1
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Choquet D, Petrel M, Fernández-Monreal M. Targeting of membrane proteins with fluoronanogold probes for high-resolution correlative microscopy. Methods Cell Biol 2024; 187:57-72. [PMID: 38705630 DOI: 10.1016/bs.mcb.2024.02.035] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 05/07/2024]
Abstract
Correlative light and electron microscopy (CLEM) can provide valuable information about a biological sample by giving information on the specific localization of a molecule of interest within an ultrastructural context. In this work, we describe a simple CLEM method to obtain high-resolution images of neurotransmitter receptor distribution in synapses by electron microscopy (EM). We use hippocampal organotypic slices from a previously reported mouse model expressing a modified AMPA receptor (AMPAR) subunit that binds biotin at the surface (Getz et al., 2022). This tag can be recognized by StreptAvidin-Fluoronanogold™ conjugates (SA-FNG), which reach receptors at synapses (synaptic cleft is 50-100nm thick). By using pre-embedding labeling, we found that SA-FNG reliably bind synaptic receptors and penetrate around 10-15μm in depth in live tissue. However, the silver enhancement was only reaching the surface of the slices. We show that permeabilization with triton is highly effective at increasing the in depth-gold amplification and that the membrane integrity is well preserved. Finally, we also apply high-resolution electron tomography, thus providing important information about the 3D organization of surface AMPA receptors in synapses at the nanoscale.
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Affiliation(s)
- Daniel Choquet
- Université de Bordeaux, CNRS, INSERM, Bordeaux Imaging Center (BIC), Bordeaux, France; Université de Bordeaux, CNRS, Interdisciplinary Institute for Neuroscience (IINS), Bordeaux, France
| | - Melina Petrel
- Université de Bordeaux, CNRS, INSERM, Bordeaux Imaging Center (BIC), Bordeaux, France
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2
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Fredericks K, Kriel J, Engelbrecht L, Mercea PA, Widhalm G, Harrington B, Vlok I, Loos B. 5-ALA localises to the autophagy compartment and increases its fluorescence upon autophagy enhancement through caloric restriction and spermidine treatment in human glioblastoma. Biochem Biophys Rep 2024; 37:101642. [PMID: 38288282 PMCID: PMC10823107 DOI: 10.1016/j.bbrep.2024.101642] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/20/2023] [Revised: 12/29/2023] [Accepted: 01/08/2024] [Indexed: 01/31/2024] Open
Abstract
Glioblastoma Multiforme (GBM) is the most invasive and prevalent Central Nervous System (CNS) malignancy. It is characterised by diffuse infiltrative growth and metabolic dysregulation that impairs the extent of surgical resection (EoR), contributing to its poor prognosis. 5-Aminolevulinic acid (5-ALA) fluorescence-guided surgical resection (FGR) takes advantage of the preferential generation of 5-ALA-derived fluorescence signal in glioma cells, thereby improving visualisation and enhancing the EoR. However, despite 5-ALA FGR is a widely used technique in the surgical management of malignant gliomas, the infiltrative tumour margins usually show only vague or no visible fluorescence and thus a significant amount of residual tumour tissue may hence remain in the resection cavity, subsequently driving tumour recurrence. To investigate the molecular mechanisms that govern the preferential accumulation of 5-ALA in glioma cells, we investigated the precise subcellular localisation of 5-ALA signal using Correlative Light and Electron Microscopy (CLEM) and colocalisation analyses in U118MG glioma cells. Our results revealed strong 5-ALA signal localisation in the autophagy compartment - specifically autolysosomes and lysosomes. Flow cytometry was employed to investigate whether autophagy enhancement through spermidine treatment (SPD) or nutrient deprivation/caloric restriction (CR) would enhance 5-ALA fluorescence signal generation. Indeed, SPD, CR and a combination of SPD/CR treatment significantly increased 5-ALA signal intensity, with a most robust increase in signal intensity observed in the combination treatment of SPD/CR. When using 3-D glioma spheroids to assess the effect of 5-ALA on cellular ultrastructure, we demonstrate that 5-ALA exposure leads to cytoplasmic disruption, vacuolarisation and large-scale mitophagy induction. These findings not only suggest a critical role for the autophagy compartment in 5-ALA engagement and signal generation but also point towards a novel and practically feasible approach to enhance 5-ALA fluorescence signal intensity. The findings may highlight that indeed autophagy control may serve as a promising avenue to promote an improved resection and GBM prognosis.
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Affiliation(s)
- Kim Fredericks
- Department of Physiological Sciences, Stellenbosch University, Stellenbosch, South Africa
| | - Jurgen Kriel
- Central Analytical Facility, Microscopy Unit, Stellenbosch University, South Africa
| | - Lize Engelbrecht
- Central Analytical Facility, Microscopy Unit, Stellenbosch University, South Africa
| | | | - Georg Widhalm
- Department of Neurosurgery, Medical University of Vienna, Vienna, Austria
| | - Brad Harrington
- Department of Neurosurgery, Stellenbosch University, Cape Town, South Africa
| | - Ian Vlok
- Department of Neurosurgery, Stellenbosch University, Cape Town, South Africa
| | - Ben Loos
- Department of Physiological Sciences, Stellenbosch University, Stellenbosch, South Africa
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3
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Uryu K, Soplop N, Sheahan TP, Catanese MT, Huynh C, Pena J, Boudreau N, Matei I, Kenific C, Hashimoto A, Hoshino A, Rice CM, Lyden D. Advancement in Cellular Topographic and Nanoparticle Capture Imaging by High Resolution Microscopy Incorporating a Freeze-Drying and Gaseous Nitrogen-based Approach. BIORXIV : THE PREPRINT SERVER FOR BIOLOGY 2023:2023.09.28.559906. [PMID: 37808646 PMCID: PMC10557753 DOI: 10.1101/2023.09.28.559906] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 10/10/2023]
Abstract
Scanning electron microscopy (SEM) offers an unparalleled view of the membrane topography of mammalian cells by using a conventional osmium (OsO4) and ethanol-based tissue preparation. However, conventional SEM methods limit optimal resolution due to ethanol and lipid interactions and interfere with visualization of fluorescent reporter proteins. Therefore, SEM correlative light and electron microscopy (CLEM) has been hindered by the adverse effects of ethanol and OsO4 on retention of fluorescence signals. To overcome this technological gap in achieving high-resolution SEM and retain fluorescent reporter signals, we developed a freeze-drying method with gaseous nitrogen (FDGN). We demonstrate that FDGN preserves cyto-architecture to allow visualization of detailed membrane topography while retaining fluorescent signals and that FDGN processing can be used in conjunction with a variety of high-resolution imaging systems to enable collection and validation of unique, high-quality data from these approaches. In particular, we show that FDGN coupled with high resolution microscopy provided detailed insight into viral or tumor-derived extracellular vesicle (TEV)-host cell interactions and may aid in designing new approaches to intervene during viral infection or to harness TEVs as therapeutic agents.
