDOP), Helmholtz-Institute for Pharmaceutical Research Saarland (HIPS) - Helmholtz Centre for Infection Research (HZI)SaarbrückenGermany
- Department of Pharmacy, Saarland UniversitySaarbrückenGermany
- Stratingh Institute for Chemistry, University of GroningenGroningenNetherlands
| | - Rinse de Boer
- Molecular Cell Biology, Groningen Biomolecular Sciences and Biotechnology Institute, University of GroningenGroningenNetherlands
| | - Ida J Van der Klei
- Molecular Cell Biology, Groningen Biomolecular Sciences and Biotechnology Institute, University of GroningenGroningenNetherlands
| | - Anna KH Hirsch
- Department of Drug Design and Optimization (DDOP), Helmholtz-Institute for Pharmaceutical Research Saarland (HIPS) - Helmholtz Centre for Infection Research (HZI)SaarbrückenGermany
- Department of Pharmacy, Saarland UniversitySaarbrückenGermany
- Stratingh Institute for Chemistry, University of GroningenGroningenNetherlands
| | - Birgit Habenstein
- Institute of Chemistry & Biology of Membranes & Nanoobjects (UMR5248 CBMN), IECB, CNRS, Université Bordeaux, Institut Polytechnique BordeauxPessacFrance
| | - Marc Bramkamp
- Biozentrum, Ludwig-Maximilians-Universität MünchenMünchenGermany
- Institute for General Microbiology, Christian-Albrechts-UniversityKielGermany
| | - Dirk-Jan Scheffers
- Molecular Microbiology, Groningen Biomolecular Sciences and Biotechnology Institute, University of GroningenGroningenNetherlands
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Wolff G, Melia CE, Snijder EJ, Bárcena M. Double-Membrane Vesicles as Platforms for Viral Replication. Trends Microbiol 2020; 28:1022-1033. [PMID: 32536523 PMCID: PMC7289118 DOI: 10.1016/j.tim.2020.05.009] [Citation(s) in RCA: 188] [Impact Index Per Article: 47.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/01/2020] [Revised: 05/09/2020] [Accepted: 05/14/2020] [Indexed: 12/12/2022]
Abstract
Viruses, as obligate intracellular parasites, exploit cellular pathways and resources in a variety of fascinating ways. A striking example of this is the remodelling of intracellular membranes into specialized structures that support the replication of positive-sense ssRNA (+RNA) viruses infecting eukaryotes. These distinct forms of virus-induced structures include double-membrane vesicles (DMVs), found during viral infections as diverse and notorious as those of coronaviruses, enteroviruses, noroviruses, or hepatitis C virus. Our understanding of these DMVs has evolved over the past 15 years thanks to advances in imaging techniques and modern molecular biology tools. In this article, we review contemporary understanding of the biogenesis, structure, and function of virus-induced DMVs as well as the open questions posed by these intriguing structures.
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Affiliation(s)
- Georg Wolff
- Section Electron Microscopy, Department of Cell and Chemical Biology, Leiden University Medical Center, Leiden, The Netherlands
| | - Charlotte E Melia
- Section Electron Microscopy, Department of Cell and Chemical Biology, Leiden University Medical Center, Leiden, The Netherlands
| | - Eric J Snijder
- Molecular Virology laboratory, Department of Medical Microbiology, Leiden University Medical Center, Leiden, The Netherlands
| | - Montserrat Bárcena
- Section Electron Microscopy, Department of Cell and Chemical Biology, Leiden University Medical Center, Leiden, The Netherlands.
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26
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Hook KA, Fisher HS. Methodological considerations for examining the relationship between sperm morphology and motility. Mol Reprod Dev 2020; 87:633-649. [PMID: 32415812 PMCID: PMC7329573 DOI: 10.1002/mrd.23346] [Citation(s) in RCA: 10] [Impact Index Per Article: 2.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/25/2020] [Revised: 04/10/2020] [Accepted: 04/21/2020] [Indexed: 12/12/2022]
Abstract
Sperm cells of all taxa share a common goal to reach and fertilize an ovum, yet sperm are one of the most diverse cell types in nature. While the structural diversity of these cells is well recognized, the functional significance of variation in sperm design remains elusive. An important function of spermatozoa is a need to migrate toward the ova, often over long distances in a foreign environment, which may include a complex and hostile female reproductive tract. Several comparative and experimental studies have attempted to address the link between sperm morphology and motility, yet the conclusions drawn from these studies are often inconsistent, even within the same taxa. Much of what we know about the functional significance of sperm design in internally fertilizing species has been gleaned from in vitro studies, for which experimental parameters often vary among studies. We propose that discordant results from these studies are in part due to a lack of consistency of methods, conditions that do not replicate those of the female reproductive tract, and the overuse of simple linear measures of sperm shape. Within this review, we provide a toolkit for imaging, quantifying, and analyzing sperm morphology and movement patterns for in vitro studies and discuss emerging approaches. Results from studies linking morphology to motility enhance our understanding of the evolution of adaptive sperm traits and the mechanisms that regulate fertility, thus offering new insights into methods used in assisted reproductive technologies in animal science, conservation and public health.
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Affiliation(s)
- Kristin A. Hook
- Department of Biology, University of Maryland, College Park, U.S.A
| | - Heidi S. Fisher
- Department of Biology, University of Maryland, College Park, U.S.A
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27
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Riesterer JL, López CS, Stempinski ES, Williams M, Loftis K, Stoltz K, Thibault G, Lanicault C, Williams T, Gray JW. A workflow for visualizing human cancer biopsies using large-format electron microscopy. Methods Cell Biol 2020; 158:163-181. [PMID: 32423648 DOI: 10.1016/bs.mcb.2020.01.005] [Citation(s) in RCA: 10] [Impact Index Per Article: 2.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/23/2023]
Abstract
Recent developments in large format electron microscopy have enabled generation of images that provide detailed ultrastructural information on normal and diseased cells and tissues. Analyses of these images increase our understanding of cellular organization and interactions and disease-related changes therein. In this manuscript, we describe a workflow for two-dimensional (2D) and three-dimensional (3D) imaging, including both optical and scanning electron microscopy (SEM) methods, that allow pathologists and cancer biology researchers to identify areas of interest from human cancer biopsies. The protocols and mounting strategies described in this workflow are compatible with 2D large format EM mapping, 3D focused ion beam-SEM and serial block face-SEM. The flexibility to use diverse imaging technologies available at most academic institutions makes this workflow useful and applicable for most life science samples. Volumetric analysis of the biopsies studied here revealed morphological, organizational and ultrastructural aspects of the tumor cells and surrounding environment that cannot be revealed by conventional 2D EM imaging. Our results indicate that although 2D EM is still an important tool in many areas of diagnostic pathology, 3D images of ultrastructural relationships between both normal and cancerous cells, in combination with their extracellular matrix, enables cancer researchers and pathologists to better understand the progression of the disease and identify potential therapeutic targets.
