1
|
Tran MV, Khuntsariya D, Fetter RD, Ferguson JW, Wang JT, Long AF, Cote LE, Wellard SR, Vázquez-Martínez N, Sallee MD, Genova M, Magiera MM, Eskinazi S, Lee JD, Peel N, Janke C, Stearns T, Shen K, Lansky Z, Magescas J, Feldman JL. MAP9/MAPH-9 supports axonemal microtubule doublets and modulates motor movement. Dev Cell 2024; 59:199-210.e11. [PMID: 38159567 PMCID: PMC11385174 DOI: 10.1016/j.devcel.2023.12.001] [Citation(s) in RCA: 3] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/09/2023] [Revised: 08/15/2023] [Accepted: 12/04/2023] [Indexed: 01/03/2024]
Abstract
Microtubule doublets (MTDs) comprise an incomplete microtubule (B-tubule) attached to the side of a complete cylindrical microtubule. These compound microtubules are conserved in cilia across the tree of life; however, the mechanisms by which MTDs form and are maintained in vivo remain poorly understood. Here, we identify microtubule-associated protein 9 (MAP9) as an MTD-associated protein. We demonstrate that C. elegans MAPH-9, a MAP9 homolog, is present during MTD assembly and localizes exclusively to MTDs, a preference that is in part mediated by tubulin polyglutamylation. We find that loss of MAPH-9 causes ultrastructural MTD defects, including shortened and/or squashed B-tubules with reduced numbers of protofilaments, dysregulated axonemal motor velocity, and perturbed cilia function. Because we find that the mammalian ortholog MAP9 localizes to axonemes in cultured mammalian cells and mouse tissues, we propose that MAP9/MAPH-9 plays a conserved role in regulating ciliary motors and supporting the structure of axonemal MTDs.
Collapse
Affiliation(s)
- Michael V Tran
- Department of Biology, Stanford University, Stanford, CA 94305, USA
| | - Daria Khuntsariya
- Institute of Biotechnology, Czech Academy of Sciences, BIOCEV, 25250 Vestec, Prague West, Czech Republic
| | - Richard D Fetter
- Howard Hughes Medical Institute, Department of Biology, Stanford University, Stanford, CA 94305, USA
| | - James W Ferguson
- Department of Biology, Stanford University, Stanford, CA 94305, USA
| | - Jennifer T Wang
- Department of Biology, Stanford University, Stanford, CA 94305, USA
| | - Alexandra F Long
- Department of Biology, Stanford University, Stanford, CA 94305, USA
| | - Lauren E Cote
- Department of Biology, Stanford University, Stanford, CA 94305, USA
| | | | | | - Maria D Sallee
- Department of Biology, Stanford University, Stanford, CA 94305, USA
| | - Mariya Genova
- Institut Curie, Université PSL, CNRS UMR3348, Orsay, France; Université Paris-Saclay, CNRS UMR3348, Orsay, France
| | - Maria M Magiera
- Institut Curie, Université PSL, CNRS UMR3348, Orsay, France; Université Paris-Saclay, CNRS UMR3348, Orsay, France
| | - Sani Eskinazi
- Department of Biology, Stanford University, Stanford, CA 94305, USA
| | | | - Nina Peel
- The College of New Jersey, Ewing, NJ 08628, USA
| | - Carsten Janke
- Institut Curie, Université PSL, CNRS UMR3348, Orsay, France; Université Paris-Saclay, CNRS UMR3348, Orsay, France
| | - Tim Stearns
- Department of Biology, Stanford University, Stanford, CA 94305, USA; Department of Genetics, Stanford University School of Medicine, Stanford, CA 94305, USA
| | - Kang Shen
- Howard Hughes Medical Institute, Department of Biology, Stanford University, Stanford, CA 94305, USA
| | - Zdenek Lansky
- Institute of Biotechnology, Czech Academy of Sciences, BIOCEV, 25250 Vestec, Prague West, Czech Republic
| | - Jérémy Magescas
- Department of Biology, Stanford University, Stanford, CA 94305, USA.
| | - Jessica L Feldman
- Department of Biology, Stanford University, Stanford, CA 94305, USA.
| |
Collapse
|
2
|
Land R, Fetter R, Liang X, Tzeng CP, Taylor CA, Shen K. Endoplasmic Reticulum Exit Sites scale with somato-dendritic size in neurons. Mol Biol Cell 2023; 34:ar106. [PMID: 37556208 PMCID: PMC10559313 DOI: 10.1091/mbc.e23-03-0090] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/13/2023] [Revised: 07/10/2023] [Accepted: 08/02/2023] [Indexed: 08/11/2023] Open
Abstract
Nervous systems exhibit dramatic diversity in cell morphology and size. How neurons regulate their biosynthetic and secretory machinery to support such diversity is not well understood. Endoplasmic reticulum exit sites (ERESs) are essential for maintaining secretory flux, and are required for normal dendrite development, but how neurons of different size regulate secretory capacity remains unknown. In Caenorhabditis elegans, we find that the ERES number is strongly correlated with the size of a neuron's dendritic arbor. The elaborately branched sensory neuron, PVD, has especially high ERES numbers. Asymmetric cell division provides PVD with a large initial cell size critical for rapid establishment of PVD's high ERES number before neurite outgrowth, and these ERESs are maintained throughout development. Maintenance of ERES number requires the cell fate transcription factor MEC-3, C. elegans TOR (ceTOR/let-363), and nutrient availability, with mec-3 and ceTOR/let-363 mutant PVDs both displaying reductions in ERES number, soma size, and dendrite size. Notably, mec-3 mutant animals exhibit reduced expression of a ceTOR/let-363 reporter in PVD, and starvation reduces ERES number and somato-dendritic size in a manner genetically redundant with ceTOR/let-363 perturbation. Our data suggest that both asymmetric cell division and nutrient sensing pathways regulate secretory capacities to support elaborate dendritic arbors.
Collapse
Affiliation(s)
- Ruben Land
- Department of Biology, Stanford University, Stanford, CA 94305
- Neurosciences IDP, Stanford University, Stanford, CA 94305
| | - Richard Fetter
- Howard Hughes Medical Institute, Stanford University, Stanford, CA 94305
| | - Xing Liang
- Department of Biology, Stanford University, Stanford, CA 94305
| | - Christopher P. Tzeng
- Department of Biology, Stanford University, Stanford, CA 94305
- Howard Hughes Medical Institute, Stanford University, Stanford, CA 94305
| | - Caitlin A. Taylor
- Department of Biology, Stanford University, Stanford, CA 94305
- Howard Hughes Medical Institute, Stanford University, Stanford, CA 94305
| | - Kang Shen
- Department of Biology, Stanford University, Stanford, CA 94305
- Howard Hughes Medical Institute, Stanford University, Stanford, CA 94305
| |
Collapse
|
3
|
Tran MV, Ferguson JW, Cote LE, Khuntsariya D, Fetter RD, Wang JT, Wellard SR, Sallee MD, Genova M, Eskinazi S, Magiera MM, Janke C, Stearns T, Lansky Z, Shen K, Magescas J, Feldman JL. MAP9/MAPH-9 supports axonemal microtubule doublets and modulates motor movement. BIORXIV : THE PREPRINT SERVER FOR BIOLOGY 2023:2023.02.23.529616. [PMID: 36865107 PMCID: PMC9980146 DOI: 10.1101/2023.02.23.529616] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 02/25/2023]
Abstract
Microtubule doublets (MTDs) are a well conserved compound microtubule structure found primarily in cilia. However, the mechanisms by which MTDs form and are maintained in vivo remain poorly understood. Here, we characterize microtubule-associated protein 9 (MAP9) as a novel MTD-associated protein. We demonstrate that C. elegans MAPH-9, a MAP9 homolog, is present during MTD assembly and localizes exclusively to MTDs, a preference that is in part mediated by tubulin polyglutamylation. Loss of MAPH-9 caused ultrastructural MTD defects, dysregulated axonemal motor velocity, and perturbed cilia function. As we found that the mammalian ortholog MAP9 localized to axonemes in cultured mammalian cells and mouse tissues, we propose that MAP9/MAPH-9 plays a conserved role in supporting the structure of axonemal MTDs and regulating ciliary motors.
Collapse
|
4
|
Bergner T, Zech F, Hirschenberger M, Stenger S, Sparrer KMJ, Kirchhoff F, Read C. Near-Native Visualization of SARS-CoV-2 Induced Membrane Remodeling and Virion Morphogenesis. Viruses 2022; 14:v14122786. [PMID: 36560790 PMCID: PMC9784144 DOI: 10.3390/v14122786] [Citation(s) in RCA: 2] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/10/2022] [Revised: 12/02/2022] [Accepted: 12/09/2022] [Indexed: 12/15/2022] Open
Abstract
Infection with the severe acute respiratory syndrome coronavirus-2 (SARS-CoV-2), the causative agent of the COVID-19 pandemic, leads to profound remodeling of cellular membranes, promoting viral replication and virion assembly. A full understanding of this drastic remodeling and the process of virion morphogenesis remains lacking. In this study, we applied room temperature transmission electron microscopy (TEM) and scanning transmission electron microscopy (STEM) tomography to visualize the SARS-CoV-2 replication factory in Vero cells, and present our results in comparison with published cryo-EM studies. We obtained cryo-EM-like clarity of the ultrastructure by employing high-pressure freezing, freeze substitution (HPF-FS) and embedding, allowing room temperature visualization of double-membrane vesicles (DMVs) in a near-native state. In addition, our data illustrate the consecutive stages of virion morphogenesis and reveal that SARS-CoV-2 ribonucleoprotein assembly and membrane curvature occur simultaneously. Finally, we show the tethering of virions to the plasma membrane in 3D, and that accumulations of virus particles lacking spike protein in large vesicles are most likely not a result of defective virion assembly at their membrane. In conclusion, this study puts forward a room-temperature EM technique providing near-native ultrastructural information about SARS-CoV-2 replication, adding to our understanding of the interaction of this pandemic virus with its host cell.
Collapse
Affiliation(s)
- Tim Bergner
- Central Facility for Electron Microscopy, Ulm University, 89081 Ulm, Germany
| | - Fabian Zech
- Institute of Molecular Virology, Ulm University Medical Center, 89081 Ulm, Germany
| | | | - Steffen Stenger
- Institute for Microbiology and Hygiene, Ulm University Medical Center, 89081 Ulm, Germany
| | | | - Frank Kirchhoff
- Institute of Molecular Virology, Ulm University Medical Center, 89081 Ulm, Germany
| | - Clarissa Read
- Central Facility for Electron Microscopy, Ulm University, 89081 Ulm, Germany
- Institute of Virology, Ulm University Medical Center, 89081 Ulm, Germany
- Correspondence:
| |
Collapse
|
5
|
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.
Collapse
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,
| |
Collapse
|
6
|
Heiligenstein X, Lucas MS. One for All, All for One: A Close Look at In-Resin Fluorescence Protocols for CLEM. Front Cell Dev Biol 2022; 10:866472. [PMID: 35846358 PMCID: PMC9280628 DOI: 10.3389/fcell.2022.866472] [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: 01/31/2022] [Accepted: 06/13/2022] [Indexed: 11/13/2022] Open
Abstract
Sample preparation is the novel bottleneck for high throughput correlative light and electron microscopy (CLEM). Protocols suitable for both imaging methods must therefore balance the requirements of each technique. For fluorescence light microscopy, a structure of interest can be targeted using: 1) staining, which is often structure or tissue specific rather than protein specific, 2) dye-coupled proteins or antibodies, or 3) genetically encoded fluorescent proteins. Each of these three methods has its own advantages. For ultrastructural investigation by electron microscopy (EM) resin embedding remains a significant sample preparation approach, as it stabilizes the sample such that it withstands the vacuum conditions of the EM, and enables long-term storage. Traditionally, samples are treated with heavy metal salts prior to resin embedding, in order to increase imaging contrast for EM. This is particularly important for volume EM (vEM) techniques. Yet, commonly used contrasting agents (e.g., osmium tetroxide, uranyl acetate) tend to impair fluorescence. The discovery that fluorescence can be preserved in resin-embedded specimens after mild heavy metal staining was a game changer for CLEM. These so-called in-resin fluorescence protocols present a significant leap forward for CLEM approaches towards high precision localization of a fluorescent signal in (volume) EM data. Integrated microscopy approaches, combining LM and EM detection into a single instrument certainly require such an “all in one” sample preparation. Preserving, or adding, dedicated fluorescence prior to resin embedding requires a compromise, which often comes at the expense of EM imaging contrast and membrane visibility. Especially vEM can be strongly hampered by a lack of heavy metal contrasting. This review critically reflects upon the fundamental aspects of resin embedding with regard to 1) specimen fixation and the physics and chemistry underlying the preservation of protein structure with respect to fluorescence and antigenicity, 2) optimization of EM contrast for transmission or scanning EM, and 3) the choice of embedding resin. On this basis, various existing workflows employing in-resin fluorescence are described, highlighting their common features, discussing advantages and disadvantages of the respective approach, and finally concluding with promising future developments for in-resin CLEM.