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Affiliation(s)
- Kunihiro Uryu
- Electron Microscopy Resource Center, The Rockefeller University, New York, NY 10065, USA
- Janelia Rearch Campsu, Ashuburn, Virginia, 20147, USA
| | - Nadine Soplop
- Electron Microscopy Resource Center, The Rockefeller University, New York, NY 10065, USA
| | - Timothy P. Sheahan
- Laboratory of Virology and Infectious Disease, The Rockefeller University, New York, NY 10065 USA
| | - Maria-Teresa Catanese
- Laboratory of Virology and Infectious Disease, The Rockefeller University, New York, NY 10065 USA
| | - Chuong Huynh
- Ion Microscopy Innovation Center, Zeiss Microscopy LLC, Peabody, MA 01960, USA
| | - John Pena
- Sonder Research X, Cornell University, Ithaca, NY 14853, USA
| | - Nancy Boudreau
- Children’s Cancer and Blood Foundation Laboratories, Departments of Pediatrics, and Cell and Developmental Biology, Drukier Institute for Children’s Health, Meyer Cancer Center, Weill Cornell Medicine, New York, NY 10065 USA
| | - Irina Matei
- Children’s Cancer and Blood Foundation Laboratories, Departments of Pediatrics, and Cell and Developmental Biology, Drukier Institute for Children’s Health, Meyer Cancer Center, Weill Cornell Medicine, New York, NY 10065 USA
| | - Candia Kenific
- Children’s Cancer and Blood Foundation Laboratories, Departments of Pediatrics, and Cell and Developmental Biology, Drukier Institute for Children’s Health, Meyer Cancer Center, Weill Cornell Medicine, New York, NY 10065 USA
| | - Ayako Hashimoto
- Children’s Cancer and Blood Foundation Laboratories, Departments of Pediatrics, and Cell and Developmental Biology, Drukier Institute for Children’s Health, Meyer Cancer Center, Weill Cornell Medicine, New York, NY 10065 USA
| | - Ayuko Hoshino
- Children’s Cancer and Blood Foundation Laboratories, Departments of Pediatrics, and Cell and Developmental Biology, Drukier Institute for Children’s Health, Meyer Cancer Center, Weill Cornell Medicine, New York, NY 10065 USA
| | - Charles M. Rice
- Laboratory of Virology and Infectious Disease, The Rockefeller University, New York, NY 10065 USA
| | - David Lyden
- Children’s Cancer and Blood Foundation Laboratories, Departments of Pediatrics, and Cell and Developmental Biology, Drukier Institute for Children’s Health, Meyer Cancer Center, Weill Cornell Medicine, New York, NY 10065 USA
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4
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Collinson LM. Developing a Cost-effective User-friendly Pipeline for Super-resolution Volume CLEM. MICROSCOPY AND MICROANALYSIS : THE OFFICIAL JOURNAL OF MICROSCOPY SOCIETY OF AMERICA, MICROBEAM ANALYSIS SOCIETY, MICROSCOPICAL SOCIETY OF CANADA 2023; 29:1212. [PMID: 37613421 DOI: 10.1093/micmic/ozad067.623] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 08/25/2023]
Affiliation(s)
- Lucy M Collinson
- Electron Microscopy Science Technology Platform, Francis Crick Institute, London, UK
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5
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de Wet S, Mangali A, Batt R, Kriel J, Vahrmeijer N, Niehaus D, Theart R, Loos B. The Highs and Lows of Memantine-An Autophagy and Mitophagy Inducing Agent That Protects Mitochondria. Cells 2023; 12:1726. [PMID: 37443760 PMCID: PMC10340721 DOI: 10.3390/cells12131726] [Citation(s) in RCA: 1] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/11/2023] [Revised: 06/16/2023] [Accepted: 06/21/2023] [Indexed: 07/15/2023] Open
Abstract
Memantine is an FDA-approved, non-competitive NMDA-receptor antagonist that has been shown to have mitochondrial protective effects, improve cell viability and enhance clearance of Aβ42 peptide. Currently, there are uncertainties regarding the precise molecular targets as well as the most favourable treatment concentrations of memantine. Here, we made use of an imaging-based approach to investigate the concentration-dependent effects of memantine on mitochondrial fission and fusion dynamics, autophagy and mitochondrial quality control using a neuronal model of CCCP-induced mitochondrial injury so as to better unpack how memantine aids in promoting neuronal health. GT1-7 murine hypothalamic cells were cultured under standard conditions, treated with a relatively high and low concentration (100 µM and 50 µM) of memantine for 48 h. Images were acquired using a Zeiss 780 PS1 platform. Utilising the mitochondrial event localiser (MEL), we demonstrated clear concentration-dependent effects of memantine causing a protective response to mitochondrial injury. Both concentrations maintained the mitochondrial network volume whilst the low concentration caused an increase in mitochondrial number as well as increased fission and fusion events following CCCP-induced injury. Additionally, we made use of a customised Python-based image processing and analysis pipeline to quantitatively assess memantine-dependent changes in the autophagosomal and lysosomal compartments. Our results revealed that memantine elicits a differential, concentration-dependent effect on autophagy pathway intermediates. Intriguingly, low but not high concentrations of memantine lead to the induction of mitophagy. Taken together, our findings have shown that memantine is able to protect the mitochondrial network by preserving its volume upon mitochondrial injury with high concentrations of memantine inducing macroautophagy, whereas low concentrations lead to the induction of mitophagy.