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Affiliation(s)
- Jessica L Riesterer
- OHSU Center for Spatial Systems Biomedicine, Oregon Health and Sciences University, Portland, OR, United States; Multiscale Microscopy Core, Oregon Health and Sciences University, Portland, OR, United States.
| | - Claudia S López
- OHSU Center for Spatial Systems Biomedicine, Oregon Health and Sciences University, Portland, OR, United States; Multiscale Microscopy Core, Oregon Health and Sciences University, Portland, OR, United States; Pacific Northwest Center for CryoEM, Oregon Health and Sciences University, Portland, OR, United States.
| | - Erin S Stempinski
- OHSU Center for Spatial Systems Biomedicine, Oregon Health and Sciences University, Portland, OR, United States; Multiscale Microscopy Core, Oregon Health and Sciences University, Portland, OR, United States
| | - Melissa Williams
- OHSU Center for Spatial Systems Biomedicine, Oregon Health and Sciences University, Portland, OR, United States; Multiscale Microscopy Core, Oregon Health and Sciences University, Portland, OR, United States
| | - Kevin Loftis
- OHSU Center for Spatial Systems Biomedicine, Oregon Health and Sciences University, Portland, OR, United States
| | - Kevin Stoltz
- OHSU Center for Spatial Systems Biomedicine, Oregon Health and Sciences University, Portland, OR, United States
| | - Guillaume Thibault
- OHSU Center for Spatial Systems Biomedicine, Oregon Health and Sciences University, Portland, OR, United States
| | - Christian Lanicault
- Department of Pathology, Oregon Health and Sciences University, Portland, OR, United States
| | - Todd Williams
- Department of Pathology, Oregon Health and Sciences University, Portland, OR, United States
| | - Joe W Gray
- OHSU Center for Spatial Systems Biomedicine, Oregon Health and Sciences University, Portland, OR, United States.
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28
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Flechsler J, Heimerl T, Pickl C, Rachel R, Stierhof YD, Klingl A. 2D and 3D immunogold localization on (epoxy) ultrathin sections with and without osmium tetroxide. Microsc Res Tech 2020; 83:691-705. [PMID: 32057162 DOI: 10.1002/jemt.23459] [Citation(s) in RCA: 9] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/31/2019] [Revised: 01/13/2020] [Accepted: 02/04/2020] [Indexed: 11/07/2022]
Abstract
For nearly 50 years immunogold labeling on ultrathin sections has been successfully used for protein localization in laboratories worldwide. In theory and in practice, this method has undergone continual improvement over time. In this study, we carefully analyzed circulating protocols for postembedding labeling to find out if they are still valid under modern laboratory conditions, and in addition, we tested unconventional protocols. For this, we investigated immunolabeling of Epon-embedded cells, immunolabeling of cells treated with osmium, and the binding behavior of differently sized gold particles. Here we show that (in contrast to widespread belief) immunolabeling of Epon-embedded cells and of cells treated with osmium tetroxide is actually working. Furthermore, we established a "speed protocol" for immunolabeling by reducing antibody incubation times. Finally, we present our results on three-dimensional immunogold labeling.
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Affiliation(s)
- Jennifer Flechsler
- Plant Development and Electron Microscopy, Department of Biology I, Munchen, Germany
| | - Thomas Heimerl
- LOEWE Centre for Synthetic Microbiology (SYNMIKRO), Philipps-Universität Marburg, Marburg, Germany
| | - Carolin Pickl
- Plant Development and Electron Microscopy, Department of Biology I, Munchen, Germany
| | - Reinhard Rachel
- Institute of Microbiology and Centre for Electron Microscopy, University of Regensburg, Regensburg, Germany
| | - York-Dieter Stierhof
- Microscopy, Center for Plant Molecular Biology, University of Tübingen, Tübingen, Germany
| | - Andreas Klingl
- Plant Development and Electron Microscopy, Department of Biology I, Munchen, Germany
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29
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Sele M, Wernitznig S, Lipovšek S, Radulović S, Haybaeck J, Birkl-Toeglhofer AM, Wodlej C, Kleinegger F, Sygulla S, Leoni M, Ropele S, Leitinger G. Optimization of ultrastructural preservation of human brain for transmission electron microscopy after long post-mortem intervals. Acta Neuropathol Commun 2019; 7:144. [PMID: 31481118 PMCID: PMC6724377 DOI: 10.1186/s40478-019-0794-3] [Citation(s) in RCA: 15] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/01/2019] [Accepted: 08/24/2019] [Indexed: 12/16/2022] Open
Abstract
Electron microscopy (EM) provides the necessary resolution to visualize the finer structures of nervous tissue morphology, which is important to understand healthy and pathological conditions in the brain. However, for the interpretation of the micrographs the tissue preservation is crucial. The quality of the tissue structure is mostly influenced by the post mortem interval (PMI), the time of death until the preservation of the tissue. Therefore, the aim of this study was to optimize the preparation-procedure for the human frontal lobe to preserve the ultrastructure as well as possible despite the long PMIs. Combining chemical pre- and post-fixation with cryo-fixation and cryo-substitution ("hybrid freezing"), it was possible to improve the preservation of the neuronal profiles of human brain samples compared to the "standard" epoxy resin embedding method. In conclusion short PMIs are generally desirable but up to a PMI of 16 h the ultrastructure can be preserved on an acceptable level with a high contrast using the "hybrid freezing" protocol described here.
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Whiddon KT, Gudneppanavar R, Hammer TJ, West DA, Konopka MC. Fluorescence-based analysis of the intracytoplasmic membranes of type I methanotrophs. Microb Biotechnol 2019; 12:1024-1033. [PMID: 31264365 PMCID: PMC6680624 DOI: 10.1111/1751-7915.13458] [Citation(s) in RCA: 13] [Impact Index Per Article: 2.6] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/20/2019] [Accepted: 06/16/2019] [Indexed: 12/15/2022] Open
Abstract
Most methanotrophic bacteria maintain intracytoplasmic membranes which house the methane-oxidizing enzyme, particulate methane monooxygenase. Previous studies have primarily used transmission electron microscopy or cryo-electron microscopy to look at the structure of these membranes or lipid extraction methods to determine the per cent of cell dry weight composed of lipids. We show an alternative approach using lipophilic membrane probes and other fluorescent dyes to assess the extent of intracytoplasmic membrane formation in living cells. This fluorescence method is sensitive enough to show not only the characteristic shift in intracytoplasmic membrane formation that is present when methanotrophs are grown with or without copper, but also differences in intracytoplasmic membrane levels at intermediate copper concentrations. This technique can also be employed to monitor dynamic intracytoplasmic membrane changes in the same cell in real time under changing growth conditions. We anticipate that this approach will be of use to researchers wishing to visualize intracytoplasmic membranes who may not have access to electron microscopes. It will also have the capability to relate membrane changes in individual living cells to other measurements by fluorescence labelling or other single-cell analysis methods.
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Affiliation(s)
| | | | - Theodore J. Hammer
- Department of ChemistryThe University of AkronAkronOHUSA
- Department of Polymer ScienceThe University of AkronAkronOHUSA
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31
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Otegui MS, Pennington JG. Electron tomography in plant cell biology. Microscopy (Oxf) 2019; 68:69-79. [PMID: 30452668 DOI: 10.1093/jmicro/dfy133] [Citation(s) in RCA: 11] [Impact Index Per Article: 2.2] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/30/2018] [Revised: 10/15/2018] [Accepted: 10/31/2018] [Indexed: 12/11/2022] Open
Abstract
Electron tomography (ET) approaches are based on the imaging of a biological specimen at different tilt angles by transmission electron microscopy (TEM). ET can be applied to both plastic-embedded and frozen samples. Technological advancements in TEM, direct electron detection, automated image collection, and imaging processing algorithms allow for 2-7-nm scale axial resolution in tomographic reconstructions of cells and organelles. In this review, we discussed the application of ET in plant cell biology and new opportunities for imaging plant cells by cryo-ET and other 3D electron microscopy approaches.