Collapse
Affiliation(s)
| | - Miriam S. Lucas
- Scientific Center for Light and Electron Microscopy (ScopeM), ETH Zurich, Zurich, Switzerland
- *Correspondence: Miriam S. Lucas,
| |
Collapse
|
7
|
Schauflinger M, Bergner T, Neusser G, Kranz C, Read C. Potassium permanganate is an excellent alternative to osmium tetroxide in freeze-substitution. Histochem Cell Biol 2022; 157:481-489. [PMID: 34984524 PMCID: PMC9001235 DOI: 10.1007/s00418-021-02070-0] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Accepted: 12/24/2021] [Indexed: 11/06/2022]
Abstract
High-pressure freezing followed by freeze-substitution is a valuable method for ultrastructural analyses of resin-embedded biological samples. The visualization of lipid membranes is one of the most critical aspects of any ultrastructural study and can be especially challenging in high-pressure frozen specimens. Historically, osmium tetroxide has been the preferred fixative and staining agent for lipid-containing structures in freeze-substitution solutions. However, osmium tetroxide is not only a rare and expensive material, but also volatile and toxic. Here, we introduce the use of a combination of potassium permanganate, uranyl acetate, and water in acetone as complementing reagents during the freeze-substitution process. This mix imparts an intense en bloc stain to cellular ultrastructure and membranes, which makes poststaining superfluous and is well suited for block-face imaging. Thus, potassium permanganate can effectively replace osmium tetroxide in the freeze-substitution solution without sacrificing the quality of ultrastructural preservation.
Collapse
Affiliation(s)
- Martin Schauflinger
- Institute of Virology, Philipps University Marburg, Hans-Meerwein-Straße 2, 35037, Marburg, Germany. .,Electron Microscopy Core Facility, University of Missouri, 1600 East Rollins Street, Columbia, MO, 65211, USA.
| | - Tim Bergner
- Central Facility for Electron Microscopy, Ulm University, Albert-Einstein-Allee 11, 89081, Ulm, Germany
| | - Gregor Neusser
- FIB Center UUlm, Institute of Analytical and Bioanalytical Chemistry, Ulm University, Albert-Einstein-Allee 11, 89081, Ulm, Germany
| | - Christine Kranz
- FIB Center UUlm, Institute of Analytical and Bioanalytical Chemistry, Ulm University, Albert-Einstein-Allee 11, 89081, Ulm, Germany
| | - Clarissa Read
- Central Facility for Electron Microscopy, Ulm University, Albert-Einstein-Allee 11, 89081, Ulm, Germany.,Institute of Virology, Ulm University Medical Center, Albert-Einstein-Allee 11, 89081, Ulm, Germany
| |
Collapse
|
8
|
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.
Collapse
Affiliation(s)
| | - Jindřiška Fišerová
- Department of Biology of the Cell Nucleus, Institute of Molecular Genetics AS CR, Prague, Czech Republic
| | | |
Collapse
|
9
|
Pipathsouk A, Brunetti RM, Town JP, Graziano BR, Breuer A, Pellett PA, Marchuk K, Tran NHT, Krummel MF, Stamou D, Weiner OD. The WAVE complex associates with sites of saddle membrane curvature. J Cell Biol 2021; 220:e202003086. [PMID: 34096975 PMCID: PMC8185649 DOI: 10.1083/jcb.202003086] [Citation(s) in RCA: 26] [Impact Index Per Article: 8.7] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/06/2020] [Revised: 04/13/2021] [Accepted: 05/18/2021] [Indexed: 12/30/2022] Open
Abstract
How local interactions of actin regulators yield large-scale organization of cell shape and movement is not well understood. Here we investigate how the WAVE complex organizes sheet-like lamellipodia. Using super-resolution microscopy, we find that the WAVE complex forms actin-independent 230-nm-wide rings that localize to regions of saddle membrane curvature. This pattern of enrichment could explain several emergent cell behaviors, such as expanding and self-straightening lamellipodia and the ability of endothelial cells to recognize and seal transcellular holes. The WAVE complex recruits IRSp53 to sites of saddle curvature but does not depend on IRSp53 for its own localization. Although the WAVE complex stimulates actin nucleation via the Arp2/3 complex, sheet-like protrusions are still observed in ARP2-null, but not WAVE complex-null, cells. Therefore, the WAVE complex has additional roles in cell morphogenesis beyond Arp2/3 complex activation. Our work defines organizing principles of the WAVE complex lamellipodial template and suggests how feedback between cell shape and actin regulators instructs cell morphogenesis.
Collapse
Affiliation(s)
- Anne Pipathsouk
- Cardiovascular Research Institute, University of California, San Francisco, San Francisco, CA
- Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, CA
| | - Rachel M. Brunetti
- Cardiovascular Research Institute, University of California, San Francisco, San Francisco, CA
- Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, CA
| | - Jason P. Town
- Cardiovascular Research Institute, University of California, San Francisco, San Francisco, CA
- Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, CA
| | - Brian R. Graziano
- Cardiovascular Research Institute, University of California, San Francisco, San Francisco, CA
| | - Artù Breuer
- Nano-Science Center and Department of Chemistry, University of Copenhagen, Copenhagen, Denmark
| | | | - Kyle Marchuk
- Department of Pathology and Biological Imaging Development CoLab, University of California, San Francisco, San Francisco, CA
| | - Ngoc-Han T. Tran
- Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, CA
| | - Matthew F. Krummel
- Department of Pathology and Biological Imaging Development CoLab, University of California, San Francisco, San Francisco, CA
| | - Dimitrios Stamou
- Nano-Science Center and Department of Chemistry, University of Copenhagen, Copenhagen, Denmark
| | - Orion D. Weiner
- Cardiovascular Research Institute, University of California, San Francisco, San Francisco, CA
- Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, CA
| |
Collapse
|
10
|
Müller A, Schmidt D, Xu CS, Pang S, D’Costa JV, Kretschmar S, Münster C, Kurth T, Jug F, Weigert M, Hess HF, Solimena M. 3D FIB-SEM reconstruction of microtubule-organelle interaction in whole primary mouse β cells. J Cell Biol 2021; 220:e202010039. [PMID: 33326005 PMCID: PMC7748794 DOI: 10.1083/jcb.202010039] [Citation(s) in RCA: 52] [Impact Index Per Article: 17.3] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/07/2020] [Revised: 11/14/2020] [Accepted: 11/18/2020] [Indexed: 11/22/2022] Open
Abstract
Microtubules play a major role in intracellular trafficking of vesicles in endocrine cells. Detailed knowledge of microtubule organization and their relation to other cell constituents is crucial for understanding cell function. However, their role in insulin transport and secretion is under debate. Here, we use FIB-SEM to image islet β cells in their entirety with unprecedented resolution. We reconstruct mitochondria, Golgi apparati, centrioles, insulin secretory granules, and microtubules of seven β cells, and generate a comprehensive spatial map of microtubule-organelle interactions. We find that microtubules form nonradial networks that are predominantly not connected to either centrioles or endomembranes. Microtubule number and length, but not microtubule polymer density, vary with glucose stimulation. Furthermore, insulin secretory granules are enriched near the plasma membrane, where they associate with microtubules. In summary, we provide the first 3D reconstructions of complete microtubule networks in primary mammalian cells together with evidence regarding their importance for insulin secretory granule positioning and thus their supportive role in insulin secretion.
Collapse
Affiliation(s)
- Andreas Müller
- Molecular Diabetology, University Hospital and Faculty of Medicine, Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany
- Paul Langerhans Institute Dresden of the Helmholtz Center Munich at the University Hospital and Faculty of Medicine, Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany
- German Center for Diabetes Research (DZD e.V.), Neuherberg, Germany
| | - Deborah Schmidt
- Center for Systems Biology Dresden, Dresden, Germany
- Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany
| | - C. Shan Xu
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA
| | - Song Pang
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA
| | - Joyson Verner D’Costa
- Molecular Diabetology, University Hospital and Faculty of Medicine, Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany
- Paul Langerhans Institute Dresden of the Helmholtz Center Munich at the University Hospital and Faculty of Medicine, Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany
- German Center for Diabetes Research (DZD e.V.), Neuherberg, Germany
| | - Susanne Kretschmar
- Center for Molecular and Cellular Bioengineering, Technology Platform, Technische Universität Dresden, Dresden, Germany
| | - Carla Münster
- Molecular Diabetology, University Hospital and Faculty of Medicine, Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany
- Paul Langerhans Institute Dresden of the Helmholtz Center Munich at the University Hospital and Faculty of Medicine, Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany
- German Center for Diabetes Research (DZD e.V.), Neuherberg, Germany
| | - Thomas Kurth
- Center for Molecular and Cellular Bioengineering, Technology Platform, Technische Universität Dresden, Dresden, Germany
| | - Florian Jug
- Center for Systems Biology Dresden, Dresden, Germany
- Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany
- Fondazione Human Technopole, Milano, Italy
| | - Martin Weigert
- Institute of Bioengineering, School of Life Sciences, École Polytechnique Fédérale de Lausanne, Lausanne, Switzerland
| | - Harald F. Hess
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA
| | - Michele Solimena
- Molecular Diabetology, University Hospital and Faculty of Medicine, Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany
- Paul Langerhans Institute Dresden of the Helmholtz Center Munich at the University Hospital and Faculty of Medicine, Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany
- German Center for Diabetes Research (DZD e.V.), Neuherberg, Germany
- Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany
| |
Collapse
|
11
|
Kuznetsov KM, Kozlov MI, Aslandukov AN, Vashchenko AA, Medved'ko AV, Latipov EV, Goloveshkin AS, Tsymbarenko DM, Utochnikova VV. Eu(tta) 3DPPZ-based organic light-emitting diodes: spin-coating vs. vacuum-deposition. Dalton Trans 2021; 50:9685-9689. [PMID: 34231618 DOI: 10.1039/d1dt01316e] [Citation(s) in RCA: 7] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/21/2022]
Abstract
The effect of the emission layer deposition method on the characteristics of OLEDs was studied on the example of the europium mixed-ligand complex Eu(tta)3DPPZ (tta: 2-thenoyltrifluoroacetone, DPPZ: dipyrido[3,2-a:2'c,3'c-c]phenazine). The maximum brightnesses of both OLEDs almost coincided, though OLED based on the spin-coated layer operated at lower voltages. The reason for that was the higher density and smoothness of the solution-processed layer.
Collapse
Affiliation(s)
- Kirill M Kuznetsov
- M.V. Lomonosov Moscow State University, 1/3 Leninskie Gory, Moscow, Russia.
| | - Makarii I Kozlov
- M.V. Lomonosov Moscow State University, 1/3 Leninskie Gory, Moscow, Russia.
| | - Andrey N Aslandukov
- Laboratory of Crystallography, University of Bayreuth, 95440 Bayreuth, Germany
| | | | - Aleksei V Medved'ko
- N.D. Zelinsky Institute of Organic Chemistry, 47 Leninsky Prospect, Moscow, Russia
| | - Egor V Latipov
- M.V. Lomonosov Moscow State University, 1/3 Leninskie Gory, Moscow, Russia.
| | | | | | - Valentina V Utochnikova
- M.V. Lomonosov Moscow State University, 1/3 Leninskie Gory, Moscow, Russia. and EVOLED Ltd, 1A-24 Puškina iela, Riga LV-1050, Latvia
| |
Collapse
|
12
|
Read C, Walther P, von Einem J. Quantitative Electron Microscopy to Study HCMV Morphogenesis. Methods Mol Biol 2021; 2244:265-289. [PMID: 33555592 DOI: 10.1007/978-1-0716-1111-1_14] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/28/2022]
Abstract
The generation and release of mature virions from human cytomegalovirus (HCMV) infected cells is a multistep process, involving a profound reorganization of cellular structures and various stages of virus particle morphogenesis in different cellular compartments. Although the general steps of HCMV morphogenesis are known, it has become clear that the detailed molecular mechanisms are complex and dependent on various viral factors and cellular pathways. The lack of a full understanding of HCMV virion morphogenesis emphasizes the need of imaging techniques to visualize the different stages of virion assembly, such as electron microscopy. Here, we describe various electron microscopy techniques and the methodology of high-pressure freezing and freeze substitution for sample preparation to visualize HCMV morphogenesis. These methods are used in our laboratory in combination with a thorough quantification to characterize phenotypic alterations and to identify the function of viral and cellular proteins for the various morphogenesis stages.