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Affiliation(s)
- Sholto de Wet
- Department of Physiological Sciences, Stellenbosch University, Stellenbosch 7600, South Africa
| | - Asandile Mangali
- Department of Physiological Sciences, Stellenbosch University, Stellenbosch 7600, South Africa
| | - Richard Batt
- Department of Electric and Electronic Engineering, Stellenbosch University, Stellenbosch 7600, South Africa
| | - Jurgen Kriel
- Microscopy Unit, Central Analytical Facility, Stellenbosch University, Stellenbosch 7600, South Africa
| | - Nicola Vahrmeijer
- Department of Physiological Sciences, Stellenbosch University, Stellenbosch 7600, South Africa
| | - Dana Niehaus
- Department of Psychiatry and Stikland Hospital, Faculty of Medicine and Health Sciences, Stellenbosch University, Cape Town 7530, South Africa
| | - Rensu Theart
- Department of Electric and Electronic Engineering, Stellenbosch University, Stellenbosch 7600, South Africa
| | - Ben Loos
- Department of Physiological Sciences, Stellenbosch University, Stellenbosch 7600, South Africa
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6
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Liv N, Fermie J, Ten Brink CBM, de Heus C, Klumperman J. Functional characterization of endo-lysosomal compartments by correlative live-cell and volume electron microscopy. Methods Cell Biol 2023; 177:301-326. [PMID: 37451771 DOI: 10.1016/bs.mcb.2022.12.022] [Citation(s) in RCA: 1] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 07/18/2023]
Abstract
Fluorescent biosensors are valuable tools to monitor protein activities and the functional state of organelles in live cells. However, the information provided by fluorescent microscopy (FM) is mostly limited in resolution and lacks ultrastructural context information. Protein activities are confined to organelle zones with a distinct membrane morphology, which can only be seen by electron microscopy (EM). EM, however, intrinsically lacks information on protein activities. The lack of methods to integrate these two imaging modalities has hampered understanding the functional organization of cellular organelles. Here we introduce "functional correlative microscopy" (functional CLEM) to directly infer functional information from live cells to EM with nanometer resolution. We label and visualize live cells with fluorescent biosensors after which they are processed for EM and imaged using a volume electron microscopy technique. Within a single dataset we correlate hundreds of fluorescent spots enabling quantitative analysis of the functional-ultrastructural data. We employ our method to monitor essential functional parameters of late endo-lysosomal compartments, i.e., pH, calcium, enzyme activities and cholesterol content. Our data reveal a steep functional difference in enzyme activity between late endosomes and lysosomes and unexpectedly high calcium levels in late endosomes. The presented CLEM workflow is compatible with a large repertoire of probes and paves the way for large scale functional studies of all types of cellular structures.
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Affiliation(s)
- Nalan Liv
- Center for Molecular Medicine, University Medical Center Utrecht, Utrecht University, Utrecht, The Netherlands.
| | - Job Fermie
- Center for Molecular Medicine, University Medical Center Utrecht, Utrecht University, Utrecht, The Netherlands; Molecular Biophysics, Debye Institute for Nanomaterials Science, Utrecht University, Utrecht, The Netherlands
| | - Corlinda B M Ten Brink
- Center for Molecular Medicine, University Medical Center Utrecht, Utrecht University, Utrecht, The Netherlands
| | - Cecilia de Heus
- Center for Molecular Medicine, University Medical Center Utrecht, Utrecht University, Utrecht, The Netherlands
| | - Judith Klumperman
- Center for Molecular Medicine, University Medical Center Utrecht, Utrecht University, Utrecht, The Netherlands
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7
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Pang S, Xu CS. Methods of enhanced FIB-SEM sample preparation and image acquisition. Methods Cell Biol 2023; 177:269-300. [PMID: 37451770 DOI: 10.1016/bs.mcb.2023.01.019] [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: 07/18/2023]
Abstract
The ability to view biomolecules in cells and measure changes in their structure, quantity, distribution, and interaction is fundamental to understanding biology. By coupling nano -scale resolution with meso and even macro scale volumes, the enhanced focused ion beam-scanning electron microscopy (FIB-SEM) pipeline has enabled numerous transformational discoveries in life science, many of which were major new landmarks in their fields. This pipeline consists of EM sample preparation, FIB-SEM sample preparation, FIB-SEM imaging, data alignment, and image analysis. While the EM sample preparation, data alignment, and image analysis are consistent with those from other volume Electron Microscopy (vEM) approaches, the enhanced FIB-SEM sample preparation and imaging are unique to the rest of comparable methods. We here illustrate the detailed methods of enhanced FIB-SEM sample preparation and image acquisition that have not been previously described. These methods can also be applied to the conventional FIB-SEM platforms for improved image acquisition quality and pipeline throughput.