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Affiliation(s)
- Marisa S Otegui
- Department of Botany, University of Wisconsin-Madison, 430 Lincoln Drive, Madison WI, USA.,Laboratory of Molecular and Cellular Biology, University of Wisconsin-Madison, 1525 Linden Drive, Madison WI, USA.,Department of Genetics, University of Wisconsin-Madison, 425 Henry Mall, Madison WI, USA
| | - Jannice G Pennington
- Institute for Molecular Virology, University of Wisconsin-Madison, 1525 Linden Drive, Madison WI, USA.,Howard Hughes Medical Institute, University of Wisconsin-Madison, Madison, USA
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32
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King GA, Goodman JS, Schick JG, Chetlapalli K, Jorgens DM, McDonald KL, Ünal E. Meiotic cellular rejuvenation is coupled to nuclear remodeling in budding yeast. eLife 2019; 8:e47156. [PMID: 31397671 PMCID: PMC6711709 DOI: 10.7554/elife.47156] [Citation(s) in RCA: 41] [Impact Index Per Article: 8.2] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/26/2019] [Accepted: 07/19/2019] [Indexed: 12/12/2022] Open
Abstract
Production of healthy gametes in meiosis relies on the quality control and proper distribution of both nuclear and cytoplasmic contents. Meiotic differentiation naturally eliminates age-induced cellular damage by an unknown mechanism. Using time-lapse fluorescence microscopy in budding yeast, we found that nuclear senescence factors - including protein aggregates, extrachromosomal ribosomal DNA circles, and abnormal nucleolar material - are sequestered away from chromosomes during meiosis II and subsequently eliminated. A similar sequestration and elimination process occurs for the core subunits of the nuclear pore complex in both young and aged cells. Nuclear envelope remodeling drives the formation of a membranous compartment containing the sequestered material. Importantly, de novo generation of plasma membrane is required for the sequestration event, preventing the inheritance of long-lived nucleoporins and senescence factors into the newly formed gametes. Our study uncovers a new mechanism of nuclear quality control and provides insight into its function in meiotic cellular rejuvenation.
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Affiliation(s)
- Grant A King
- Department of Molecular and Cell BiologyUniversity of California, BerkeleyBerkeleyUnited States
| | - Jay S Goodman
- Department of Molecular and Cell BiologyUniversity of California, BerkeleyBerkeleyUnited States
| | - Jennifer G Schick
- Department of Molecular and Cell BiologyUniversity of California, BerkeleyBerkeleyUnited States
| | - Keerthana Chetlapalli
- Department of Molecular and Cell BiologyUniversity of California, BerkeleyBerkeleyUnited States
| | - Danielle M Jorgens
- Electron Microscope LabUniversity of California, BerkeleyBerkeleyUnited States
| | - Kent L McDonald
- Electron Microscope LabUniversity of California, BerkeleyBerkeleyUnited States
| | - Elçin Ünal
- Department of Molecular and Cell BiologyUniversity of California, BerkeleyBerkeleyUnited States
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33
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Keene DR, Tufa SF. Connective Tissue Ultrastructure: A Direct Comparison between Conventional Specimen Preparation and High-Pressure Freezing/Freeze-Substitution. Anat Rec (Hoboken) 2019; 303:1514-1526. [PMID: 31251834 DOI: 10.1002/ar.24211] [Citation(s) in RCA: 4] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/31/2018] [Revised: 12/28/2018] [Accepted: 01/18/2019] [Indexed: 11/11/2022]
Abstract
It is generally agreed within the microscopy community that the quality of ultrastructure within the connective tissue matrix resulting from high-pressure freezing followed by freeze-substitution (HPF/FS) far exceeds that gained following the "conventional" preparation method, which includes aqueous fixation, dehydration, and embedding. Exposure to cryogen at high pressure is the only cryopreservation method capable of vitrifying tissue structure to a depth exceeding 200 μm. Cells within connective tissues prepared by HPF/FS are universally larger, filling the commonly seen void at the juncture between cell and matrix. Without significant shrinkage of cells and the coincident extraction of the cytosolic components, well-resolved organelles are less clustered within an expanded cytosol. Much of the artifact from "conventional" methods occurs as large space filling and also smaller fibril-associated proteoglycans are extracted during fixation. However, the visualization of some matrix features by electron microscopy is actually dependent on the collapse or extraction of these "masking" components. Herein, we argue that an impression of ultrastructure within commonly studied matrices, in particular skin, is best gained following the evaluation of both conventional preparations and tissue prepared by HPF/FS. Anat Rec, 2019. © 2019 American Association for Anatomy.
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Affiliation(s)
- Douglas R Keene
- Shriners Hospital for Children Micro-Imaging Center, Portland, Oregon.,Department of Biomechanical Engineering, Oregon Health Sciences University, Portland, Oregon.,Department of Medical Genetics, Oregon Health Sciences University, Portland, Oregon
| | - Sara F Tufa
- Shriners Hospital for Children Micro-Imaging Center, Portland, Oregon
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34
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The architecture of cell differentiation in choanoflagellates and sponge choanocytes. PLoS Biol 2019; 17:e3000226. [PMID: 30978201 PMCID: PMC6481868 DOI: 10.1371/journal.pbio.3000226] [Citation(s) in RCA: 44] [Impact Index Per Article: 8.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/01/2018] [Revised: 04/24/2019] [Accepted: 03/27/2019] [Indexed: 11/19/2022] Open
Abstract
Although collar cells are conserved across animals and their closest relatives, the choanoflagellates, little is known about their ancestry, their subcellular architecture, or how they differentiate. The choanoflagellate Salpingoeca rosetta expresses genes necessary for animal development and can alternate between unicellular and multicellular states, making it a powerful model for investigating the origin of animal multicellularity and mechanisms underlying cell differentiation. To compare the subcellular architecture of solitary collar cells in S. rosetta with that of multicellular ‘rosette’ colonies and collar cells in sponges, we reconstructed entire cells in 3D through transmission electron microscopy on serial ultrathin sections. Structural analysis of our 3D reconstructions revealed important differences between single and colonial choanoflagellate cells, with colonial cells exhibiting a more amoeboid morphology consistent with higher levels of macropinocytotic activity. Comparison of multiple reconstructed rosette colonies highlighted the variable nature of cell sizes, cell–cell contact networks, and colony arrangement. Importantly, we uncovered the presence of elongated cells in some rosette colonies that likely represent a distinct and differentiated cell type, pointing toward spatial cell differentiation. Intercellular bridges within choanoflagellate colonies displayed a variety of morphologies and connected some but not all neighbouring cells. Reconstruction of sponge choanocytes revealed ultrastructural commonalities but also differences in major organelle composition in comparison to choanoflagellates. Together, our comparative reconstructions uncover the architecture of cell differentiation in choanoflagellates and sponge choanocytes and constitute an important step in reconstructing the cell biology of the last common ancestor of animals. 3D electron microscopy of choanoflagellates and sponge choanocytes reveals a remarkable variety of cell architecture and suggests that cell type differentiation may have been present in the stem lineage leading to the animals. Choanoflagellates are microscopic aquatic organisms that can alternate between single-celled and multicellular states, and sequencing of their genomes has revealed that choanoflagellates are the closest single-celled relatives of animals. Moreover, choanoflagellates are a form of ‘collar cell’—a cell type crowned by an array of finger-like microvilli and a single, whip-like flagellum. This cell type is also found throughout the animal kingdom; therefore, studying the structure of the choanoflagellate collar cell can shed light on how this cell type and animal multicellularity might have evolved. We used electron microscopy to reconstruct in 3D the total subcellular composition of single-celled and multicellular choanoflagellates as well as the collar cells from a marine sponge, which represents an early-branching animal lineage. We found differences between single-celled and multicellular choanoflagellates in structures associated with cellular energetics, membrane trafficking, and cell morphology. Likewise, we describe a complex system of cell–cell connections associated with multicellular choanoflagellates. Finally, comparison of choanoflagellates and sponge collar cells revealed subcellular differences associated with feeding and cellular energetics. Taken together, this study is an important step forward in reconstructing the biology of the last common ancestor of the animals.