Collapse
Affiliation(s)
- Clarissa Read
- Institute of Virology, Ulm University Medical Center, Ulm, Germany.,Central Facility for Electron Microscopy, Ulm University, Ulm, Germany
| | - Paul Walther
- Central Facility for Electron Microscopy, Ulm University, Ulm, Germany
| | - Jens von Einem
- Institute of Virology, Ulm University Medical Center, Ulm, Germany.
| |
Collapse
|
13
|
Javid A, Roudbari A, Yousefi N, Fard MA, Barkdoll B, Talebi SS, Nazemi S, Ghanbarian M, Ghadiri SK. Modeling of chromium (VI) removal from aqueous solution using modified green-Graphene: RSM-CCD approach, optimization, isotherm, and kinetic studies. JOURNAL OF ENVIRONMENTAL HEALTH SCIENCE & ENGINEERING 2020; 18:515-529. [PMID: 33312580 PMCID: PMC7721790 DOI: 10.1007/s40201-020-00479-8] [Citation(s) in RCA: 15] [Impact Index Per Article: 3.8] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 10/23/2019] [Accepted: 04/14/2020] [Indexed: 05/07/2023]
Abstract
BACKGROUND The aim of this study was to investigate the removal of Cr (VI) using Green-Graphene Nanosheets (GGN) synthesized from rice straw. METHODS Synthesis of the GGN was optimized using response surface methodology and central composite design (CCD). The effect of two independent variables including KOH-to-raw rice ash (KOH/RRA) ratio and temperature on the specific surface area of the GGN was determined. To have better removal of Cr (VI), GGN was modified using the grafting amine group method. In the Cr (VI) removal process, the effects of four independent variables including initial Cr (VI) concentration, adsorbent dosage, contact time, and initial solution pH were studied. RESULTS The results of this study showed that the optimum values of the KOH/RRA ratio and temperature for the preparation of GGN were 10.85 and 749.61 °C, respectively. The maximum amount of SSA obtained at optimum conditions for GGN was 551.14 ± 3.83 m 2 /g. The optimum conditions for Cr (VI) removal were 48.35 mg/L, 1.46 g/L, 44.30 min, and 6.87 for Cr (VI) concentration, adsorbent dosage, contact time, and pH, respectively. Based on variance analysis, the adsorbent dose was the most sensitive factor for Cr (VI) removal. Langmuir isotherm (R2 = 0.991) and Pseudo-second-order kinetic models (R2 = 0.999) were the best fit for the study results and the Q max was 138.89 mg/g. CONCLUSIONS It can be concluded that the predicted conditions from the GGN synthesis model and the optimum conditions from the Cr (VI) removal model both agreed with the experimental findings.
Collapse
Affiliation(s)
- Allahbakhsh Javid
- Department of Environmental Health Engineering, School of Public Health, Shahroud University of Medical Sciences, Shahroud, Iran
| | - Aliakbar Roudbari
- Department of Environmental Health Engineering, School of Public Health, Shahroud University of Medical Sciences, Shahroud, Iran
| | - Nader Yousefi
- Department of Environmental Health Engineering, School of Public Health, Tehran University of Medical Sciences, Tehran, Iran
| | - Mohammad Alizadeh Fard
- Department of Civil and Environmental Engineering, Michigan Technological University, Houghton, MI USA
| | - Brian Barkdoll
- Department of Civil and Environmental Engineering, Michigan Technological University, Houghton, MI USA
| | - Seyedeh Solmaz Talebi
- Department of Epidemiology, School of Medicine, Shahroud University of Medical Sciences, Shahroud, Iran
| | - Saeed Nazemi
- Department of Environmental Health Engineering, School of Public Health, Shahroud University of Medical Sciences, Shahroud, Iran
| | - Marjan Ghanbarian
- Department of Environmental Health Engineering, School of Public Health, Shahroud University of Medical Sciences, Shahroud, Iran
| | - Seid Kamal Ghadiri
- Department of Environmental Health Engineering, School of Public Health, Shahroud University of Medical Sciences, Shahroud, Iran
| |
Collapse
|
14
|
McDonald NA, Fetter RD, Shen K. Assembly of synaptic active zones requires phase separation of scaffold molecules. Nature 2020; 588:454-458. [PMID: 33208945 DOI: 10.1038/s41586-020-2942-0] [Citation(s) in RCA: 77] [Impact Index Per Article: 19.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/07/2020] [Accepted: 09/03/2020] [Indexed: 12/20/2022]
Abstract
The formation of synapses during neuronal development is essential for establishing neural circuits and a nervous system1. Every presynapse builds a core 'active zone' structure, where ion channels cluster and synaptic vesicles release their neurotransmitters2. Although the composition of active zones is well characterized2,3, it is unclear how active-zone proteins assemble together and recruit the machinery required for vesicle release during development. Here we find that the core active-zone scaffold proteins SYD-2 (also known as liprin-α) and ELKS-1 undergo phase separation during an early stage of synapse development, and later mature into a solid structure. We directly test the in vivo function of phase separation by using mutant SYD-2 and ELKS-1 proteins that specifically lack this activity. These mutant proteins remain enriched at synapses in Caenorhabditis elegans, but show defects in active-zone assembly and synapse function. The defects are rescued by introducing a phase-separation motif from an unrelated protein. In vitro, we reconstitute the SYD-2 and ELKS-1 liquid-phase scaffold, and find that it is competent to bind and incorporate downstream active-zone components. We find that the fluidity of SYD-2 and ELKS-1 condensates is essential for efficient mixing and incorporation of active-zone components. These data reveal that a developmental liquid phase of scaffold molecules is essential for the assembly of the synaptic active zone, before maturation into a stable final structure.
Collapse
Affiliation(s)
| | - Richard D Fetter
- Howard Hughes Medical Institute, Department of Biology, Stanford University, Stanford, CA, USA
| | - Kang Shen
- Department of Biology, Stanford University, Stanford, CA, USA. .,Howard Hughes Medical Institute, Department of Biology, Stanford University, Stanford, CA, USA.
| |
Collapse
|
15
|
Parvate A, Sengupta R, Williams EP, Xue Y, Chu YK, Stahelin RV, Jonsson CB. Cryofixation of Inactivated Hantavirus-Infected Cells as a Method for Obtaining High-Quality Ultrastructural Preservation for Electron Microscopic Studies. Front Cell Infect Microbiol 2020; 10:580339. [PMID: 33240823 PMCID: PMC7677528 DOI: 10.3389/fcimb.2020.580339] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/05/2020] [Accepted: 09/25/2020] [Indexed: 12/17/2022] Open
Abstract
Hantaviruses rewire the host cell and induce extensive membrane rearrangements for their replication and the morphogenesis of the virion. Transmission electron microscopy (TEM) is a powerful technique for imaging these pathological membrane changes especially when combined with large volume electron tomography. Excellent preservation of membrane structure can be obtained when chemical fixation is combined with cryofixation via high pressure freezing making the samples amenable to serial-section tomographic reconstruction. Taking advantage of this, we have optimized a hybrid method that employs aldehyde fixation, a step that is essential for virus inactivation, followed by high-pressure freezing for ultrastructural study of Hantaan (HTN) and Andes (AND) virus infected Vero E6 cells. HTNV and ANDV are two species of the Orthohantavirus, from the Old and New World, respectively, and the causative agents of hemorrhagic fever with renal syndrome and hantavirus pulmonary syndrome in humans. We applied the method for the qualitative assessment of the perturbation of the endomembrane system induced by HTNV and ANDV in infected vs. mock-infected cells. Screening of serial-sections revealed consistency of membrane preservation across large volumes indicating potential of these samples for tomographic studies. Images revealed large-scale perturbations of the endomembrane system following HTNV-infection that included the dilation of the rough endoplasmic reticulum and fragmentation of the Golgi apparatus. Infected cells exhibited a tendency to accumulate large numbers of vacuoles that were especially apparent in ANDV. In summary, our hybrid method provides a path for the study of BSL-3 pathogens using cutting edge 3D-imaging technologies.
Collapse
Affiliation(s)
- Amar Parvate
- Department of Biological Sciences, Purdue University, West Lafayette, IN, United States
| | - Ranjan Sengupta
- Department of Biological Sciences, Purdue University, West Lafayette, IN, United States
- Medicinal Chemistry and Molecular Pharmacology and the Purdue Institute for Inflammation, Immunology and Infectious Disease, Purdue University, West Lafayette, IN, United States
| | - Evan P. Williams
- Department of Microbiology, Immunology and Biochemistry, University of Tennessee Health Science Center, Memphis, TN, United States
| | - Yi Xue
- Department of Microbiology, Immunology and Biochemistry, University of Tennessee Health Science Center, Memphis, TN, United States
| | - Yong-Kyu Chu
- Center for Predictive Medicine, University of Louisville, Louisville, KY, United States
| | - Robert V. Stahelin
- Medicinal Chemistry and Molecular Pharmacology and the Purdue Institute for Inflammation, Immunology and Infectious Disease, Purdue University, West Lafayette, IN, United States
| | - Colleen B. Jonsson
- Department of Microbiology, Immunology and Biochemistry, University of Tennessee Health Science Center, Memphis, TN, United States
| |
Collapse
|
16
|
Liang X, Kokes M, Fetter RD, Sallee MD, Moore AW, Feldman JL, Shen K. Growth cone-localized microtubule organizing center establishes microtubule orientation in dendrites. eLife 2020; 9:e56547. [PMID: 32657271 PMCID: PMC7375809 DOI: 10.7554/elife.56547] [Citation(s) in RCA: 30] [Impact Index Per Article: 7.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/02/2020] [Accepted: 07/09/2020] [Indexed: 01/01/2023] Open
Abstract
A polarized arrangement of neuronal microtubule arrays is the foundation of membrane trafficking and subcellular compartmentalization. Conserved among both invertebrates and vertebrates, axons contain exclusively 'plus-end-out' microtubules while dendrites contain a high percentage of 'minus-end-out' microtubules, the origins of which have been a mystery. Here we show that in Caenorhabditis elegans the dendritic growth cone contains a non-centrosomal microtubule organizing center (MTOC), which generates minus-end-out microtubules along outgrowing dendrites and plus-end-out microtubules in the growth cone. RAB-11-positive endosomes accumulate in this region and co-migrate with the microtubule nucleation complex γ-TuRC. The MTOC tracks the extending growth cone by kinesin-1/UNC-116-mediated endosome movements on distal plus-end-out microtubules and dynein clusters this advancing MTOC. Critically, perturbation of the function or localization of the MTOC causes reversed microtubule polarity in dendrites. These findings unveil the endosome-localized dendritic MTOC as a critical organelle for establishing axon-dendrite polarity.
Collapse
Affiliation(s)
- Xing Liang
- Department of Biology, Stanford UniversityStanfordUnited States
- Howard Hughes Medical Institute, Stanford UniversityStanfordUnited States
| | - Marcela Kokes
- Department of Biology, Stanford UniversityStanfordUnited States
- Howard Hughes Medical Institute, Stanford UniversityStanfordUnited States
| | - Richard D Fetter
- Howard Hughes Medical Institute, Stanford UniversityStanfordUnited States
| | | | | | | | - Kang Shen
- Department of Biology, Stanford UniversityStanfordUnited States
- Howard Hughes Medical Institute, Stanford UniversityStanfordUnited States
| |
Collapse
|
17
|
Bauer A, Frascaroli G, Walther P. Megapinosome: Morphological description of a novel organelle. J Struct Biol 2020; 210:107505. [DOI: 10.1016/j.jsb.2020.107505] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/20/2020] [Revised: 03/19/2020] [Accepted: 03/30/2020] [Indexed: 01/08/2023]
|
18
|
Thornton SM, Samararatne VD, Skeate JG, Buser C, Lühen KP, Taylor JR, Da Silva DM, Kast WM. The Essential Role of anxA2 in Langerhans Cell Birbeck Granules Formation. Cells 2020; 9:cells9040974. [PMID: 32326440 PMCID: PMC7227008 DOI: 10.3390/cells9040974] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/05/2020] [Revised: 04/01/2020] [Accepted: 04/12/2020] [Indexed: 12/15/2022] Open
Abstract
Langerhans cells (LC) are the resident antigen presenting cells of the mucosal epithelium and play an essential role in initiating immune responses. LC are the only cells in the body to contain Birbeck granules (BG), which are unique cytoplasmic organelles comprised of c-type lectin langerin. Studies of BG have historically focused on morphological characterizations, but BG have also been implicated in viral antigen processing which suggests that they can serve a function in antiviral immunity. This study focused on investigating proteins that could be involved in BG formation to further characterize their structure using transmission electron microscopy (TEM). Here, we report a critical role for the protein annexin A2 (anxA2) in the proper formation of BG structures. When anxA2 expression is downregulated, langerin expression decreases, cytoplasmic BG are nearly ablated, and the presence of malformed BG-like structures increases. Furthermore, in the absence of anxA2, we found langerin was no longer localized to BG or BG-like structures. Taken together, these results indicate an essential role for anxA2 in facilitating the proper formation of BG.