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Affiliation(s)
- Song Pang
- Yale School of Medicine, New Haven, CT, United States.
| | - C Shan Xu
- Department of Cellular and Molecular Physiology, Yale School of Medicine, New Haven, CT, United States
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8
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Bélanger S, Berensmann H, Baena V, Duncan K, Meyers BC, Narayan K, Czymmek KJ. A versatile enhanced freeze-substitution protocol for volume electron microscopy. Front Cell Dev Biol 2022; 10:933376. [PMID: 36003147 PMCID: PMC9393620 DOI: 10.3389/fcell.2022.933376] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/30/2022] [Accepted: 07/08/2022] [Indexed: 11/18/2022] Open
Abstract
Volume electron microscopy, a powerful approach to generate large three-dimensional cell and tissue volumes at electron microscopy resolutions, is rapidly becoming a routine tool for understanding fundamental and applied biological questions. One of the enabling factors for its adoption has been the development of conventional fixation protocols with improved heavy metal staining. However, freeze-substitution with organic solvent-based fixation and staining has not realized the same level of benefit. Here, we report a straightforward approach including osmium tetroxide, acetone and up to 3% water substitution fluid (compatible with traditional or fast freeze-substitution protocols), warm-up and transition from organic solvent to aqueous 2% osmium tetroxide. Once fully hydrated, samples were processed in aqueous based potassium ferrocyanide, thiocarbohydrazide, osmium tetroxide, uranyl acetate and lead acetate before resin infiltration and polymerization. We observed a consistent and substantial increase in heavy metal staining across diverse and difficult-to-fix test organisms and tissue types, including plant tissues (Hordeum vulgare), nematode (Caenorhabditis elegans) and yeast (Saccharomyces cerevisiae). Our approach opens new possibilities to combine the benefits of cryo-preservation with enhanced contrast for volume electron microscopy in diverse organisms.
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Affiliation(s)
| | - Heather Berensmann
- Center for Molecular Microscopy, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD, United States
- Cancer Research Technology Program, Frederick National Laboratory for Cancer Research, Frederick, MD, United States
| | - Valentina Baena
- Center for Molecular Microscopy, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD, United States
- Cancer Research Technology Program, Frederick National Laboratory for Cancer Research, Frederick, MD, United States
| | - Keith Duncan
- Donald Danforth Plant Science Center, Saint Louis, MO, United States
| | - Blake C. Meyers
- Donald Danforth Plant Science Center, Saint Louis, MO, United States
- Division of Plant Science and Technology, University of Missouri–Columbia, Columbia, MO, United States
| | - Kedar Narayan
- Center for Molecular Microscopy, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD, United States
- Cancer Research Technology Program, Frederick National Laboratory for Cancer Research, Frederick, MD, United States
| | - Kirk J. Czymmek
- Donald Danforth Plant Science Center, Saint Louis, MO, United States
- Advanced Bioimaging Laboratory, Donald Danforth Plant Science Center, Saint Louis, MO, United States
- *Correspondence: Kirk J. Czymmek,
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9
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Peddie CJ, Genoud C, Kreshuk A, Meechan K, Micheva KD, Narayan K, Pape C, Parton RG, Schieber NL, Schwab Y, Titze B, Verkade P, Aubrey A, Collinson LM. Volume electron microscopy. NATURE REVIEWS. METHODS PRIMERS 2022; 2:51. [PMID: 37409324 PMCID: PMC7614724 DOI: 10.1038/s43586-022-00131-9] [Citation(s) in RCA: 35] [Impact Index Per Article: 17.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Accepted: 05/10/2022] [Indexed: 07/07/2023]
Abstract
Life exists in three dimensions, but until the turn of the century most electron microscopy methods provided only 2D image data. Recently, electron microscopy techniques capable of delving deep into the structure of cells and tissues have emerged, collectively called volume electron microscopy (vEM). Developments in vEM have been dubbed a quiet revolution as the field evolved from established transmission and scanning electron microscopy techniques, so early publications largely focused on the bioscience applications rather than the underlying technological breakthroughs. However, with an explosion in the uptake of vEM across the biosciences and fast-paced advances in volume, resolution, throughput and ease of use, it is timely to introduce the field to new audiences. In this Primer, we introduce the different vEM imaging modalities, the specialized sample processing and image analysis pipelines that accompany each modality and the types of information revealed in the data. We showcase key applications in the biosciences where vEM has helped make breakthrough discoveries and consider limitations and future directions. We aim to show new users how vEM can support discovery science in their own research fields and inspire broader uptake of the technology, finally allowing its full adoption into mainstream biological imaging.
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Affiliation(s)
- Christopher J. Peddie
- Electron Microscopy Science Technology Platform, The Francis Crick Institute, London, UK
| | - Christel Genoud
- Electron Microscopy Facility, Faculty of Biology and Medicine, University of Lausanne, Lausanne, Switzerland
| | - Anna Kreshuk
- Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Kimberly Meechan
- Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
- Present address: Faculty of Biosciences, Heidelberg University, Heidelberg, Germany
| | - Kristina D. Micheva
- Department of Molecular and Cellular Physiology, Stanford University, Palo Alto, CA, USA
| | - Kedar Narayan
- Center for Molecular Microscopy, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA
- Cancer Research Technology Program, Frederick National Laboratory for Cancer Research, Frederick, MD, USA
| | - Constantin Pape
- Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Robert G. Parton
- The Institute for Molecular Bioscience, The University of Queensland, Brisbane, Queensland, Australia
- Centre for Microscopy and Microanalysis, The University of Queensland, Brisbane, Queensland, Australia
| | - Nicole L. Schieber
- Centre for Microscopy and Microanalysis, The University of Queensland, Brisbane, Queensland, Australia
| | - Yannick Schwab
- Cell Biology and Biophysics Unit/ Electron Microscopy Core Facility, European Molecular Biology Laboratory, Heidelberg, Germany
| | | | - Paul Verkade
- School of Biochemistry, University of Bristol, Bristol, UK
| | - Aubrey Aubrey
- Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, VA, USA
| | - Lucy M. Collinson
- Electron Microscopy Science Technology Platform, The Francis Crick Institute, London, UK
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10
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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.