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35
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Mulcahy B, Witvliet D, Holmyard D, Mitchell J, Chisholm AD, Meirovitch Y, Samuel ADT, Zhen M. A Pipeline for Volume Electron Microscopy of the Caenorhabditis elegans Nervous System. Front Neural Circuits 2018; 12:94. [PMID: 30524248 PMCID: PMC6262311 DOI: 10.3389/fncir.2018.00094] [Citation(s) in RCA: 24] [Impact Index Per Article: 4.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/16/2018] [Accepted: 10/08/2018] [Indexed: 01/01/2023] Open
Abstract
The “connectome,” a comprehensive wiring diagram of synaptic connectivity, is achieved through volume electron microscopy (vEM) analysis of an entire nervous system and all associated non-neuronal tissues. White et al. (1986) pioneered the fully manual reconstruction of a connectome using Caenorhabditis elegans. Recent advances in vEM allow mapping new C. elegans connectomes with increased throughput, and reduced subjectivity. Current vEM studies aim to not only fill the remaining gaps in the original connectome, but also address fundamental questions including how the connectome changes during development, the nature of individuality, sexual dimorphism, and how genetic and environmental factors regulate connectivity. Here we describe our current vEM pipeline and projected improvements for the study of the C. elegans nervous system and beyond.
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Affiliation(s)
- Ben Mulcahy
- Lunenfeld-Tanenbaum Research Institute, Mount Sinai Hospital, Toronto, ON, Canada
| | - Daniel Witvliet
- Lunenfeld-Tanenbaum Research Institute, Mount Sinai Hospital, Toronto, ON, Canada.,Department of Molecular Genetics, University of Toronto, Toronto, ON, Canada
| | - Douglas Holmyard
- Department of Pathology and Laboratory Medicine, Mount Sinai Hospital, Toronto, ON, Canada.,Nanoscale Biomedical Imaging Facility, The Hospital for Sick Children, Peter Gilgan Centre for Research and Learning, Toronto, ON, Canada
| | - James Mitchell
- Center for Brain Science, Department of Physics, Harvard University, Cambridge, MA, United States
| | - Andrew D Chisholm
- Section of Cell and Developmental Biology, Division of Biological Sciences, University of California, San Diego, La Jolla, CA, United States
| | - Yaron Meirovitch
- Department of Physiology, University of Toronto, Toronto, ON, Canada
| | - Aravinthan D T Samuel
- Center for Brain Science, Department of Physics, Harvard University, Cambridge, MA, United States
| | - Mei Zhen
- Lunenfeld-Tanenbaum Research Institute, Mount Sinai Hospital, Toronto, ON, Canada.,Department of Molecular Genetics, University of Toronto, Toronto, ON, Canada.,Department of Physiology, University of Toronto, Toronto, ON, Canada.,Department of Cell and Systems Biology, University of Toronto, Toronto, ON, Canada
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36
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Enhanced method for High Spatial Resolution surface imaging and analysis of fungal spores using Scanning Electron Microscopy. Sci Rep 2018; 8:16278. [PMID: 30390022 PMCID: PMC6214942 DOI: 10.1038/s41598-018-34629-8] [Citation(s) in RCA: 11] [Impact Index Per Article: 1.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/12/2018] [Accepted: 10/16/2018] [Indexed: 11/25/2022] Open
Abstract
Efficient, fast and new micro-analytical methods for characterization of ultrastructures of fungal spores with electron microscopy are very much required and essential. SEM analysis of biological materials, especially fungi, requires optimal preparation of the specimen and often requires the usage of dried samples which demands a challenging sample preparation. In the present investigation, we described a fast and improved method for the preparation of fungal specimen for scanning electron microscopy (SEM). The fungus, Curvularia lunata was grown on the surface of sterile Whatman No.1 filter paper which was overlaid on Potato Dextrose Agar (PDA) medium, gold coated immediately after removal from the growth medium and subjected to imaging. Generally, SEM imaging is done with samples that were fixed with chemical fixatives, dehydrated and gold coated specimens, but here we describe an easy and more efficient sample preparation for SEM which enabled enhanced image quality and precision visualization of fungal cells, especially the spores. The developed method has enabled the analysis of even the robust samples like fungal spores that to eliminating special temperature requirement. The ultimate goal was to develop an improved protocol/method applied to analysis of fungal spores with greater coverage about fungal specimen preparation. This method permits the use of rapid sample preparation and will allow us to imaging of individual spore or conidia structures in the context of fungal cell architecture which clarifies our understanding in fungal taxonomy/biology.
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37
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Abstract
Cell-cell adhesion is important for cell growth, tissue development, and neural network formation. Structures of cell adhesion molecules have been widely studied by crystallography, revealing the molecular details of adhesion interfaces. However, due to technical limitations, the overall structure and organization of adhesion molecules at cell adhesion interfaces has not been fully investigated. Here, we combine electron microscopy and other biophysical methods to characterize the structure of cell-cell adhesion mediated by the cell adhesion molecule Sidekick (Sidekick-1 and Sidekick-2) and obtain 3D views of the Sidekick-mediated adhesion interfaces as well as the organization of Sidekick molecules between cell membranes by electron tomography. The results suggest that the Ig-like domains and the fibronectin III (FnIII) domains of Sidekicks play different roles in cell adhesion. The Ig-like domains mediate the homophilic transinteractions bridging adjacent cells, while the FnIII domains interact with membranes, resulting in a tight adhesion interface between cells that may contribute to the specificity and plasticity of cell-cell contacts during cell growth and neural development.
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38
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Reipert S, Goldammer H, Richardson C, Goldberg MW, Hawkins TJ, Hollergschwandtner E, Kaufmann WA, Antreich S, Stierhof YD. Agitation Modules: Flexible Means to Accelerate Automated Freeze Substitution. J Histochem Cytochem 2018; 66:903-921. [PMID: 29969056 DOI: 10.1369/0022155418786698] [Citation(s) in RCA: 17] [Impact Index Per Article: 2.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/14/2022] Open
Abstract
For ultrafast fixation of biological samples to avoid artifacts, high-pressure freezing (HPF) followed by freeze substitution (FS) is preferred over chemical fixation at room temperature. After HPF, samples are maintained at low temperature during dehydration and fixation, while avoiding damaging recrystallization. This is a notoriously slow process. McDonald and Webb demonstrated, in 2011, that sample agitation during FS dramatically reduces the necessary time. Then, in 2015, we (H.G. and S.R.) introduced an agitation module into the cryochamber of an automated FS unit and demonstrated that the preparation of algae could be shortened from days to a couple of hours. We argued that variability in the processing, reproducibility, and safety issues are better addressed using automated FS units. For dissemination, we started low-cost manufacturing of agitation modules for two of the most widely used FS units, the Automatic Freeze Substitution Systems, AFS(1) and AFS2, from Leica Microsystems, using three dimensional (3D)-printing of the major components. To test them, several labs independently used the modules on a wide variety of specimens that had previously been processed by manual agitation, or without agitation. We demonstrate that automated processing with sample agitation saves time, increases flexibility with respect to sample requirements and protocols, and produces data of at least as good quality as other approaches.