Collapse
Affiliation(s)
- Shantae M. Thornton
- Department of Molecular Microbiology and Immunology, University of Southern California, Los Angeles, CA 90033, USA; (S.M.T.); (V.D.S.); (J.G.S.); (J.R.T.)
| | - Varsha D. Samararatne
- Department of Molecular Microbiology and Immunology, University of Southern California, Los Angeles, CA 90033, USA; (S.M.T.); (V.D.S.); (J.G.S.); (J.R.T.)
| | - Joseph G. Skeate
- Department of Molecular Microbiology and Immunology, University of Southern California, Los Angeles, CA 90033, USA; (S.M.T.); (V.D.S.); (J.G.S.); (J.R.T.)
| | | | - Kim P. Lühen
- Norris Comprehensive Cancer Center, University of Southern California, Los Angeles, CA 90033, USA; (K.P.L.); (D.M.D.S.)
| | - Julia R. Taylor
- Department of Molecular Microbiology and Immunology, University of Southern California, Los Angeles, CA 90033, USA; (S.M.T.); (V.D.S.); (J.G.S.); (J.R.T.)
| | - Diane M. Da Silva
- Norris Comprehensive Cancer Center, University of Southern California, Los Angeles, CA 90033, USA; (K.P.L.); (D.M.D.S.)
- Department of Obstetrics & Gynecology, University of Southern California, Los Angeles, CA 90033, USA
| | - W. Martin Kast
- Department of Molecular Microbiology and Immunology, University of Southern California, Los Angeles, CA 90033, USA; (S.M.T.); (V.D.S.); (J.G.S.); (J.R.T.)
- Norris Comprehensive Cancer Center, University of Southern California, Los Angeles, CA 90033, USA; (K.P.L.); (D.M.D.S.)
- Department of Obstetrics & Gynecology, University of Southern California, Los Angeles, CA 90033, USA
- Correspondence: ; Tel.: +1-323-442-3870
| |
Collapse
|
19
|
VELAMOOR S, RICHENA M, MITCHELL A, LEQUEUX S, BOSTINA M, HARLAND D. High‐pressure freezing followed by freeze substitution of a complex and variable density miniorgan: the wool follicle. J Microsc 2020; 278:18-28. [DOI: 10.1111/jmi.12875] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/17/2019] [Revised: 01/16/2020] [Accepted: 02/06/2020] [Indexed: 01/15/2023]
Affiliation(s)
- S. VELAMOOR
- Lincoln Research Centre, Food & Bio‐Based ProductsAgresearch Limited Lincoln New Zealand
- Department of Immunology and MicrobiologyUniversity of Otago Dunedin New Zealand
| | - M. RICHENA
- Lincoln Research Centre, Food & Bio‐Based ProductsAgresearch Limited Lincoln New Zealand
| | - A. MITCHELL
- Otago Micro and Nano Imaging UnitUniversity of Otago Dunedin New Zealand
| | - S. LEQUEUX
- Otago Micro and Nano Imaging UnitUniversity of Otago Dunedin New Zealand
| | - M. BOSTINA
- Department of Immunology and MicrobiologyUniversity of Otago Dunedin New Zealand
- Otago Micro and Nano Imaging UnitUniversity of Otago Dunedin New Zealand
| | - D. HARLAND
- Lincoln Research Centre, Food & Bio‐Based ProductsAgresearch Limited Lincoln New Zealand
| |
Collapse
|
20
|
Triffo WJ, Palsdottir H, Song J, Morgan DG, McDonald KL, Auer M, Raphael RM. 3D Ultrastructure of the Cochlear Outer Hair Cell Lateral Wall Revealed By Electron Tomography. Front Cell Neurosci 2019; 13:560. [PMID: 31920560 PMCID: PMC6933316 DOI: 10.3389/fncel.2019.00560] [Citation(s) in RCA: 9] [Impact Index Per Article: 1.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/08/2019] [Accepted: 12/04/2019] [Indexed: 11/17/2022] Open
Abstract
Outer Hair Cells (OHCs) in the mammalian cochlea display a unique type of voltage-induced mechanical movement termed electromotility, which amplifies auditory signals and contributes to the sensitivity and frequency selectivity of mammalian hearing. Electromotility occurs in the OHC lateral wall, but it is not fully understood how the supramolecular architecture of the lateral wall enables this unique form of cellular motility. Employing electron tomography of high-pressure frozen and freeze-substituted OHCs, we visualized the 3D structure and organization of the membrane and cytoskeletal components of the OHC lateral wall. The subsurface cisterna (SSC) is a highly prominent feature, and we report that the SSC membranes and lumen possess hexagonally ordered arrays of particles. We also find the SSC is tightly connected to adjacent actin filaments by short filamentous protein connections. Pillar proteins that join the plasma membrane to the cytoskeleton appear as variable structures considerably thinner than actin filaments and significantly more flexible than actin-SSC links. The structurally rich organization and rigidity of the SSC coupled with apparently weaker mechanical connections between the plasma membrane (PM) and cytoskeleton reveal that the membrane-cytoskeletal architecture of the OHC lateral wall is more complex than previously appreciated. These observations are important for our understanding of OHC mechanics and need to be considered in computational models of OHC electromotility that incorporate subcellular features.
Collapse
Affiliation(s)
- William Jeffrey Triffo
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, United States.,Department of Bioengineering, George R. Brown School of Engineering, Rice University, Houston, TX, United States.,Department of Radiology, Geisinger, Danville, PA, United States
| | - Hildur Palsdottir
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, United States
| | - Junha Song
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, United States
| | - David Gene Morgan
- Interdisciplinary Center for Electron Microscopy, University of California, Davis, Davis, CA, United States
| | - Kent L McDonald
- Electron Microscope Laboratory, University of California, Berkeley, Berkeley, CA, United States
| | - Manfred Auer
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, United States
| | - Robert M Raphael
- Department of Bioengineering, George R. Brown School of Engineering, Rice University, Houston, TX, United States
| |
Collapse
|
21
|
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.
Collapse
|
22
|
Müller MT, Schempp R, Lutz A, Felder T, Felder E, Miklavc P. Interaction of microtubules and actin during the post-fusion phase of exocytosis. Sci Rep 2019; 9:11973. [PMID: 31427591 PMCID: PMC6700138 DOI: 10.1038/s41598-019-47741-0] [Citation(s) in RCA: 8] [Impact Index Per Article: 1.6] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/11/2019] [Accepted: 07/09/2019] [Indexed: 01/24/2023] Open
Abstract
Exocytosis is the intracellular trafficking step where a secretory vesicle fuses with the plasma membrane to release vesicle content. Actin and microtubules both play a role in exocytosis; however, their interplay is not understood. Here we study the interaction of actin and microtubules during exocytosis in lung alveolar type II (ATII) cells that secrete surfactant from large secretory vesicles. Surfactant extrusion is facilitated by an actin coat that forms on the vesicle shortly after fusion pore opening. Actin coat compression allows hydrophobic surfactant to be released from the vesicle. We show that microtubules are localized close to actin coats and stay close to the coats during their compression. Inhibition of microtubule polymerization by colchicine and nocodazole affected the kinetics of actin coat formation and the extent of actin polymerisation on fused vesicles. In addition, microtubule and actin cross-linking protein IQGAP1 localized to fused secretory vesicles and IQGAP1 silencing influenced actin polymerisation after vesicle fusion. This study demonstrates that microtubules can influence actin coat formation and actin polymerization on secretory vesicles during exocytosis.
Collapse
Affiliation(s)
- M Tabitha Müller
- Institute of General Physiology, Ulm University, Albert-Einstein Allee 11, 89081, Ulm, Germany
| | - Rebekka Schempp
- Institute of General Physiology, Ulm University, Albert-Einstein Allee 11, 89081, Ulm, Germany
| | - Anngrit Lutz
- Institute of General Physiology, Ulm University, Albert-Einstein Allee 11, 89081, Ulm, Germany
| | - Tatiana Felder
- Institute of General Physiology, Ulm University, Albert-Einstein Allee 11, 89081, Ulm, Germany
| | - Edward Felder
- Institute of General Physiology, Ulm University, Albert-Einstein Allee 11, 89081, Ulm, Germany
| | - Pika Miklavc
- School of Environment and Life Sciences, University of Salford, The Crescent, M54WT, Salford, United Kingdom.
| |
Collapse
|
23
|
Ding K, Han Y, Seid TW, Buser C, Karigo T, Zhang S, Dickman DK, Anderson DJ. Imaging neuropeptide release at synapses with a genetically engineered reporter. eLife 2019; 8:e46421. [PMID: 31241464 PMCID: PMC6609332 DOI: 10.7554/elife.46421] [Citation(s) in RCA: 19] [Impact Index Per Article: 3.8] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/27/2019] [Accepted: 06/25/2019] [Indexed: 12/26/2022] Open
Abstract
Research on neuropeptide function has advanced rapidly, yet there is still no spatio-temporally resolved method to measure the release of neuropeptides in vivo. Here we introduce Neuropeptide Release Reporters (NPRRs): novel genetically-encoded sensors with high temporal resolution and genetic specificity. Using the Drosophila larval neuromuscular junction (NMJ) as a model, we provide evidence that NPRRs recapitulate the trafficking and packaging of native neuropeptides, and report stimulation-evoked neuropeptide release events as real-time changes in fluorescence intensity, with sub-second temporal resolution.
Collapse
Affiliation(s)
- Keke Ding
- Division of Biology and Biological EngineeringCalifornia Institute of TechnologyPasadenaUnited States
| | - Yifu Han
- Department of NeurobiologyUniversity of Southern CaliforniaLos AngelesUnited States
- Neuroscience Graduate ProgramUniversity of Southern CaliforniaLos AngelesUnited States
| | - Taylor W Seid
- Division of Biology and Biological EngineeringCalifornia Institute of TechnologyPasadenaUnited States
| | | | - Tomomi Karigo
- Division of Biology and Biological EngineeringCalifornia Institute of TechnologyPasadenaUnited States
| | - Shishuo Zhang
- Division of Biology and Biological EngineeringCalifornia Institute of TechnologyPasadenaUnited States
| | - Dion K Dickman
- Department of NeurobiologyUniversity of Southern CaliforniaLos AngelesUnited States
| | - David J Anderson
- Division of Biology and Biological EngineeringCalifornia Institute of TechnologyPasadenaUnited States
- Howard Hughes Medical Institute, California Institute of TechnologyPasadenaUnited States
- Tianqiao and Chrissy Chen Institute for Neuroscience, California Institute of TechnologyPasadenaUnited States
| |
Collapse
|
24
|
Chang CL, Weigel AV, Ioannou MS, Pasolli HA, Xu CS, Peale DR, Shtengel G, Freeman M, Hess HF, Blackstone C, Lippincott-Schwartz J. Spastin tethers lipid droplets to peroxisomes and directs fatty acid trafficking through ESCRT-III. J Cell Biol 2019; 218:2583-2599. [PMID: 31227594 PMCID: PMC6683741 DOI: 10.1083/jcb.201902061] [Citation(s) in RCA: 130] [Impact Index Per Article: 26.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/11/2019] [Revised: 04/29/2019] [Accepted: 05/28/2019] [Indexed: 12/22/2022] Open
Abstract
Lipid droplets (LDs) are neutral lipid storage organelles that transfer lipids to various organelles including peroxisomes. Here, we show that the hereditary spastic paraplegia protein M1 Spastin, a membrane-bound AAA ATPase found on LDs, coordinates fatty acid (FA) trafficking from LDs to peroxisomes through two interrelated mechanisms. First, M1 Spastin forms a tethering complex with peroxisomal ABCD1 to promote LD-peroxisome contact formation. Second, M1 Spastin recruits the membrane-shaping ESCRT-III proteins IST1 and CHMP1B to LDs via its MIT domain to facilitate LD-to-peroxisome FA trafficking, possibly through IST1- and CHMP1B-dependent modifications in LD membrane morphology. Furthermore, LD-to-peroxisome FA trafficking mediated by M1 Spastin is required to relieve LDs of lipid peroxidation. M1 Spastin's dual roles in tethering LDs to peroxisomes and in recruiting ESCRT-III components to LD-peroxisome contact sites for FA trafficking may underlie the pathogenesis of diseases associated with defective FA metabolism in LDs and peroxisomes.