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Affiliation(s)
| | - Miriam S. Lucas
- Scientific Center for Light and Electron Microscopy (ScopeM), ETH Zurich, Zurich, Switzerland
- *Correspondence: Miriam S. Lucas,
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11
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van den Dries K, Fransen J, Cambi A. Fluorescence CLEM in biology: historic developments and current super-resolution applications. FEBS Lett 2022; 596:2486-2496. [PMID: 35674424 DOI: 10.1002/1873-3468.14421] [Citation(s) in RCA: 8] [Impact Index Per Article: 4.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/14/2022] [Revised: 05/24/2022] [Accepted: 05/25/2022] [Indexed: 11/12/2022]
Abstract
Correlative light and electron microscopy (CLEM) is a powerful imaging approach that allows the direct correlation of information obtained on a light and an electron microscope. There is a growing interest in the application of CLEM in biology, mainly attributable to technical advances in field of fluorescence microscopy in the past two decades. In this review, we summarize the important developments in CLEM for biological applications, focusing on the combination of fluorescence microscopy and electron microscopy. We first provide a brief overview of the early days of fluorescence CLEM usage starting with the initial rise in the late 1970s and the subsequent optimization of CLEM workflows during the following two decades. Next, we describe how the engineering of fluorescent proteins and the development of super-resolution fluorescence microscopy have significantly renewed the interest in CLEM resulting in the present application of fluorescence CLEM in many different areas of cellular and molecular biology. Lastly, we present the promises and challenges for the future of fluorescence CLEM discussing novel workflows, probe development and quantification possibilities.
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Affiliation(s)
- Koen van den Dries
- Department of Cell Biology, Radboud Institute for Molecular Life Sciences, Radboudumc, Nijmegen, The Netherlands
| | - Jack Fransen
- Department of Cell Biology, Radboud Institute for Molecular Life Sciences, Radboudumc, Nijmegen, The Netherlands.,Microscopic Imaging Center, Radboud Institute for Molecular Life Sciences, Radboudumc, Nijmegen, The Netherlands
| | - Alessandra Cambi
- Department of Cell Biology, Radboud Institute for Molecular Life Sciences, Radboudumc, Nijmegen, The Netherlands
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12
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Khare D, Chandwadkar P, Acharya C. Structural Analysis of Gliding Motility of a Bacteroidetes Bacterium by Correlative Light and Scanning Electron Microscopy (CLSEM). MICROSCOPY AND MICROANALYSIS : THE OFFICIAL JOURNAL OF MICROSCOPY SOCIETY OF AMERICA, MICROBEAM ANALYSIS SOCIETY, MICROSCOPICAL SOCIETY OF CANADA 2022; 28:1-7. [PMID: 35105420 DOI: 10.1017/s1431927622000095] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/14/2023]
Abstract
The members of the Bacteroidetes phylum move on surfaces by gliding motility in the absence of external motility appendages, leading to the formation of spreading colonies. Here, the structural features of the spreading colony were assessed in a uranium-tolerant Bacteroidetes bacterium, Chryseobacterium sp. strain PMSZPI, by using correlative light and scanning electron microscopy (CLSEM). We developed a simple and convenient workflow for CLSEM using a shuttle and find software module and a correlative sample holding slide designed to transport samples between the light/fluorescence microscope (LM/FM) and the scanning electron microscope (SEM) to image spreading colony edges. The datasets from the CLSEM studies allowed convenient examination of the colonial organization by LM/FM followed by ultrastructural analysis by SEM. The regions of interest (ROIs) of the spreading colony edges that were observed in LM/FM in the absence and presence of uranium could be re-identified in the SEM quickly without prolonged searching. Perfect correlation between LM and SEM could be achieved with minimum preparation steps. Subsequently, imaging of the correlated regions was done at higher resolution in SEM to obtain more comprehensive information. We further showed the association of uranium with the gliding PMSZPI cells by energy-dispersive X-ray spectroscopy (EDS) attached to SEM.
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Affiliation(s)
- Devanshi Khare
- Molecular Biology Division, Bhabha Atomic Research Centre, Trombay, Mumbai400085, India
- Homi Bhabha National Institute, Anushakti Nagar, Mumbai400094, India
| | - Pallavi Chandwadkar
- Molecular Biology Division, Bhabha Atomic Research Centre, Trombay, Mumbai400085, India
| | - Celin Acharya
- Molecular Biology Division, Bhabha Atomic Research Centre, Trombay, Mumbai400085, India
- Homi Bhabha National Institute, Anushakti Nagar, Mumbai400094, India
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13
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Lane R, Wolters AHG, Giepmans BNG, Hoogenboom JP. Integrated Array Tomography for 3D Correlative Light and Electron Microscopy. Front Mol Biosci 2022; 8:822232. [PMID: 35127826 PMCID: PMC8809480 DOI: 10.3389/fmolb.2021.822232] [Citation(s) in RCA: 4] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/25/2021] [Accepted: 12/15/2021] [Indexed: 12/22/2022] Open
Abstract
Volume electron microscopy (EM) of biological systems has grown exponentially in recent years due to innovative large-scale imaging approaches. As a standalone imaging method, however, large-scale EM typically has two major limitations: slow rates of acquisition and the difficulty to provide targeted biological information. We developed a 3D image acquisition and reconstruction pipeline that overcomes both of these limitations by using a widefield fluorescence microscope integrated inside of a scanning electron microscope. The workflow consists of acquiring large field of view fluorescence microscopy (FM) images, which guide to regions of interest for successive EM (integrated correlative light and electron microscopy). High precision EM-FM overlay is achieved using cathodoluminescent markers. We conduct a proof-of-concept of our integrated workflow on immunolabelled serial sections of tissues. Acquisitions are limited to regions containing biological targets, expediting total acquisition times and reducing the burden of excess data by tens or hundreds of GBs.