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Affiliation(s)
- Siegfried Reipert
- Core Facility Cell Imaging and Ultrastructure Research, University of Vienna, Vienna, Austria
| | - Helmuth Goldammer
- Core Facility Cell Imaging and Ultrastructure Research, University of Vienna, Vienna, Austria
| | | | - Martin W Goldberg
- Department of Biosciences, Durham University, Durham, United Kingdom
| | - Timothy J Hawkins
- Department of Biosciences, Durham University, Durham, United Kingdom
| | | | - Walter A Kaufmann
- Electron Microscopy Facility, Institute of Science and Technology Austria, Klosterneuburg, Austria
| | - Sebastian Antreich
- Core Facility Cell Imaging and Ultrastructure Research, University of Vienna, Vienna, Austria
| | - York-Dieter Stierhof
- Center for Plant Molecular Biology (ZMBP), Microscopy, University of Tübingen, Tübingen, Germany
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39
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Huebinger J, Grabenbauer M. Self-Pressurized Rapid Freezing as Cryo-Fixation Method for Electron Microscopy and Cryopreservation of Living Cells. ACTA ACUST UNITED AC 2018; 79:e47. [PMID: 29924483 DOI: 10.1002/cpcb.47] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/08/2022]
Abstract
Reduction or complete prevention of ice crystal formation during freezing of biological specimens is mandatory for two important biological applications: (1) cryopreservation of living cells or tissues for long-term storage, and (2) cryo-fixation for ultrastructural investigations by electron microscopy. Here, a protocol that is fast, easy-to-use, and suitable for both cryo-fixation and cryopreservation is described. Samples are rapidly cooled in tightly sealed metal tubes of high thermal diffusivity and then plunged into a liquid cryogen. Due to the fast cooling speed and high-pressure buildup internally in the confined volume of the metal tubes, ice crystal formation is reduced or completely prevented, resulting in vitrification of the sample. For cryopreservation, however, a similar principle applies to prevent ice crystal formation during re-warming. A detailed description of procedures for cooling (and re-warming) of biological samples using this technique is provided. © 2018 by John Wiley & Sons, Inc.
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Affiliation(s)
- Jan Huebinger
- Department of Systemic Cell Biology, Max-Planck-Institute of Molecular Physiology, Dortmund, Germany
| | - Markus Grabenbauer
- Institute for Anatomy and Cell Biology, University of Heidelberg, Heidelberg, Germany
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Suhard D, Tessier C, Manens L, Rebière F, Tack K, Agarande M, Guéguen Y. Intracellular uranium distribution: Comparison of cryogenic fixation versus chemical fixation methods for SIMS analysis. Microsc Res Tech 2018; 81:855-864. [PMID: 29737608 PMCID: PMC6221105 DOI: 10.1002/jemt.23047] [Citation(s) in RCA: 6] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/20/2017] [Revised: 01/30/2018] [Accepted: 04/17/2018] [Indexed: 12/30/2022]
Abstract
Localization of uranium within cells is mandatory for the comprehension of its cellular mechanism of toxicity. Secondary Ion Mass Spectrometry (SIMS) has recently shown its interest to detect and localize uranium at very low levels within the cells. This technique requires a specific sample preparation similar to the one used for Transmission Electronic Microscopy, achieved by implementing different chemical treatments to preserve as much as possible the living configuration uranium distribution into the observed sample. This study aims to compare the bioaccumulation sites of uranium within liver or kidney cells after chemical fixation and cryomethods preparations of the samples: SIMS analysis of theses samples show the localization of uranium soluble forms in the cell cytoplasm and nucleus with a more homogenous distribution when using cryopreparation probably due to the diffusible portion of uranium inside the cytoplasm.
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Affiliation(s)
- D Suhard
- Institut de Radioprotection et de Sûreté Nucléaire (IRSN), PSE-SANTE/SESANE, Fontenay-aux-Roses, France
| | - C Tessier
- Institut de Radioprotection et de Sûreté Nucléaire (IRSN), PSE-SANTE/SESANE, Fontenay-aux-Roses, France
| | - L Manens
- Institut de Radioprotection et de Sûreté Nucléaire (IRSN), PSE-SANTE/SESANE, Fontenay-aux-Roses, France
| | - F Rebière
- Institut de Radioprotection et de Sûreté Nucléaire (IRSN), PSE-SANTE/SESANE, Fontenay-aux-Roses, France
| | - K Tack
- Institut de Radioprotection et de Sûreté Nucléaire (IRSN), PSE-SANTE/SESANE, Fontenay-aux-Roses, France
| | - M Agarande
- Institut de Radioprotection et de Sûreté Nucléaire (IRSN), PSE-ENV/SAME, Le Vésinet, France
| | - Y Guéguen
- Institut de Radioprotection et de Sûreté Nucléaire (IRSN), PSE-SANTE/SESANE, Fontenay-aux-Roses, France
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Webb RI, Schieber NL. Volume Scanning Electron Microscopy: Serial Block-Face Scanning Electron Microscopy Focussed Ion Beam Scanning Electron Microscopy. BIOLOGICAL AND MEDICAL PHYSICS, BIOMEDICAL ENGINEERING 2018. [DOI: 10.1007/978-3-319-68997-5_5] [Citation(s) in RCA: 7] [Impact Index Per Article: 1.2] [Reference Citation Analysis] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 12/12/2022]
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Pinkert MA, Hortensius RA, Ogle BM, Eliceiri KW. Imaging the Cardiac Extracellular Matrix. ADVANCES IN EXPERIMENTAL MEDICINE AND BIOLOGY 2018; 1098:21-44. [PMID: 30238364 DOI: 10.1007/978-3-319-97421-7_2] [Citation(s) in RCA: 9] [Impact Index Per Article: 1.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 12/11/2022]
Abstract
Cardiovascular disease is the global leading cause of death. One route to address this problem is using biomedical imaging to measure the molecules and structures that surround cardiac cells. This cellular microenvironment, known as the cardiac extracellular matrix, changes in composition and organization during most cardiac diseases and in response to many cardiac treatments. Measuring these changes with biomedical imaging can aid in understanding, diagnosing, and treating heart disease. This chapter supports those efforts by reviewing representative methods for imaging the cardiac extracellular matrix. It first describes the major biological targets of ECM imaging, including the primary imaging target of fibrillar collagen. Then it discusses the imaging methods, describing their current capabilities and limitations. It categorizes the imaging methods into two main categories: organ-scale noninvasive methods and cellular-scale invasive methods. Noninvasive methods can be used on patients, but only a few are clinically available, and others require further development to be used in the clinic. Invasive methods are the most established and can measure a variety of properties, but they cannot be used on live patients. Finally, the chapter concludes with a perspective on future directions and applications of biomedical imaging technologies.