Collapse
Affiliation(s)
- Chi-Lun Chang
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA
| | - Aubrey V Weigel
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA
| | - Maria S Ioannou
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA
| | - H Amalia Pasolli
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA
| | - C Shan Xu
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA
| | - David R Peale
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA
| | - Gleb Shtengel
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA
| | - Melanie Freeman
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA
| | - Harald F Hess
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA
| | - Craig Blackstone
- Neurogenetics Branch, National Institute of Neurological Disorders and Stroke, National Institutes of Health, Bethesda, MD
| | | |
Collapse
|
25
|
Brockmann SJ, Freischmidt A, Oeckl P, Müller K, Ponna SK, Helferich AM, Paone C, Reinders J, Kojer K, Orth M, Jokela M, Auranen M, Udd B, Hermann A, Danzer KM, Lichtner P, Walther P, Ludolph AC, Andersen PM, Otto M, Kursula P, Just S, Weishaupt JH. CHCHD10 mutations p.R15L and p.G66V cause motoneuron disease by haploinsufficiency. Hum Mol Genet 2019; 27:706-715. [PMID: 29315381 DOI: 10.1093/hmg/ddx436] [Citation(s) in RCA: 24] [Impact Index Per Article: 4.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/17/2017] [Accepted: 12/18/2017] [Indexed: 12/21/2022] Open
Abstract
Mutations in the mitochondrially located protein CHCHD10 cause motoneuron disease by an unknown mechanism. In this study, we investigate the mutations p.R15L and p.G66V in comparison to wild-type CHCHD10 and the non-pathogenic variant p.P34S in vitro, in patient cells as well as in the vertebrate in vivo model zebrafish. We demonstrate a reduction of CHCHD10 protein levels in p.R15L and p.G66V mutant patient cells to approximately 50%. Quantitative real-time PCR revealed that expression of CHCHD10 p.R15L, but not of CHCHD10 p.G66V, is already abrogated at the mRNA level. Altered secondary structure and rapid protein degradation are observed with regard to the CHCHD10 p.G66V mutant. In contrast, no significant differences in expression, degradation rate or secondary structure of non-pathogenic CHCHD10 p.P34S are detected when compared with wild-type protein. Knockdown of CHCHD10 expression in zebrafish to about 50% causes motoneuron pathology, abnormal myofibrillar structure and motility deficits in vivo. Thus, our data show that the CHCHD10 mutations p.R15L and p.G66V cause motoneuron disease primarily based on haploinsufficiency of CHCHD10.
Collapse
Affiliation(s)
| | | | - Patrick Oeckl
- Department of Neurology, Ulm University, 89081 Ulm, Germany
| | - Kathrin Müller
- Department of Neurology, Ulm University, 89081 Ulm, Germany
| | - Srinivas K Ponna
- Faculty of Biochemistry and Molecular Medicine, University of Oulu, 90014 Oulu, Finland
| | | | - Christoph Paone
- Molecular Cardiology, Department of Internal Medicine II, Ulm University Medical Center, 89081 Ulm, Germany
| | - Jörg Reinders
- Institute for Functional Genomics, University Regensburg, 93053 Regensburg, Germany
| | - Kerstin Kojer
- Department of Neurology, Ulm University, 89081 Ulm, Germany
| | - Michael Orth
- Department of Neurology, Ulm University, 89081 Ulm, Germany
| | - Manu Jokela
- Neuromuscular Research Center, Tampere University and University Hospital, 33014 Tampere, Finland
| | - Mari Auranen
- Neurological Department, Helsinki University Hospital, 00029 Helsinki, Finland
| | - Bjarne Udd
- Neuromuscular Research Center, Tampere University and University Hospital, 33014 Tampere, Finland
| | - Andreas Hermann
- Department of Neurology, Technische Universität Dresden, 01307 Dresden, Germany.,German Center for Neurodegenerative Diseases, Dresden Research Site, 01307 Dresden, Germany.,Center for Regenerative Therapies Dresden (CRTD), Technische Universität Dresden, 01307 Dresden, Germany
| | - Karin M Danzer
- Department of Neurology, Ulm University, 89081 Ulm, Germany
| | - Peter Lichtner
- Institute of Human Genetics, Helmholtz Zentrum München, 85764 Neuherberg, Germany
| | - Paul Walther
- Zentrale Einrichtung Elektronenmikroskopie, Universitaet Ulm, 89081 Ulm, Germany
| | | | - Peter M Andersen
- Department of Pharmacology and Clinical Neuroscience, Umeå University, 90187 Umeå, Sweden
| | - Markus Otto
- Department of Neurology, Ulm University, 89081 Ulm, Germany
| | - Petri Kursula
- Faculty of Biochemistry and Molecular Medicine, University of Oulu, 90014 Oulu, Finland.,Department of Biomedicine, University of Bergen, 5020 Bergen, Norway
| | - Steffen Just
- Molecular Cardiology, Department of Internal Medicine II, Ulm University Medical Center, 89081 Ulm, Germany
| | | |
Collapse
|
26
|
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.
Collapse
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
| |
Collapse
|
27
|
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.
Collapse
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
| |
Collapse
|
28
|
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.
Collapse
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
| |
Collapse
|
29
|
Haller T, Cerrada A, Pfaller K, Braubach P, Felder E. Polarized light microscopy reveals physiological and drug-induced changes in surfactant membrane assembly in alveolar type II pneumocytes. BIOCHIMICA ET BIOPHYSICA ACTA-BIOMEMBRANES 2018; 1860:1152-1161. [DOI: 10.1016/j.bbamem.2018.01.010] [Citation(s) in RCA: 16] [Impact Index Per Article: 2.7] [Reference Citation Analysis] [Track Full Text] [Subscribe] [Scholar Register] [Received: 08/11/2017] [Revised: 12/05/2017] [Accepted: 01/04/2018] [Indexed: 12/16/2022]
|
30
|
Sander P, Mostafa H, Soboh A, Schneider JM, Pala A, Baron AK, Moepps B, Wirtz CR, Georgieff M, Schneider M. Vacquinol-1 inducible cell death in glioblastoma multiforme is counter regulated by TRPM7 activity induced by exogenous ATP. Oncotarget 2018; 8:35124-35137. [PMID: 28410232 PMCID: PMC5471040 DOI: 10.18632/oncotarget.16703] [Citation(s) in RCA: 27] [Impact Index Per Article: 4.5] [Reference Citation Analysis] [Abstract] [Key Words] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/03/2017] [Accepted: 03/15/2017] [Indexed: 12/29/2022] Open
Abstract
Glioblastomas (GBM) are the most malignant brain tumors in humans and have a very poor prognosis. New therapeutic options are urgently needed. A novel drug, Vacquinol-1 (Vac), a quinolone derivative, displays promising properties by inducing rapid cell death in GBM but not in non-transformed tissues. Features of this type of cell death are compatible with a process termed methuosis. Here we tested Vac on a highly malignant glioma cell line observed by long-term video microscopy. Human dental-pulp stem cells (DPSCs) served as controls. A major finding was that an exogenous ATP concentration of as little as 1 μM counter regulated the Vac-induced cell death. Studies using carvacrol, an inhibitor of transient receptor potential cation channel, subfamily M, member 7 (TRPM7), demonstrated that the ATP-inducible inhibitory effect is likely to be via TRPM7. Exogenous ATP is of relevance in GBM with large necrotic areas. Our results support the use of GBM cultures with different grades of malignancy to address their sensitivity to methuosis. The video-microscopy approach presented here allows decoding of signaling pathways as well as mechanisms of chemotherapeutic resistance by long-term observation. Before implementing Vac as a novel therapeutic drug in GBM, cells from each individual patient need to be assessed for their ATP sensitivity. In summary, the current investigation supports the concept of methuosis, described as non-apoptotic cell death and a promising approach for GBM treatment. Tissue-resident ATP/necrosis may interfere with this cell-death pathway but can be overcome by a natural compound, carvacrol that even penetrates the blood-brain barrier.
Collapse
Affiliation(s)
- Philip Sander
- Division of Experimental Anesthesiology, University Hospital Ulm, 89081 Ulm, Germany
| | - Haouraa Mostafa
- Division of Experimental Anesthesiology, University Hospital Ulm, 89081 Ulm, Germany
| | - Ayman Soboh
- Division of Experimental Anesthesiology, University Hospital Ulm, 89081 Ulm, Germany
| | - Julian M Schneider
- Division of Experimental Anesthesiology, University Hospital Ulm, 89081 Ulm, Germany
| | - Andrej Pala
- Department of Neurosurgery, Bezirkskrankenhaus Guenzburg, 89312 Guenzburg, Germany
| | - Ann-Kathrin Baron
- Department of Operative Dentistry and Periodontology, University Hospital Ulm, 89081 Ulm, Germany
| | - Barbara Moepps
- Institute of Pharmacology and Toxicology, University Hospital Ulm, 89081 Ulm, Germany
| | - C Rainer Wirtz
- Department of Neurosurgery, Bezirkskrankenhaus Guenzburg, 89312 Guenzburg, Germany
| | - Michael Georgieff
- Department of Anesthesiology, University Hospital Ulm, 89081 Ulm, Germany
| | - Marion Schneider
- Division of Experimental Anesthesiology, University Hospital Ulm, 89081 Ulm, Germany
| |
Collapse
|
31
|
Schaefer PM, Hilpert D, Niederschweiberer M, Neuhauser L, Kalinina S, Calzia E, Rueck A, von Einem B, von Arnim CAF. Mitochondrial matrix pH as a decisive factor in neurometabolic imaging. NEUROPHOTONICS 2017; 4:045004. [PMID: 29181426 PMCID: PMC5685807 DOI: 10.1117/1.nph.4.4.045004] [Citation(s) in RCA: 18] [Impact Index Per Article: 2.6] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Figures] [Subscribe] [Scholar Register] [Received: 08/14/2017] [Accepted: 10/16/2017] [Indexed: 06/07/2023]
Abstract
Alterations of cellular bioenergetics are a common feature in most neurodegenerative disorders. However, there is a selective vulnerability of different brain regions, cell types, and even mitochondrial populations to these metabolic disturbances. Thus, the aim of our study was to establish and validate an in vivo metabolic imaging technique to screen for mitochondrial function on the subcellular level. Based on nicotinamide adenine dinucleotide (phosphate) fluorescence lifetime imaging microscopy [NAD(P)H FLIM], we performed a quantitative correlation to high-resolution respirometry. Thereby, we revealed mitochondrial matrix pH as a decisive factor in imaging NAD(P)H redox state. By combining both parameters, we illustrate a quantitative, high-resolution assessment of mitochondrial function in metabolically modified cells as well as in an amyloid precursor protein-overexpressing model of Alzheimer's disease. Our metabolic imaging technique provides the basis for dissecting mitochondrial deficits not only in a range of neurodegenerative diseases, shedding light onto bioenergetic failures of cells remaining in their metabolic microenvironment.
Collapse
Affiliation(s)
| | - Diana Hilpert
- Ulm University, Department of Neurology, Ulm, Germany
| | | | | | - Sviatlana Kalinina
- Ulm University, Core Facility Confocal and Multiphoton Microscopy, Ulm, Germany
| | - Enrico Calzia
- University Medical School, Institute of Anesthesiological Pathophysiology and Process Engineering, Ulm, Germany
| | - Angelika Rueck
- Ulm University, Core Facility Confocal and Multiphoton Microscopy, Ulm, Germany
| | | | | |
Collapse
|
32
|
Becher A, Eiseler T, Porzner M, Walther P, Keil R, Bobrovich S, Hatzfeld M, Seufferlein T. The armadillo protein p0071 controls KIF3 motor transport. J Cell Sci 2017; 130:3374-3387. [PMID: 28808088 DOI: 10.1242/jcs.200170] [Citation(s) in RCA: 5] [Impact Index Per Article: 0.7] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/30/2016] [Accepted: 08/02/2017] [Indexed: 01/17/2023] Open
Abstract
We here report a novel function of the armadillo protein p0071 (also known as PKP4) during transport mediated by the KIF3 transport complex. Secretion of chromogranin A and matrix metallopeptidase 9 from pancreatic neuroendocrine tumor cells or pancreatic cancer cells, respectively, was substantially reduced following knockdown of p0071. Vesicle tracking indicated that there was impaired directional persistence of vesicle movement upon p0071 depletion. This suggests a disturbed balance between plus- and minus-end directed microtubule transport in cells lacking p0071. p0071 directly interacts with the KIF3 motor subunit KIF3B. Our data indicate that p0071 also interacts with the kinesin cargo adaptor protein KAP3 (also known as KIFAP3) acting as a stabilizing linker between KIF3B and its KAP3 cargo-binding entity. Thus, p0071 is required for directional vesicle movement and secretion of different KIF3-transported carriers, thereby regulating the transport of intracellular membrane vesicles along microtubules.