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Affiliation(s)
- Ryan Lane
- Imaging Physics, Delft University of Technology, Delft, Netherlands
| | - Anouk H. G. Wolters
- Department of Biomedical Sciences of Cells and Systems, University Groningen, University Medical Center Groningen, Groningen, Netherlands
| | - Ben N. G. Giepmans
- Department of Biomedical Sciences of Cells and Systems, University Groningen, University Medical Center Groningen, Groningen, Netherlands
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14
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Baatsen P, Gabarre S, Vints K, Wouters R, Vandael D, Goodchild R, Munck S, Gounko NV. Preservation of Fluorescence Signal and Imaging Optimization for Integrated Light and Electron Microscopy. Front Cell Dev Biol 2022; 9:737621. [PMID: 34977003 PMCID: PMC8715528 DOI: 10.3389/fcell.2021.737621] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/07/2021] [Accepted: 11/25/2021] [Indexed: 11/13/2022] Open
Abstract
Life science research often needs to define where molecules are located within the complex environment of a cell or tissue. Genetically encoded fluorescent proteins and or fluorescence affinity-labeling are the go-to methods. Although recent fluorescent microscopy methods can provide localization of fluorescent molecules with relatively high resolution, an ultrastructural context is missing. This is solved by imaging a region of interest with correlative light and electron microscopy (CLEM). We have adopted a protocol that preserves both genetically-encoded and antibody-derived fluorescent signals in resin-embedded cell and tissue samples and provides high-resolution electron microscopy imaging of the same thin section. This method is particularly suitable for dedicated CLEM instruments that combine fluorescence and electron microscopy optics. In addition, we optimized scanning EM imaging parameters for samples of varying thicknesses. These protocols will enable rapid acquisition of CLEM information from samples and can be adapted for three-dimensional EM.
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Affiliation(s)
- Pieter Baatsen
- VIB-KU Leuven Center for Brain and Disease Research, Electron Microscopy Platform and VIB-Bioimaging Core, Leuven, Belgium.,KU Leuven Department of Neurosciences, Leuven Brain Institute, Leuven, Belgium
| | - Sergio Gabarre
- VIB-KU Leuven Center for Brain and Disease Research, Electron Microscopy Platform and VIB-Bioimaging Core, Leuven, Belgium.,KU Leuven Department of Neurosciences, Leuven Brain Institute, Leuven, Belgium.,VIB-KU Leuven Center for Brain and Disease Research, Light Microscopy Expertise Unit and VIB Bioimaging Core, Leuven, Belgium
| | - Katlijn Vints
- VIB-KU Leuven Center for Brain and Disease Research, Electron Microscopy Platform and VIB-Bioimaging Core, Leuven, Belgium.,KU Leuven Department of Neurosciences, Leuven Brain Institute, Leuven, Belgium
| | - Rosanne Wouters
- KU Leuven Department of Neurosciences, Leuven Brain Institute, Leuven, Belgium.,VIB-KU Leuven Center for Brain and Disease Research, Laboratory for Membrane Trafficking, Leuven, Belgium
| | - Dorien Vandael
- VIB-KU Leuven Center for Brain and Disease Research, Electron Microscopy Platform and VIB-Bioimaging Core, Leuven, Belgium.,KU Leuven Department of Neurosciences, Leuven Brain Institute, Leuven, Belgium
| | - Rose Goodchild
- KU Leuven Department of Neurosciences, Leuven Brain Institute, Leuven, Belgium.,VIB-KU Leuven Center for Brain and Disease Research, Laboratory for Dystonia Research, Leuven, Belgium
| | - Sebastian Munck
- KU Leuven Department of Neurosciences, Leuven Brain Institute, Leuven, Belgium.,VIB-KU Leuven Center for Brain and Disease Research, Light Microscopy Expertise Unit and VIB Bioimaging Core, Leuven, Belgium
| | - Natalia V Gounko
- VIB-KU Leuven Center for Brain and Disease Research, Electron Microscopy Platform and VIB-Bioimaging Core, Leuven, Belgium.,KU Leuven Department of Neurosciences, Leuven Brain Institute, Leuven, Belgium
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15
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Richardson AC, Fišerová J, Goldberg MW. NPC Structure in Model Organisms: Transmission Electron Microscopy and Immunogold Labeling Using High-Pressure Freezing/Freeze Substitution of Yeast, Worms, and Plants. Methods Mol Biol 2022; 2502:439-459. [PMID: 35412255 DOI: 10.1007/978-1-0716-2337-4_28] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 06/14/2023]
Abstract
The nuclear pore complex (NPC) is a large elaborate structure embedded within the nuclear envelope, and intimately linked to the cytoskeleton, nucleoskeleton, and chromatin. Many different cargoes pass through its central channel and along the membrane at its periphery. The NPC is dismantled and reassembly, fully or partially, every cell cycle. In post-mitotic cells it consists of a combination of hyper-stable and highly dynamic proteins. Because of its size, dynamics, heterogeneity and integration, it is not possible to understand its structure and molecular function by any one, or even several, methods. For decades, and to this day, thin section transmission electron microscopy (TEM) has been a central tool for understanding the NPC, its associations, dynamics and role in transport as it can uniquely answer questions concerning fine structural detail within a cellular context. Using immunogold labeling specific components can also be identified within the ultrastructural context. Model organisms such as Saccharomyces cerevisiae are also central to NPC studies but have not been used extensively in structural work. This is because the cell wall presents difficulties with structural preservation and processing for TEM. In recent years, high-pressure freezing and freeze substitution have overcome these problems, as well as opened up methods to combine immunogold labeling with detailed structural analysis. Other model organisms such as the worm Caenorhabditis elegans and the plant Arabidopsis thaliana have been underused for similar reasons, but with similar solutions, which we present here. There are also many advantages to using these methods, adapted for use in mammalian systems, due to the instant nature of the initial fixation, to capture rapid processes such as nuclear transport, and preservation of dynamic membranes.