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Affiliation(s)
- Michael A Pinkert
- Laboratory for Optical and Computational Instrumentation and Department of Medical Physics, University of Wisconsin at Madison, Madison, WI, USA.,Morgridge Institute for Research, Madison, WI, USA
| | - Rebecca A Hortensius
- Department of Biomedical Engineering, University of Minnesota, Minneapolis, MN, USA
| | - Brenda M Ogle
- Department of Biomedical Engineering, University of Minnesota, Minneapolis, MN, USA
| | - Kevin W Eliceiri
- Laboratory for Optical and Computational Instrumentation and Department of Medical Physics, University of Wisconsin at Madison, Madison, WI, USA. .,Morgridge Institute for Research, Madison, WI, USA.
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Erickson HP. How bacterial cell division might cheat turgor pressure - a unified mechanism of septal division in Gram-positive and Gram-negative bacteria. Bioessays 2017; 39. [PMID: 28699183 DOI: 10.1002/bies.201700045] [Citation(s) in RCA: 22] [Impact Index Per Article: 3.1] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/09/2022]
Abstract
An important question for bacterial cell division is how the invaginating septum can overcome the turgor force generated by the high osmolarity of the cytoplasm. I suggest that it may not need to. Several studies in Gram-negative bacteria have shown that the periplasm is isoosmolar with the cytoplasm. Indirect evidence suggests that this is also true for Gram-positive bacteria. In this case the invagination of the septum takes place within the uniformly high osmotic pressure environment, and does not have to fight turgor pressure. A related question is how the V-shaped constriction of Gram-negative bacteria relates to the plate-like septum of Gram-positive bacteria. I collected evidence that Gram-negative bacteria have a latent capability of forming plate-like septa, and present a model in which septal division is the basic mechanism in both Gram-positive and Gram-negative bacteria.
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Affiliation(s)
- Harold P Erickson
- Department of Cell Biology, Duke University Medical School, Durham, NC 27710, USA
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Blancard C, Salin B. Plunge Freezing: A Tool for the Ultrastructural and Immunolocalization Studies of Suspension Cells in Transmission Electron Microscopy. J Vis Exp 2017. [PMID: 28518127 DOI: 10.3791/54874] [Citation(s) in RCA: 5] [Impact Index Per Article: 0.7] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 10/31/2022] Open
Abstract
Transmission Electron Microscopy (TEM) is an extraordinary tool for studying cell ultrastructure, in order to localize proteins and visualize macromolecular complexes at very high resolution. However, to get as close as possible to the native state, perfect sample preservation is required. Conventional electron microscopy (EM) fixation with aldehydes, for instance, does not provide good ultrastructural preservation. The slow penetration of fixatives induces cell reorganization and loss of various cell components. Therefore, conventional EM fixation does not allow for an instantaneous stabilization and preservation of structures and antigenicity. The best choice for examining intracellular events is to use cryofixation followed by the freeze-substitution fixation method that keeps cells in their native state. High-pressure freezing/freeze-substitution, which preserves the integrity of cellular ultrastructure, is the most commonly used method, but requires expensive equipment. Here, an easy-to-use and low-cost freeze fixation method followed by freeze-substitution for suspension cell cultures is presented.
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Affiliation(s)
- Corinne Blancard
- Institut de Biochimie et Génétique Cellulaires, Centre National de la Recherche Scientifique, UMR 5095, Université de Bordeaux
| | - Bénédicte Salin
- Institut de Biochimie et Génétique Cellulaires, Centre National de la Recherche Scientifique, UMR 5095, Université de Bordeaux;
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Gang Y, Zhou H, Jia Y, Liu L, Liu X, Rao G, Li L, Wang X, Lv X, Xiong H, Yang Z, Luo Q, Gong H, Zeng S. Embedding and Chemical Reactivation of Green Fluorescent Protein in the Whole Mouse Brain for Optical Micro-Imaging. Front Neurosci 2017; 11:121. [PMID: 28352214 PMCID: PMC5349086 DOI: 10.3389/fnins.2017.00121] [Citation(s) in RCA: 27] [Impact Index Per Article: 3.9] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/09/2017] [Accepted: 02/27/2017] [Indexed: 02/02/2023] Open
Abstract
Resin embedding has been widely applied to fixing biological tissues for sectioning and imaging, but has long been regarded as incompatible with green fluorescent protein (GFP) labeled sample because it reduces fluorescence. Recently, it has been reported that resin-embedded GFP-labeled brain tissue can be imaged with high resolution. In this protocol, we describe an optimized protocol for resin embedding and chemical reactivation of fluorescent protein labeled mouse brain, we have used mice as experiment model, but the protocol should be applied to other species. This method involves whole brain embedding and chemical reactivation of the fluorescent signal in resin-embedded tissue. The whole brain embedding process takes a total of 7 days. The duration of chemical reactivation is ~2 min for penetrating 4 μm below the surface in the resin-embedded brain. This protocol provides an efficient way to prepare fluorescent protein labeled sample for high-resolution optical imaging. This kind of sample was demonstrated to be imaged by various optical micro-imaging methods. Fine structures labeled with GFP across a whole brain can be detected.
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Affiliation(s)
- Yadong Gang
- Britton Chance Center for Biomedical Photonics, Wuhan National Laboratory for Optoelectronics-Huazhong University of Science and TechnologyWuhan, China
- Key Laboratory of Biomedical Photonics of Ministry of Education, Department of Biomedical Engineering, Huazhong University of Science and TechnologyWuhan, China
| | - Hongfu Zhou
- Britton Chance Center for Biomedical Photonics, Wuhan National Laboratory for Optoelectronics-Huazhong University of Science and TechnologyWuhan, China
- Key Laboratory of Biomedical Photonics of Ministry of Education, Department of Biomedical Engineering, Huazhong University of Science and TechnologyWuhan, China
| | - Yao Jia
- Britton Chance Center for Biomedical Photonics, Wuhan National Laboratory for Optoelectronics-Huazhong University of Science and TechnologyWuhan, China
- Key Laboratory of Biomedical Photonics of Ministry of Education, Department of Biomedical Engineering, Huazhong University of Science and TechnologyWuhan, China
| | - Ling Liu
- Britton Chance Center for Biomedical Photonics, Wuhan National Laboratory for Optoelectronics-Huazhong University of Science and TechnologyWuhan, China
- Key Laboratory of Biomedical Photonics of Ministry of Education, Department of Biomedical Engineering, Huazhong University of Science and TechnologyWuhan, China
| | - Xiuli Liu
- Britton Chance Center for Biomedical Photonics, Wuhan National Laboratory for Optoelectronics-Huazhong University of Science and TechnologyWuhan, China
- Key Laboratory of Biomedical Photonics of Ministry of Education, Department of Biomedical Engineering, Huazhong University of Science and TechnologyWuhan, China
| | - Gong Rao
- Britton Chance Center for Biomedical Photonics, Wuhan National Laboratory for Optoelectronics-Huazhong University of Science and TechnologyWuhan, China
- Key Laboratory of Biomedical Photonics of Ministry of Education, Department of Biomedical Engineering, Huazhong University of Science and TechnologyWuhan, China
| | - Longhui Li
- Britton Chance Center for Biomedical Photonics, Wuhan National Laboratory for Optoelectronics-Huazhong University of Science and TechnologyWuhan, China