Collapse
Affiliation(s)
- Alexander Becher
- Department of Internal Medicine I, Ulm University, Albert-Einstein-Allee 23, 89081 Ulm, Germany
| | - Tim Eiseler
- Department of Internal Medicine I, Ulm University, Albert-Einstein-Allee 23, 89081 Ulm, Germany
| | - Marc Porzner
- Department of Internal Medicine I, Ulm University, Albert-Einstein-Allee 23, 89081 Ulm, Germany
| | - Paul Walther
- Central Facility for Electron Microscopy, Ulm University, Albert-Einstein-Allee 11, 89081, Ulm, Germany
| | - René Keil
- Institute of Molecular Medicine, Division of Pathobiochemistry, Martin-Luther-University of Halle-Wittenberg, D-06114 Halle, Germany
| | - Susanne Bobrovich
- Department of Internal Medicine I, Ulm University, Albert-Einstein-Allee 23, 89081 Ulm, Germany
| | - Mechthild Hatzfeld
- Institute of Molecular Medicine, Division of Pathobiochemistry, Martin-Luther-University of Halle-Wittenberg, D-06114 Halle, Germany
| | - Thomas Seufferlein
- Department of Internal Medicine I, Ulm University, Albert-Einstein-Allee 23, 89081 Ulm, Germany
| |
Collapse
|
33
|
Shah PNM, Stanifer ML, Höhn K, Engel U, Haselmann U, Bartenschlager R, Kräusslich HG, Krijnse-Locker J, Boulant S. Genome packaging of reovirus is mediated by the scaffolding property of the microtubule network. Cell Microbiol 2017; 19. [PMID: 28672089 DOI: 10.1111/cmi.12765] [Citation(s) in RCA: 19] [Impact Index Per Article: 2.7] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/07/2017] [Revised: 06/19/2017] [Accepted: 06/28/2017] [Indexed: 12/12/2022]
Abstract
Reovirus replication occurs in the cytoplasm of the host cell, in virally induced mini-organelles called virus factories. On the basis of the serotype of the virus, the virus factories can manifest as filamentous (type 1 Lang strain) or globular structures (type 3 Dearing strain). The filamentous factories morphology is dependent on the microtubule cytoskeleton; however, the exact function of the microtubule network in virus replication remains unknown. Using a combination of fluorescent microscopy, electron microscopy, and tomography of high-pressure frozen and freeze-substituted cells, we determined the ultrastructural organisation of reovirus factories. Cells infected with the reovirus microtubule-dependent strain display paracrystalline arrays of progeny virions resulting from their tiered organisation around microtubule filaments. On the contrary, in cells infected with the microtubule-independent strain, progeny virions lacked organisation. Conversely to the microtubule-dependent strain, around half of the viral particles present in these viral factories did not contain genomes (genome-less particles). Complementarily, interference with the microtubule filaments in cells infected with the microtubule-dependent strain resulted in a significant increase of genome-less particle number. This decrease of genome packaging efficiency could be rescued by rerouting viral factories on the actin cytoskeleton. These findings demonstrate that the scaffolding properties of the microtubule, and not biochemical nature of tubulin, are critical determinants for reovirus efficient genome packaging. This work establishes, for the first time, a functional correlation between ultrastructural organisation of reovirus factories with genome packaging efficiency and provides novel information on how viruses coordinate assembly of progeny particles.
Collapse
Affiliation(s)
- Pranav N M Shah
- Department of Infectious Diseases, Virology, Heidelberg University Hospital, Heidelberg, Germany.,Schaller Research Group at CellNetworks and DKFZ, Heidelberg, Germany
| | - Megan L Stanifer
- Department of Infectious Diseases, Virology, Heidelberg University Hospital, Heidelberg, Germany.,Schaller Research Group at CellNetworks and DKFZ, Heidelberg, Germany
| | - Katharina Höhn
- Department of Infectious Diseases, Virology, Heidelberg University Hospital, Heidelberg, Germany
| | - Ulrike Engel
- Nikon Imaging Center, Heidelberg University, Heidelberg, Germany
| | - Uta Haselmann
- Department of Infectious Diseases, Molecular Virology, Heidelberg University Hospital, Germany
| | - Ralf Bartenschlager
- Department of Infectious Diseases, Molecular Virology, Heidelberg University Hospital, Germany
| | - Hans-Georg Kräusslich
- Department of Infectious Diseases, Virology, Heidelberg University Hospital, Heidelberg, Germany
| | - Jacomine Krijnse-Locker
- Department of Infectious Diseases, Virology, Heidelberg University Hospital, Heidelberg, Germany.,Ultrapole, Ultrastructural Bio-imaging, Center for Innovation and Technological Research, Institut Pasteur, Paris, France
| | - Steeve Boulant
- Department of Infectious Diseases, Virology, Heidelberg University Hospital, Heidelberg, Germany.,Schaller Research Group at CellNetworks and DKFZ, Heidelberg, Germany
| |
Collapse
|
34
|
The sleeping beauty kissed awake: new methods in electron microscopy to study cellular membranes. Biochem J 2017; 474:1041-1053. [DOI: 10.1042/bcj20160990] [Citation(s) in RCA: 6] [Impact Index Per Article: 0.9] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/08/2016] [Revised: 01/03/2017] [Accepted: 01/23/2017] [Indexed: 01/12/2023]
Abstract
Electron microscopy (EM) for biological samples, developed in the 1940–1950s, changed our conception about the architecture of eukaryotic cells. It was followed by a period where EM applied to cell biology had seemingly fallen asleep, even though new methods with important implications for modern EM were developed. Among these was the discovery that samples can be preserved by chemical fixation and most importantly by rapid freezing without the formation of crystalline ice, giving birth to the world of cryo-EM. The past 15–20 years are hallmarked by a tremendous interest in EM, driven by important technological advances. Cryo-EM, in particular, is now capable of revealing structures of proteins at a near-atomic resolution owing to improved sample preparation methods, microscopes and cameras. In this review, we focus on the challenges associated with the imaging of membranes by EM and give examples from the field of host–pathogen interactions, in particular of virus-infected cells. Despite the advantages of imaging membranes under native conditions in cryo-EM, conventional EM will remain an important complementary method, in particular if large volumes need to be imaged.
Collapse
|
35
|
Müller A, Neukam M, Ivanova A, Sönmez A, Münster C, Kretschmar S, Kalaidzidis Y, Kurth T, Verbavatz JM, Solimena M. A Global Approach for Quantitative Super Resolution and Electron Microscopy on Cryo and Epoxy Sections Using Self-labeling Protein Tags. Sci Rep 2017; 7:23. [PMID: 28154417 PMCID: PMC5428382 DOI: 10.1038/s41598-017-00033-x] [Citation(s) in RCA: 33] [Impact Index Per Article: 4.7] [Reference Citation Analysis] [Abstract] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/01/2016] [Accepted: 12/20/2016] [Indexed: 01/19/2023] Open
Abstract
Correlative light and electron microscopy (CLEM) is a powerful approach to investigate the molecular ultrastructure of labeled cell compartments. However, quantitative CLEM studies are rare, mainly due to small sample sizes and the sensitivity of fluorescent proteins to strong fixatives and contrasting reagents for EM. Here, we show that fusion of a self-labeling protein to insulin allows for the quantification of age-distinct insulin granule pools in pancreatic beta cells by a combination of super resolution and transmission electron microscopy on Tokuyasu cryosections. In contrast to fluorescent proteins like GFP organic dyes covalently bound to self-labeling proteins retain their fluorescence also in epoxy resin following high pressure freezing and freeze substitution, or remarkably even after strong chemical fixation. This enables for the assessment of age-defined granule morphology and degradation. Finally, we demonstrate that this CLEM protocol is highly versatile, being suitable for single and dual fluorescent labeling and detection of different proteins with optimal ultrastructure preservation and contrast.
Collapse
Affiliation(s)
- Andreas Müller
- Molecular Diabetology, University Hospital and Faculty of Medicine Carl Gustav Carus, TU Dresden, Dresden, Germany.,Paul Langerhans Institute Dresden (PLID) of the Helmholtz Center Munich at the University Hospital Carl Gustav Carus and Faculty of Medicine of the TU Dresden, Dresden, Germany
| | - Martin Neukam
- Molecular Diabetology, University Hospital and Faculty of Medicine Carl Gustav Carus, TU Dresden, Dresden, Germany.,Paul Langerhans Institute Dresden (PLID) of the Helmholtz Center Munich at the University Hospital Carl Gustav Carus and Faculty of Medicine of the TU Dresden, Dresden, Germany
| | - Anna Ivanova
- Molecular Diabetology, University Hospital and Faculty of Medicine Carl Gustav Carus, TU Dresden, Dresden, Germany.,Paul Langerhans Institute Dresden (PLID) of the Helmholtz Center Munich at the University Hospital Carl Gustav Carus and Faculty of Medicine of the TU Dresden, Dresden, Germany
| | - Anke Sönmez
- Molecular Diabetology, University Hospital and Faculty of Medicine Carl Gustav Carus, TU Dresden, Dresden, Germany.,Paul Langerhans Institute Dresden (PLID) of the Helmholtz Center Munich at the University Hospital Carl Gustav Carus and Faculty of Medicine of the TU Dresden, Dresden, Germany
| | - Carla Münster
- Molecular Diabetology, University Hospital and Faculty of Medicine Carl Gustav Carus, TU Dresden, Dresden, Germany.,Paul Langerhans Institute Dresden (PLID) of the Helmholtz Center Munich at the University Hospital Carl Gustav Carus and Faculty of Medicine of the TU Dresden, Dresden, Germany
| | - Susanne Kretschmar
- Center for Regenerative Therapies Dresden (CRTD), TU Dresden, Dresden, Germany.,Biotechnology Center of the TU Dresden (BIOTEC), Dresden, Germany
| | - Yannis Kalaidzidis
- Max Planck Institute of Molecular Cell Biology and Genetics (MPI-CBG), Dresden, Germany.,Faculty of Bioengineering and Bioinformatics, Moscow State University, Moscow, Russia
| | - Thomas Kurth
- Center for Regenerative Therapies Dresden (CRTD), TU Dresden, Dresden, Germany.,Biotechnology Center of the TU Dresden (BIOTEC), Dresden, Germany
| | - Jean-Marc Verbavatz
- Max Planck Institute of Molecular Cell Biology and Genetics (MPI-CBG), Dresden, Germany.,Institut Jacques Monod, Université Paris Diderot, Paris, France
| | - Michele Solimena
- Molecular Diabetology, University Hospital and Faculty of Medicine Carl Gustav Carus, TU Dresden, Dresden, Germany. .,Paul Langerhans Institute Dresden (PLID) of the Helmholtz Center Munich at the University Hospital Carl Gustav Carus and Faculty of Medicine of the TU Dresden, Dresden, Germany. .,Max Planck Institute of Molecular Cell Biology and Genetics (MPI-CBG), Dresden, Germany.
| |
Collapse
|
36
|
Weil MT, Ruhwedel T, Möbius W, Simons M. Intracerebral Injections and Ultrastructural Analysis of High-Pressure Frozen Brain Tissue. ACTA ACUST UNITED AC 2017; 78:2.27.1-2.27.18. [PMID: 28046202 DOI: 10.1002/cpns.22] [Citation(s) in RCA: 5] [Impact Index Per Article: 0.7] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/10/2022]
Abstract
Intracerebral injections are an invasive method to bypass the blood brain barrier and are widely used to study molecular and cellular mechanisms of the central nervous system. The administered substances are injected directly at the site of interest, executing their effect locally. By combining injections in the rat brain with state-of-the-art electron microscopy, subtle changes in ultrastructure of the nervous tissue can be detected prior to overt damage or disease. The protocol presented here involves stereotactic injection into the corpus callosum of Lewis rats and the cryopreparation of freshly dissected tissue for electron microscopy. The localization of the injection site in tissue sections during the sample preparation for transmission electron microscopy is explained and possible artifacts of the method are indicated. With the help of this powerful combination of injections and electron microscopy, subtle effects of the applied substances on the biology of neural cells can be identified and monitored over time. © 2017 by John Wiley & Sons, Inc.