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Affiliation(s)
| | - Jindřiška Fišerová
- Department of Biology of the Cell Nucleus, Institute of Molecular Genetics AS CR, Prague, Czech Republic
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16
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Capturing the third dimension in drug discovery: Spatially-resolved tools for interrogation of complex 3D cell models. Biotechnol Adv 2021; 55:107883. [PMID: 34875362 DOI: 10.1016/j.biotechadv.2021.107883] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.7] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/15/2021] [Revised: 11/22/2021] [Accepted: 11/30/2021] [Indexed: 02/07/2023]
Abstract
Advanced three-dimensional (3D) cell models have proven to be capable of depicting architectural and microenvironmental features of several tissues. By providing data of higher physiological and pathophysiological relevance, 3D cell models have been contributing to a better understanding of human development, pathology onset and progression mechanisms, as well as for 3D cell-based assays for drug discovery. Nonetheless, the characterization and interrogation of these tissue-like structures pose major challenges on the conventional analytical methods, pushing the development of spatially-resolved technologies. Herein, we review recent advances and pioneering technologies suitable for the interrogation of multicellular 3D models, while capable of retaining biological spatial information. We focused on imaging technologies and omics tools, namely transcriptomics, proteomics and metabolomics. The advantages and shortcomings of these novel methodologies are discussed, alongside the opportunities to intertwine data from the different tools.
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17
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Parlanti P, Cappello V. Microscopes, tools, probes, and protocols: A guide in the route of correlative microscopy for biomedical investigation. Micron 2021; 152:103182. [PMID: 34801960 DOI: 10.1016/j.micron.2021.103182] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/28/2021] [Revised: 11/12/2021] [Accepted: 11/14/2021] [Indexed: 12/11/2022]
Abstract
In the last decades, the advancements of microscopes technology, together with the development of new imaging approaches, are trying to address some biological questions that have been unresolved in the past: the need to combine in the same analysis temporal, functional and morphological information on the biological sample has become pressing. For this reason, the use of correlative microscopy, in which two or more imaging techniques are combined in the same analysis, is getting increasingly widespread. In fact, correlative microscopy can overcome limitations of a single imaging method, giving access to a larger amount of information from the same specimen. However, correlative microscopy can be challenging, and appropriate protocols for sample preparation and imaging methods must be selected. Here we review the state of the art of correlating electron microscopy with different imaging methods, focusing on sample preparation, tools, and labeling methods, with the aim to provide a comprehensive guide for those scientists who are approaching the field of correlative methods.
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Affiliation(s)
- Paola Parlanti
- Istituto Italiano di Tecnologia, Center for Materials Interfaces, Electron Crystallography, Viale Rinaldo Piaggio 34, I-56025, Pontedera (PI), Italy.
| | - Valentina Cappello
- Istituto Italiano di Tecnologia, Center for Materials Interfaces, Electron Crystallography, Viale Rinaldo Piaggio 34, I-56025, Pontedera (PI), Italy.
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18
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Partridge TA. Enhancing Interrogation of Skeletal Muscle Samples for Informative Quantitative Data. J Neuromuscul Dis 2021; 8:S257-S269. [PMID: 34511511 PMCID: PMC8673506 DOI: 10.3233/jnd-210736] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/17/2022]
Abstract
Careful quantitative analysis of histological preparations of muscle samples is crucial to accurate investigation of myopathies in man and of interpretation of data from animals subjected to experimental or potentially therapeutic treatments. Protocols for measuring cell numbers are subject to problems arising from biases associated with preparative and analytical techniques. Prominent among these is the effect of polarized structure of skeletal muscle on sampling bias. It is also common in this tissue to collect data as ratios to convenient reference dominators, the fundamental bases of which are ill-defined, or unrecognized or not accurately assessable. Use of such 'floating' denominators raises a barrier to estimation of the absolute values that assume practical importance in medical research, where accurate comparison between different scenarios in different species is essential to the aim of translating preclinical research findings in animal models to clinical utility in Homo sapiens.This review identifies some of the underappreciated problems with current morphometric practice, some of which are exacerbated in skeletal muscle, and evaluates the extent of their intrusiveness into the of building an objective, accurate, picture of the structure of the muscle sample. It also contains recommendations for eliminating or at least minimizing these problems. Principal among these, would be the use of stereological procedures to avoid the substantial counting biases arising from inter-procedure differences in object size and section thickness.Attention is also drawn to the distortions of interpretation arising from use of undefined or inappropriate denominators.
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Affiliation(s)
- Terence A Partridge
- Professor of Integrative Systemic Biology, George Washington University, Washington DC.,Honorary Professor, Institute of Child Health, University College London
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19
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Ronchi P, Mizzon G, Machado P, D’Imprima E, Best BT, Cassella L, Schnorrenberg S, Montero MG, Jechlinger M, Ephrussi A, Leptin M, Mahamid J, Schwab Y. High-precision targeting workflow for volume electron microscopy. J Cell Biol 2021; 220:e202104069. [PMID: 34160561 PMCID: PMC8225610 DOI: 10.1083/jcb.202104069] [Citation(s) in RCA: 24] [Impact Index Per Article: 8.0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/16/2021] [Revised: 05/27/2021] [Accepted: 06/06/2021] [Indexed: 02/07/2023] Open
Abstract
Cells are 3D objects. Therefore, volume EM (vEM) is often crucial for correct interpretation of ultrastructural data. Today, scanning EM (SEM) methods such as focused ion beam (FIB)-SEM are frequently used for vEM analyses. While they allow automated data acquisition, precise targeting of volumes of interest within a large sample remains challenging. Here, we provide a workflow to target FIB-SEM acquisition of fluorescently labeled cells or subcellular structures with micrometer precision. The strategy relies on fluorescence preservation during sample preparation and targeted trimming guided by confocal maps of the fluorescence signal in the resin block. Laser branding is used to create landmarks on the block surface to position the FIB-SEM acquisition. Using this method, we acquired volumes of specific single cells within large tissues such as 3D cultures of mouse mammary gland organoids, tracheal terminal cells in Drosophila melanogaster larvae, and ovarian follicular cells in adult Drosophila, discovering ultrastructural details that could not be appreciated before.