- Key Laboratory of Biomedical Photonics of Ministry of Education, Department of Biomedical Engineering, Huazhong University of Science and TechnologyWuhan, China
| | - Xiaojun Wang
- Britton Chance Center for Biomedical Photonics, Wuhan National Laboratory for Optoelectronics-Huazhong University of Science and TechnologyWuhan, China
- Key Laboratory of Biomedical Photonics of Ministry of Education, Department of Biomedical Engineering, Huazhong University of Science and TechnologyWuhan, China
| | - Xiaohua Lv
- Britton Chance Center for Biomedical Photonics, Wuhan National Laboratory for Optoelectronics-Huazhong University of Science and TechnologyWuhan, China
- Key Laboratory of Biomedical Photonics of Ministry of Education, Department of Biomedical Engineering, Huazhong University of Science and TechnologyWuhan, China
| | - Hanqing Xiong
- Britton Chance Center for Biomedical Photonics, Wuhan National Laboratory for Optoelectronics-Huazhong University of Science and TechnologyWuhan, China
- Key Laboratory of Biomedical Photonics of Ministry of Education, Department of Biomedical Engineering, Huazhong University of Science and TechnologyWuhan, China
| | - Zhongqin Yang
- Britton Chance Center for Biomedical Photonics, Wuhan National Laboratory for Optoelectronics-Huazhong University of Science and TechnologyWuhan, China
- Key Laboratory of Biomedical Photonics of Ministry of Education, Department of Biomedical Engineering, Huazhong University of Science and TechnologyWuhan, China
| | - Qingming Luo
- Britton Chance Center for Biomedical Photonics, Wuhan National Laboratory for Optoelectronics-Huazhong University of Science and TechnologyWuhan, China
- Key Laboratory of Biomedical Photonics of Ministry of Education, Department of Biomedical Engineering, Huazhong University of Science and TechnologyWuhan, China
| | - Hui Gong
- Britton Chance Center for Biomedical Photonics, Wuhan National Laboratory for Optoelectronics-Huazhong University of Science and TechnologyWuhan, China
- Key Laboratory of Biomedical Photonics of Ministry of Education, Department of Biomedical Engineering, Huazhong University of Science and TechnologyWuhan, China
| | - Shaoqun Zeng
- Britton Chance Center for Biomedical Photonics, Wuhan National Laboratory for Optoelectronics-Huazhong University of Science and TechnologyWuhan, China
- Key Laboratory of Biomedical Photonics of Ministry of Education, Department of Biomedical Engineering, Huazhong University of Science and TechnologyWuhan, China
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Johnson ME, Hanna SK, Montoro Bustos AR, Sims CM, Elliott LCC, Lingayat A, Johnston AC, Nikoobakht B, Elliott JT, Holbrook RD, Scott KCK, Murphy KE, Petersen EJ, Yu LL, Nelson BC. Separation, Sizing, and Quantitation of Engineered Nanoparticles in an Organism Model Using Inductively Coupled Plasma Mass Spectrometry and Image Analysis. ACS NANO 2017; 11:526-540. [PMID: 27983787 PMCID: PMC5459480 DOI: 10.1021/acsnano.6b06582] [Citation(s) in RCA: 25] [Impact Index Per Article: 3.6] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 05/03/2023]
Abstract
For environmental studies assessing uptake of orally ingested engineered nanoparticles (ENPs), a key step in ensuring accurate quantification of ingested ENPs is efficient separation of the organism from ENPs that are either nonspecifically adsorbed to the organism and/or suspended in the dispersion following exposure. Here, we measure the uptake of 30 and 60 nm gold nanoparticles (AuNPs) by the nematode, Caenorhabditis elegans, using a sucrose density gradient centrifugation protocol to remove noningested AuNPs. Both conventional inductively coupled plasma mass spectrometry (ICP-MS) and single particle (sp)ICP-MS are utilized to measure the total mass and size distribution, respectively, of ingested AuNPs. Scanning electron microscopy/energy-dispersive X-ray spectroscopy (SEM/EDS) imaging confirmed that traditional nematode washing procedures were ineffective at removing excess suspended and/or adsorbed AuNPs after exposure. Water rinsing procedures had AuNP removal efficiencies ranging from 57 to 97% and 22 to 83%, while the sucrose density gradient procedure had removal efficiencies of 100 and 93 to 98%, respectively, for the 30 and 60 nm AuNP exposure conditions. Quantification of total Au uptake was performed following acidic digestion of nonexposed and Au-exposed nematodes, whereas an alkaline digestion procedure was optimized for the liberation of ingested AuNPs for spICP-MS characterization. Size distributions and particle number concentrations were determined for AuNPs ingested by nematodes with corresponding confirmation of nematode uptake via high-pressure freezing/freeze substitution resin preparation and large-area SEM imaging. Methods for the separation and in vivo quantification of ENPs in multicellular organisms will facilitate robust studies of ENP uptake, biotransformation, and hazard assessment in the environment.
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Affiliation(s)
- Monique E Johnson
- Chemical Science Division, ‡Biosystems and Biomaterials Division, and §Materials Measurement Science Division, Material Measurement Laboratory, National Institute of Standards and Technology , Gaithersburg, Maryland 20899, United States
| | - Shannon K Hanna
- Chemical Science Division, ‡Biosystems and Biomaterials Division, and §Materials Measurement Science Division, Material Measurement Laboratory, National Institute of Standards and Technology , Gaithersburg, Maryland 20899, United States
| | - Antonio R Montoro Bustos
- Chemical Science Division, ‡Biosystems and Biomaterials Division, and §Materials Measurement Science Division, Material Measurement Laboratory, National Institute of Standards and Technology , Gaithersburg, Maryland 20899, United States
| | - Christopher M Sims
- Chemical Science Division, ‡Biosystems and Biomaterials Division, and §Materials Measurement Science Division, Material Measurement Laboratory, National Institute of Standards and Technology , Gaithersburg, Maryland 20899, United States
| | - Lindsay C C Elliott
- Chemical Science Division, ‡Biosystems and Biomaterials Division, and §Materials Measurement Science Division, Material Measurement Laboratory, National Institute of Standards and Technology , Gaithersburg, Maryland 20899, United States
| | - Akshay Lingayat
- Chemical Science Division, ‡Biosystems and Biomaterials Division, and §Materials Measurement Science Division, Material Measurement Laboratory, National Institute of Standards and Technology , Gaithersburg, Maryland 20899, United States
| | - Adrian C Johnston
- Chemical Science Division, ‡Biosystems and Biomaterials Division, and §Materials Measurement Science Division, Material Measurement Laboratory, National Institute of Standards and Technology , Gaithersburg, Maryland 20899, United States
| | - Babak Nikoobakht
- Chemical Science Division, ‡Biosystems and Biomaterials Division, and §Materials Measurement Science Division, Material Measurement Laboratory, National Institute of Standards and Technology , Gaithersburg, Maryland 20899, United States
| | - John T Elliott
- Chemical Science Division, ‡Biosystems and Biomaterials Division, and §Materials Measurement Science Division, Material Measurement Laboratory, National Institute of Standards and Technology , Gaithersburg, Maryland 20899, United States
| | - R David Holbrook
- Chemical Science Division, ‡Biosystems and Biomaterials Division, and §Materials Measurement Science Division, Material Measurement Laboratory, National Institute of Standards and Technology , Gaithersburg, Maryland 20899, United States
| | - Keana C K Scott
- Chemical Science Division, ‡Biosystems and Biomaterials Division, and §Materials Measurement Science Division, Material Measurement Laboratory, National Institute of Standards and Technology , Gaithersburg, Maryland 20899, United States
| | - Karen E Murphy
- Chemical Science Division, ‡Biosystems and Biomaterials Division, and §Materials Measurement Science Division, Material Measurement Laboratory, National Institute of Standards and Technology , Gaithersburg, Maryland 20899, United States