Collapse
Affiliation(s)
- Marie-Theres Weil
- Department of Cellular Neuroscience, Max-Planck Institute for Experimental Medicine, Göttingen, Germany.,Department of Neurogenetics, Max-Planck Institute for Experimental Medicine, Göttingen, Germany.,Center Nanoscale Microscopy and Molecular Physiology of the Brain (CNMPB), Göttingen, Germany
| | - Torben Ruhwedel
- Department of Neurogenetics, Max-Planck Institute for Experimental Medicine, Göttingen, Germany
| | - Wiebke Möbius
- Department of Neurogenetics, Max-Planck Institute for Experimental Medicine, Göttingen, Germany.,Center Nanoscale Microscopy and Molecular Physiology of the Brain (CNMPB), Göttingen, Germany
| | - Mikael Simons
- Department of Cellular Neuroscience, Max-Planck Institute for Experimental Medicine, Göttingen, Germany.,Institute of Neuronal Cell Biology, The Technical University of Munich, Munich, Germany.,German Center for Neurodegenerative Disease (DZNE), Munich, Germany.,Munich Cluster for Systems Neurology (SyNergy), Munich, Germany
| |
Collapse
|
37
|
Schaefer PM, von Einem B, Walther P, Calzia E, von Arnim CAF. Metabolic Characterization of Intact Cells Reveals Intracellular Amyloid Beta but Not Its Precursor Protein to Reduce Mitochondrial Respiration. PLoS One 2016; 11:e0168157. [PMID: 28005987 PMCID: PMC5178995 DOI: 10.1371/journal.pone.0168157] [Citation(s) in RCA: 19] [Impact Index Per Article: 2.4] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/22/2016] [Accepted: 11/24/2016] [Indexed: 12/17/2022] Open
Abstract
One hallmark of Alzheimer´s disease are senile plaques consisting of amyloid beta (Aβ), which derives from the processing of the amyloid precursor protein (APP). Mitochondrial dysfunction has been linked to the pathogenesis of Alzheimer´s disease and both Aβ and APP have been reported to affect mitochondrial function in isolated systems. However, in intact cells, considering a physiological localization of APP and Aβ, it is pending what triggers the mitochondrial defect. Thus, the aim of this study was to dissect the impact of APP versus Aβ in inducing mitochondrial alterations with respect to their subcellular localization. We performed an overexpression of APP or beta-site amyloid precursor protein cleaving enzyme 1 (BACE1), increasing APP and Aβ levels or Aβ alone, respectively. Conducting a comprehensive metabolic characterization we demonstrate that only APP overexpression reduced mitochondrial respiration, despite lower extracellular Aβ levels compared to BACE overexpression. Surprisingly, this could be rescued by a gamma secretase inhibitor, oppositionally indicating an Aβ-mediated mitochondrial toxicity. Analyzing Aβ localization revealed that intracellular levels of Aβ and an increased spatial association of APP/Aβ with mitochondria are associated with reduced mitochondrial respiration. Thus, our data provide marked evidence for a prominent role of intracellular Aβ accumulation in Alzheimer´s disease associated mitochondrial dysfunction. Thereby it highlights the importance of the localization of APP processing and intracellular transport as a decisive factor for mitochondrial function, linking two prominent hallmarks of neurodegenerative diseases.
Collapse
Affiliation(s)
| | | | - Paul Walther
- Central Facility for Electron Microscopy, Ulm University, Ulm, Germany
| | - Enrico Calzia
- Institut für Anästhesiologische Pathophysiologie und Verfahrensentwicklung, Universitätsklinikum Ulm, Ulm, Germany
| | | |
Collapse
|
38
|
Ultrastructural Characterization of Phagophores Using Electron Tomography on Cryoimmobilized and Freeze Substituted Samples. Methods Enzymol 2016; 587:331-349. [PMID: 28253964 DOI: 10.1016/bs.mie.2016.09.063] [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: 02/17/2023]
Abstract
Electron tomography has significantly contributed to recent findings regarding the biogenesis of the phagophore, an organelle which initiates autophagic sequestration. The information obtained from 1.9nm slices through the tomograms have revealed that during biogenesis the phagophore is in contact with the membranes of apposing organelles to form tubular connections and membrane contact sites (MCSs). The most reported and established tubular connections occur between the phagophore and the endoplasmic reticulum. However, as the phagophore continues to grow and expand, connections and MCSs have also been reported to occur between the phagophore and several other organelles in a possible attempt to utilize lipids for membrane expansion from alternative sources. Since the lifespan of the phagophore is only a few minutes and membrane connections and MCSs are very dynamic, capturing these two events requires precision during fixation. Up to date there is no quicker alternative for sample preservation in transmission electron microscopy than cryoimmobilization. In this report, we describe our protocol for cryoimmobilization using high-pressure freezing and freeze substitution, and report our first findings on phagophore morphology using this approach.
Collapse
|
39
|
Reversible Cryopreservation of Living Cells Using an Electron Microscopy Cryo-Fixation Method. PLoS One 2016; 11:e0164270. [PMID: 27711254 PMCID: PMC5053471 DOI: 10.1371/journal.pone.0164270] [Citation(s) in RCA: 10] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/01/2016] [Accepted: 09/22/2016] [Indexed: 02/01/2023] Open
Abstract
Rapid cooling of aqueous solutions is a useful approach for two important biological applications: (I) cryopreservation of cells and tissues for long-term storage, and (II) cryofixation for ultrastructural investigations by electron and cryo-electron microscopy. Usually, both approaches are very different in methodology. Here we show that a novel, fast and easy to use cryofixation technique called self-pressurized rapid freezing (SPRF) is–after some adaptations–also a useful and versatile technique for cryopreservation. Sealed metal tubes with high thermal diffusivity containing the samples are plunged into liquid cryogen. Internal pressure builds up reducing ice crystal formation and therefore supporting reversible cryopreservation through vitrification of cells. After rapid rewarming of pressurized samples, viability rates of > 90% can be reached, comparable to best-performing of the established rapid cooling devices tested. In addition, the small SPRF tubes allow for space-saving sample storage and the sealed containers prevent contamination from or into the cryogen during freezing, storage, or thawing.
Collapse
|
40
|
Nafeey S, Martin I, Felder T, Walther P, Felder E. Branching of keratin intermediate filaments. J Struct Biol 2016; 194:415-22. [PMID: 27039023 DOI: 10.1016/j.jsb.2016.03.023] [Citation(s) in RCA: 11] [Impact Index Per Article: 1.4] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/03/2015] [Revised: 03/22/2016] [Accepted: 03/30/2016] [Indexed: 11/18/2022]
Abstract
Keratin intermediate filaments (IFs) are crucial to maintain mechanical stability in epithelial cells. Since little is known about the network architecture that provides this stiffness and especially about branching properties of filaments, we addressed this question with different electron microscopic (EM) methods. Using EM tomography of high pressure frozen keratinocytes, we investigated the course of several filaments in a branching of a filament bundle. Moreover we found several putative bifurcations in individual filaments. To verify our observation we also visualized the keratin network in detergent extracted keratinocytes with scanning EM. Here bifurcations of individual filaments could unambiguously be identified additionally to bundle branchings. Interestingly, identical filament bifurcations were also found in purified keratin 8/18 filaments expressed in Escherichia coli which were reassembled in vitro. This excludes that an accessory protein contributes to the branch formation. Measurements of the filament cross sectional areas showed various ratios between the three bifurcation arms. This demonstrates that intermediate filament furcation is very different from actin furcation where an entire new filament is attached to an existing filament. Instead, the architecture of intermediate filament bifurcations is less predetermined and hence consistent with the general concept of IF formation.
Collapse
Affiliation(s)
- Soufi Nafeey
- Central Facility for Electron Microscopy, Ulm University, 89081 Ulm, Germany
| | - Ines Martin
- Institute of Experimental Physics, Ulm University, 89081 Ulm, Germany
| | - Tatiana Felder
- Institute of General Physiology, Ulm University, 89081 Ulm, Germany
| | - Paul Walther
- Central Facility for Electron Microscopy, Ulm University, 89081 Ulm, Germany.
| | - Edward Felder
- Institute of General Physiology, Ulm University, 89081 Ulm, Germany
| |
Collapse
|
41
|
Crauwels P, Bohn R, Thomas M, Gottwalt S, Jäckel F, Krämer S, Bank E, Tenzer S, Walther P, Bastian M, van Zandbergen G. Apoptotic-like Leishmania exploit the host's autophagy machinery to reduce T-cell-mediated parasite elimination. Autophagy 2016; 11:285-97. [PMID: 25801301 PMCID: PMC4502818 DOI: 10.1080/15548627.2014.998904] [Citation(s) in RCA: 50] [Impact Index Per Article: 6.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/19/2022] Open
Abstract
Apoptosis is a well-defined cellular process in which a cell dies, characterized by cell shrinkage and DNA fragmentation. In parasites like Leishmania, the process of apoptosis-like cell death has been described. Moreover upon infection, the apoptotic-like population is essential for disease development, in part by silencing host phagocytes. Nevertheless, the exact mechanism of how apoptosis in unicellular organisms may support infectivity remains unclear. Therefore we investigated the fate of apoptotic-like Leishmania parasites in human host macrophages. Our data showed—in contrast to viable parasites—that apoptotic-like parasites enter an LC3+, autophagy-like compartment. The compartment was found to consist of a single lipid bilayer, typical for LC3-associated phagocytosis (LAP). As LAP can provoke anti-inflammatory responses and autophagy modulates antigen presentation, we analyzed how the presence of apoptotic-like parasites affected the adaptive immune response. Macrophages infected with viable Leishmania induced proliferation of CD4+ T-cells, leading to a reduced intracellular parasite survival. Remarkably, the presence of apoptotic-like parasites in the inoculum significantly reduced T-cell proliferation. Chemical induction of autophagy in human monocyte-derived macrophage (hMDM), infected with viable parasites only, had an even stronger proliferation-reducing effect, indicating that host cell autophagy and not parasite viability limits the T-cell response and enhances parasite survival. Concluding, our data suggest that apoptotic-like Leishmania hijack the host cells´ autophagy machinery to reduce T-cell proliferation. Furthermore, the overall population survival is guaranteed, explaining the benefit of apoptosis-like cell death in a single-celled parasite and defining the host autophagy pathway as a potential therapeutic target in treating Leishmaniasis.
Collapse
Key Words
- ANXA5, annexin V
- CFSE, carboxyfluorescein succinimidyl ester
- CM, complete medium
- IF, immunofluorescence
- IL, interleukin
- LAP
- LAP, LC3-associated phagocytosis
- Lm, Leishmania
- MACS, magnetic-associated cell sorting
- MAP1LC3/LC3, microtubule-associated protein 1 light chain 3
- MFI, mean fluorescence intensity
- MHC, major histocompatibility complex
- MOI, multiplicity of infection
- PBMCs, peripheral blood mononuclear cells
- PS, phosphatidylserine
- T-cell proliferation
- TGFB, transforming growth factor
- anti-inflammatory
- apoptotic-like Leishmania
- autophagy
- hMDM, human monocyte derived macrophage
- human primary macrophages
- immune evasion
- log.ph, logarithmic phase
- stat.ph, stationary phase
- β; TT, tetanus toxoid
Collapse
Affiliation(s)
- Peter Crauwels
- a Division of Immunology ; Paul-Ehrlich-Institute ; Langen , Germany
| | | | | | | | | | | | | | | | | | | | | |
Collapse
|
42
|
Bauer A, Subramanian N, Villinger C, Frascaroli G, Mertens T, Walther P. Megapinocytosis: a novel endocytic pathway. Histochem Cell Biol 2016; 145:617-27. [DOI: 10.1007/s00418-015-1395-2] [Citation(s) in RCA: 9] [Impact Index Per Article: 1.1] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Accepted: 12/15/2015] [Indexed: 01/13/2023]
|
43
|
Celler K, Fujita M, Kawamura E, Ambrose C, Herburger K, Holzinger A, Wasteneys GO. Microtubules in Plant Cells: Strategies and Methods for Immunofluorescence, Transmission Electron Microscopy, and Live Cell Imaging. Methods Mol Biol 2016; 1365:155-84. [PMID: 26498784 DOI: 10.1007/978-1-4939-3124-8_8] [Citation(s) in RCA: 12] [Impact Index Per Article: 1.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/22/2022]
Abstract
Microtubules (MTs) are required throughout plant development for a wide variety of processes, and different strategies have evolved to visualize and analyze them. This chapter provides specific methods that can be used to analyze microtubule organization and dynamic properties in plant systems and summarizes the advantages and limitations for each technique. We outline basic methods for preparing samples for immunofluorescence labeling, including an enzyme-based permeabilization method, and a freeze-shattering method, which generates microfractures in the cell wall to provide antibodies access to cells in cuticle-laden aerial organs such as leaves. We discuss current options for live cell imaging of MTs with fluorescently tagged proteins (FPs), and provide chemical fixation, high-pressure freezing/freeze substitution, and post-fixation staining protocols for preserving MTs for transmission electron microscopy and tomography.
Collapse
Affiliation(s)
- Katherine Celler
- Department of Botany, The University of British Columbia, Vancouver, BC, Canada
| | - Miki Fujita
- Department of Botany, The University of British Columbia, Vancouver, BC, Canada
| | - Eiko Kawamura
- Western College of Veterinary Medicine, University of Saskatchewan, Saskatoon, SK, Canada
| | - Chris Ambrose
- Department of Biology, University of Saskatchewan, Saskatoon, SK, Canada
| | - Klaus Herburger
- Functional Plant Biology, Institute of Botany, University of Innsbruck, Sternwartestraße 15, 6020, Innsbruck, Austria
| | - Andreas Holzinger
- Functional Plant Biology, Institute of Botany, University of Innsbruck, Sternwartestraße 15, 6020, Innsbruck, Austria.
| | | |
Collapse
|
44
|
A new role for an old drug: Ambroxol triggers lysosomal exocytosis via pH-dependent Ca²⁺ release from acidic Ca²⁺ stores. Cell Calcium 2015; 58:628-37. [PMID: 26560688 DOI: 10.1016/j.ceca.2015.10.002] [Citation(s) in RCA: 36] [Impact Index Per Article: 4.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/15/2015] [Revised: 10/20/2015] [Accepted: 10/22/2015] [Indexed: 12/21/2022]
Abstract
Ambroxol (Ax) is a frequently prescribed drug used to facilitate mucociliary clearance, but its mode of action is yet poorly understood. Here we show by X-ray spectroscopy that Ax accumulates in lamellar bodies (LBs), the surfactant storing, secretory lysosomes of type II pneumocytes. Using lyso- and acidotropic substances in combination with fluorescence imaging we confirm that these vesicles belong to the class of acidic Ca(2+) stores. Ax lead to a significant neutralization of LB pH, followed by intracellular Ca(2+) release, and to a dose-dependent surfactant exocytosis. Ax-induced Ca(2+) release was significantly reduced and slowed down by pretreatment of the cells with bafilomycin A1 (Baf A1), an inhibitor of the vesicular H(+) ATPase. These results could be nearly reproduced with NH3/NH4(+). The findings suggest that Ax accumulates within LBs and severely affects their H(+) and Ca(2+) homeostasis. This is further supported by an Ax-induced change of nanostructural assembly of surfactant layers. We conclude that Ax profoundly affects LBs presumably by disordering lipid bilayers and by acting as a weak base. The pH change triggers - at least in part - Ca(2+) release from stores and secretion of surfactant from type II cells. This novel mechanism of Ax as a lysosomal secretagogue may also play a role for its recently discussed use for lysosomal storage and other degenerative diseases.
Collapse
|
45
|
Loussert Fonta C, Humbel BM. Correlative microscopy. Arch Biochem Biophys 2015; 581:98-110. [DOI: 10.1016/j.abb.2015.05.017] [Citation(s) in RCA: 63] [Impact Index Per Article: 7.0] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/09/2015] [Revised: 05/26/2015] [Accepted: 05/29/2015] [Indexed: 11/15/2022]
|
46
|
Suarez C, Andres G, Kolovou A, Hoppe S, Salas ML, Walther P, Krijnse Locker J. African swine fever virus assembles a single membrane derived from rupture of the endoplasmic reticulum. Cell Microbiol 2015; 17:1683-98. [PMID: 26096327 DOI: 10.1111/cmi.12468] [Citation(s) in RCA: 32] [Impact Index Per Article: 3.6] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/26/2015] [Revised: 05/05/2015] [Accepted: 05/19/2015] [Indexed: 12/13/2022]
Abstract
Collective evidence argues that two members of the nucleocytoplasmic large DNA viruses (NCLDVs) acquire their membrane from open membrane intermediates, postulated to be derived from membrane rupture. We now study membrane acquisition of the NCLDV African swine fever virus. By electron tomography (ET), the virion assembles a single bilayer, derived from open membrane precursors that collect as ribbons in the cytoplasm. Biochemically, lumenal endoplasmic reticulum (ER) proteins are released into the cytosol, arguing that the open intermediates are ruptured ER membranes. ET shows that viral capsid assembles on the convex side of the open viral membrane to shape it into an icosahedron. The viral capsid is composed of tiny spikes with a diameter of ∼5 nm, connected to the membrane by a 6 nm wide structure displaying thin striations, as observed by several complementary electron microscopy imaging methods. Immature particles display an opening that closes after uptake of the viral genome and core proteins, followed by the formation of the mature virion. Together with our previous data, this study shows a common principle of NCLDVs to build a single internal envelope from open membrane intermediates. Our data now provide biochemical evidence that these open intermediates result from rupture of a cellular membrane, the ER.
Collapse
Affiliation(s)
- Cristina Suarez
- Electron Microscopy (EM) Core Facility and Department of Infectious Diseases, Heidelberg University Hospital, Heidelberg, Germany
| | - German Andres
- Electron Microscopy (EM) Unit, Centro de Biologia Molecular Severo Ochoa, Consejo Superior de Investigaciones Científicas-Universidad Autónoma de Madrid, Madrid, Spain
| | - Androniki Kolovou
- Electron Microscopy (EM) Core Facility and Department of Infectious Diseases, Heidelberg University Hospital, Heidelberg, Germany
| | - Simone Hoppe
- Electron Microscopy (EM) Core Facility and Department of Infectious Diseases, Heidelberg University Hospital, Heidelberg, Germany
| | - Maria L Salas
- Virology Department, Centro de Biologia Molecular Severo Ochoa, Consejo Superior de Investigaciones Científicas-Universidad Autónoma de Madrid, Madrid, Spain
| | - Paul Walther
- Electron Microscopy (EM) Core Facility, University of Ulm, Ulm, Germany
| | - Jacomine Krijnse Locker
- Electron Microscopy (EM) Core Facility and Department of Infectious Diseases, Heidelberg University Hospital, Heidelberg, Germany
| |
Collapse
|
47
|
McDonald KL. Out with the old and in with the new: rapid specimen preparation procedures for electron microscopy of sectioned biological material. PROTOPLASMA 2014; 251:429-448. [PMID: 24258967 DOI: 10.1007/s00709-013-0575-y] [Citation(s) in RCA: 55] [Impact Index Per Article: 5.5] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Received: 10/21/2013] [Accepted: 10/22/2013] [Indexed: 06/02/2023]
Abstract
This article presents the best current practices for preparation of biological samples for examination as thin sections in an electron microscope. The historical development of fixation, dehydration, and embedding procedures for biological materials are reviewed for both conventional and low temperature methods. Conventional procedures for processing cells and tissues are usually done over days and often produce distortions, extractions, and other artifacts that are not acceptable for today's structural biology standards. High-pressure freezing and freeze substitution can minimize some of these artifacts. New methods that reduce the times for freeze substitution and resin embedding to a few hours are discussed as well as a new rapid room temperature method for preparing cells for on-section immunolabeling without the use of aldehyde fixatives.
Collapse
Affiliation(s)
- Kent L McDonald
- Electron Microscope Laboratory, University of California, 26 Giannini Hall, Berkeley, CA, 94720, USA,
| |
Collapse
|
48
|
Wilkat M, Herdoiza E, Forsbach-Birk V, Walther P, Essig A. Electron tomography and cryo-SEM characterization reveals novel ultrastructural features of host-parasite interaction during Chlamydia abortus infection. Histochem Cell Biol 2014; 142:171-84. [PMID: 24522393 DOI: 10.1007/s00418-014-1189-y] [Citation(s) in RCA: 12] [Impact Index Per Article: 1.2] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Accepted: 01/22/2014] [Indexed: 01/06/2023]
Abstract
Chlamydia (C.) abortus is a widely spread pathogen among ruminants that can be transmitted to women during pregnancy leading to severe systemic infection with consecutive abortion. As a member of the Chlamydiaceae, C. abortus shares the characteristic feature of an obligate intracellular biphasic developmental cycle with two morphological forms including elementary bodies (EBs) and reticulate bodies (RBs). In contrast to other chlamydial species, C. abortus ultrastructure has not been investigated yet. To do so, samples were fixed by high-pressure freezing and processed by different electron microscopic methods. Freeze-substituted samples were analysed by transmission electron microscopy, scanning transmission electron microscopical tomography and immuno-electron microscopy, and freeze-fractured samples were analysed by cryo-scanning electron microscopy. Here, we present three ultrastructural features of C. abortus that have not been reported up to now. Firstly, the morphological evidence that C. abortus is equipped with the type three secretion system. Secondly, the accumulation and even coating of whole inclusion bodies by membrane complexes consisting of multiple closely adjacent membranes which seems to be a C. abortus specific feature. Thirdly, the formation of small vesicles in the periplasmic space of RBs in the second half of the developmental cycle. Concerning the time point of their formation and the fact that they harbour chlamydial components, these vesicles might be morphological correlates of an intermediate step during the process of redifferentiation of RBs into EBs. As this feature has also been shown for C. trachomatis and C. pneumoniae, it might be a common characteristic of the family of Chlamydiaceae.
Collapse
Affiliation(s)
- M Wilkat
- Institute of Medical Microbiology and Hygiene, University Hospital of Ulm, Albert-Einstein-Allee 23, 89081, Ulm, Germany,
| | | | | | | | | |
Collapse
|
49
|
Abstract
High-pressure freeze fixation is the method of choice to arrest instantly all dynamic and physiological processes inside cells, tissues, and small organisms. Embedded in vitreous ice, such samples can be further processed by freeze substitution or directly analyzed in their fully hydrated state by cryo-electron microscopy of vitreous sections (CEMOVIS) to explore cellular ultrastructure as close as possible to the native state. Here, we describe the procedure of self-pressurized rapid freezing as fast, easy-to-use, and low-cost freeze fixation method, avoiding the usage of a high-pressure freezing (HPF) apparatus. Cells or small organisms are placed in capillary metal tubes, which are tightly closed and plunged directly into liquid ethane cooled by liquid nitrogen. In parts of the tube, crystalline ice is formed and builds up pressure sufficient for the liquid-glass transition of the remaining specimen. The quality of samples is equivalent to preparations by conventional HPF apparatus, allowing for high-resolution cryo-EM applications or for freeze substitution and plastic embedding.
Collapse
Affiliation(s)
- Markus Grabenbauer
- Institute for Anatomy and Cell Biology Germany, University of Heidelberg, Heidelberg, Germany
| | | | | |
Collapse
|
50
|
Abstract
In this chapter we describe three different approaches for three-dimensional imaging of electron microscopic samples: serial sectioning transmission electron microscopy (TEM), scanning transmission electron microscopy (STEM) tomography, and focused ion beam/scanning electron microscopy (FIB/SEM) tomography. With these methods, relatively large volumes of resin-embedded biological structures can be analyzed at resolutions of a few nm within a reasonable expenditure of time. The traditional method is serial sectioning and imaging the same area in all sections. Another method is TEM tomography that involves tilting a section in the electron beam and then reconstruction of the volume by back projection of the images. When the scanning transmission (STEM) mode is used, thicker sections (up to 1 μm) can be analyzed. The third approach presented here is focused ion beam/scanning electron microscopy (FIB/SEM) tomography, in which a sample is repeatedly milled with a focused ion beam (FIB) and each newly produced block face is imaged with the scanning electron microscope (SEM). This process can be repeated ad libitum in arbitrary small increments allowing 3D analysis of relatively large volumes such as eukaryotic cells. We show that resolution of this approach is considerably improved when the secondary electron signal is used. However, the most important prerequisite for three-dimensional imaging is good specimen preparation. For all three imaging methods, cryo-fixed (high-pressure frozen) and freeze-substituted samples have been used.
Collapse
|