| | - Elijah J Petersen
- Chemical Science Division, ‡Biosystems and Biomaterials Division, and §Materials Measurement Science Division, Material Measurement Laboratory, National Institute of Standards and Technology , Gaithersburg, Maryland 20899, United States
| | - Lee L Yu
- Chemical Science Division, ‡Biosystems and Biomaterials Division, and §Materials Measurement Science Division, Material Measurement Laboratory, National Institute of Standards and Technology , Gaithersburg, Maryland 20899, United States
| | - Bryant C Nelson
- Chemical Science Division, ‡Biosystems and Biomaterials Division, and §Materials Measurement Science Division, Material Measurement Laboratory, National Institute of Standards and Technology , Gaithersburg, Maryland 20899, United States
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48
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Claeys M, Yushin VV, Leunissen JL, Claeys J, Bert W. Self-Pressurised Rapid Freezing (SPRF): an easy-to-use and low-cost alternative cryo-fixation method for nematodes. NEMATOLOGY 2017. [DOI: 10.1163/15685411-00003093] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/19/2022]
Abstract
Self-Pressurised Rapid Freezing (SPRF), an easy-to-use and low-cost alternative cryo-fixation method, was evaluated based on a comparative analysis of the ultrastructure of spermatozoa of the nematodes Acrobeles complexus and Caenorhabditis elegans. Sealed copper tubes, packed with active nematodes in water, were plunged into nitrogen slush, a semi-solid form of nitrogen. The water inside the capillary copper tube expands upon cooling due to the formation of hexagonal ice, thereby generating high pressure intrinsically for cryo-fixation of the sample. For sperm cells cryo-fixed by SPRF, the preservation of the ultrastructure was comparable to that achieved with high pressure freezing. This was evidenced by the clear details in mitochondria, membranous organelles and cytoskeleton in the pseudopod. It was demonstrated that SPRF fixation did not destroy antigenicity, based on the results of the immunolocalisation of the major sperm protein in both species. In conclusion, SPRF is a low-cost alternative cryo-fixation method for nematodes.
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Affiliation(s)
- Myriam Claeys
- Nematology Research Unit, Department of Biology, Ghent University, Belgium
| | - Vladimir V. Yushin
- National Scientific Center of Marine Biology, Far Eastern Branch, Russian Academy of Sciences, Vladivostok 690041, Russia
- Far Eastern Federal University, Vladivostok 690950, Russia
| | | | - Jef Claeys
- Nematology Research Unit, Department of Biology, Ghent University, Belgium
| | - Wim Bert
- Nematology Research Unit, Department of Biology, Ghent University, Belgium
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Shamir ER, Coutinho K, Georgess D, Auer M, Ewald AJ. Twist1-positive epithelial cells retain adhesive and proliferative capacity throughout dissemination. Biol Open 2016; 5:1216-28. [PMID: 27402962 PMCID: PMC5051642 DOI: 10.1242/bio.019703] [Citation(s) in RCA: 10] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/17/2023] Open
Abstract
Dissemination is the process by which cells detach and migrate away from a multicellular tissue. The epithelial-to-mesenchymal transition (EMT) conceptualizes dissemination in a stepwise fashion, with downregulation of E-cadherin leading to loss of intercellular junctions, induction of motility, and then escape from the epithelium. This gain of migratory activity is proposed to be mutually exclusive with proliferation. We previously developed a dissemination assay based on inducible expression of the transcription factor Twist1 and here utilize it to characterize the timing and dynamics of intercellular adhesion, proliferation and migration during dissemination. Surprisingly, Twist1(+) epithelium displayed extensive intercellular junctions, and Twist1(-) luminal epithelial cells could still adhere to disseminating Twist1(+) cells. Although proteolysis and proliferation were both observed throughout dissemination, neither was absolutely required. Finally, Twist1(+) cells exhibited a hybrid migration mode; their morphology and nuclear deformation were characteristic of amoeboid cells, whereas their dynamic protrusive activity, pericellular proteolysis and migration speeds were more typical of mesenchymal cells. Our data reveal that epithelial cells can disseminate while retaining competence to adhere and proliferate.
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Affiliation(s)
- Eliah R Shamir
- Departments of Cell Biology and Oncology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, 855 N. Wolfe St, Baltimore, MD 21205, USA
| | - Kester Coutinho
- Departments of Cell Biology and Oncology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, 855 N. Wolfe St, Baltimore, MD 21205, USA Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, 1 Cyclotron Road, MS Donner, Berkeley, CA 94720, USA
| | - Dan Georgess
- Departments of Cell Biology and Oncology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, 855 N. Wolfe St, Baltimore, MD 21205, USA
| | - Manfred Auer
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, 1 Cyclotron Road, MS Donner, Berkeley, CA 94720, USA
| | - Andrew J Ewald
- Departments of Cell Biology and Oncology, Center for Cell Dynamics, Johns Hopkins University School of Medicine, 855 N. Wolfe St, Baltimore, MD 21205, USA
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50
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Wolff G, Hagen C, Grünewald K, Kaufmann R. Towards correlative super-resolution fluorescence and electron cryo-microscopy. Biol Cell 2016; 108:245-58. [PMID: 27225383 PMCID: PMC5524168 DOI: 10.1111/boc.201600008] [Citation(s) in RCA: 64] [Impact Index Per Article: 8.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/14/2016] [Revised: 05/20/2016] [Accepted: 05/23/2016] [Indexed: 12/31/2022]
Abstract
Correlative light and electron microscopy (CLEM) has become a powerful tool in life sciences. Particularly cryo-CLEM, the combination of fluorescence cryo-microscopy (cryo-FM) permitting for non-invasive specific multi-colour labelling, with electron cryo-microscopy (cryo-EM) providing the undisturbed structural context at a resolution down to the Ångstrom range, has enabled a broad range of new biological applications. Imaging rare structures or events in crowded environments, such as inside a cell, requires specific fluorescence-based information for guiding cryo-EM data acquisition and/or to verify the identity of the structure of interest. Furthermore, cryo-CLEM can provide information about the arrangement of specific proteins in the wider structural context of their native nano-environment. However, a major obstacle of cryo-CLEM currently hindering many biological applications is the large resolution gap between cryo-FM (typically in the range of ∼400 nm) and cryo-EM (single nanometre to the Ångstrom range). Very recently, first proof of concept experiments demonstrated the feasibility of super-resolution cryo-FM imaging and the correlation with cryo-EM. This opened the door towards super-resolution cryo-CLEM, and thus towards direct correlation of structural details from both imaging modalities.
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Affiliation(s)
- Georg Wolff
- Division of Structural Biology, Wellcome Trust Centre for Human Genetics, University of Oxford, Oxford, UK
| | - Christoph Hagen
- Division of Structural Biology, Wellcome Trust Centre for Human Genetics, University of Oxford, Oxford, UK
| | - Kay Grünewald
- Division of Structural Biology, Wellcome Trust Centre for Human Genetics, University of Oxford, Oxford, UK
| | - Rainer Kaufmann
- Division of Structural Biology, Wellcome Trust Centre for Human Genetics, University of Oxford, Oxford, UK.
- Department of Biochemistry, University of Oxford, Oxford, UK.
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