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Affiliation(s)
- Paolo Ronchi
- Electron Microscopy Core Facility, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Giulia Mizzon
- Electron Microscopy Core Facility, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Pedro Machado
- Electron Microscopy Core Facility, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Edoardo D’Imprima
- Structural and Computational Biology Unit, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Benedikt T. Best
- Directors’ Research, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Lucia Cassella
- Developmental Biology Unit, European Molecular Biology Laboratory, Heidelberg, Germany
- Faculty of Biosciences, Heidelberg University, Heidelberg, Germany
| | - Sebastian Schnorrenberg
- Advanced Light Microscopy Facility, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Marta G. Montero
- Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Martin Jechlinger
- Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Anne Ephrussi
- Developmental Biology Unit, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Maria Leptin
- Directors’ Research, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Julia Mahamid
- Structural and Computational Biology Unit, European Molecular Biology Laboratory, Heidelberg, Germany
| | - Yannick Schwab
- Electron Microscopy Core Facility, European Molecular Biology Laboratory, Heidelberg, Germany
- Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
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20
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Dorone Y, Boeynaems S, Flores E, Jin B, Hateley S, Bossi F, Lazarus E, Pennington JG, Michiels E, De Decker M, Vints K, Baatsen P, Bassel GW, Otegui MS, Holehouse AS, Exposito-Alonso M, Sukenik S, Gitler AD, Rhee SY. A prion-like protein regulator of seed germination undergoes hydration-dependent phase separation. Cell 2021; 184:4284-4298.e27. [PMID: 34233164 PMCID: PMC8513799 DOI: 10.1016/j.cell.2021.06.009] [Citation(s) in RCA: 87] [Impact Index Per Article: 29.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/16/2020] [Revised: 03/22/2021] [Accepted: 06/04/2021] [Indexed: 12/22/2022]
Abstract
Many organisms evolved strategies to survive desiccation. Plant seeds protect dehydrated embryos from various stressors and can lay dormant for millennia. Hydration is the key trigger to initiate germination, but the mechanism by which seeds sense water remains unresolved. We identified an uncharacterized Arabidopsis thaliana prion-like protein we named FLOE1, which phase separates upon hydration and allows the embryo to sense water stress. We demonstrate that biophysical states of FLOE1 condensates modulate its biological function in vivo in suppressing seed germination under unfavorable environments. We find intragenic, intraspecific, and interspecific natural variation in FLOE1 expression and phase separation and show that intragenic variation is associated with adaptive germination strategies in natural populations. This combination of molecular, organismal, and ecological studies uncovers FLOE1 as a tunable environmental sensor with direct implications for the design of drought-resistant crops, in the face of climate change.
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Affiliation(s)
- Yanniv Dorone
- Department of Plant Biology, Carnegie Institution for Science, Stanford, CA 94305, USA; Department of Biology, Stanford University, Stanford, CA 94305, USA
| | - Steven Boeynaems
- Department of Genetics, Stanford University School of Medicine, Stanford, CA 94305, USA
| | - Eduardo Flores
- Department of Chemistry and Chemical Biology, UC Merced, Merced, CA 95340, USA
| | - Benjamin Jin
- Department of Plant Biology, Carnegie Institution for Science, Stanford, CA 94305, USA
| | - Shannon Hateley
- Department of Plant Biology, Carnegie Institution for Science, Stanford, CA 94305, USA
| | - Flavia Bossi
- Department of Plant Biology, Carnegie Institution for Science, Stanford, CA 94305, USA
| | - Elena Lazarus
- Department of Plant Biology, Carnegie Institution for Science, Stanford, CA 94305, USA
| | - Janice G Pennington
- Center for Quantitative Cell Imaging, University of Wisconsin, Madison, WI 53706, USA
| | - Emiel Michiels
- EM-platform@VIB Bio Imaging Core and VIB Center for Brain and Disease Research, KU Leuven, 3000 Leuven, Belgium; Switch Laboratory, Department of Cellular and Molecular Medicine, KU Leuven, 3000 Leuven, Belgium
| | - Mathias De Decker
- EM-platform@VIB Bio Imaging Core and VIB Center for Brain and Disease Research, KU Leuven, 3000 Leuven, Belgium; KU Leuven - University of Leuven, Department of Neurosciences, Experimental Neurology, and Leuven Brain Institute (LBI), 3000 Leuven, Belgium
| | - Katlijn Vints
- EM-platform@VIB Bio Imaging Core and VIB Center for Brain and Disease Research, KU Leuven, 3000 Leuven, Belgium
| | - Pieter Baatsen
- EM-platform@VIB Bio Imaging Core and VIB Center for Brain and Disease Research, KU Leuven, 3000 Leuven, Belgium
| | - George W Bassel
- School of Life Sciences, University of Warwick, Coventry CV4 7AL, UK
| | - Marisa S Otegui
- Center for Quantitative Cell Imaging, University of Wisconsin, Madison, WI 53706, USA; Department of Botany, University of Wisconsin, Madison, WI 53706, USA
| | - Alex S Holehouse
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO 63110, USA; Center for Science and Engineering of Living Systems (CSELS), Washington University in St. Louis, St. Louis, MO 63130, USA
| | - Moises Exposito-Alonso
- Department of Plant Biology, Carnegie Institution for Science, Stanford, CA 94305, USA; Department of Biology, Stanford University, Stanford, CA 94305, USA
| | - Shahar Sukenik
- Department of Chemistry and Chemical Biology, UC Merced, Merced, CA 95340, USA
| | - Aaron D Gitler
- Department of Genetics, Stanford University School of Medicine, Stanford, CA 94305, USA.
| | - Seung Y Rhee
- Department of Plant Biology, Carnegie Institution for Science, Stanford, CA 94305, USA.
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21
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Gabarre S, Vernaillen F, Baatsen P, Vints K, Cawthorne C, Boeynaems S, Michiels E, Vandael D, Gounko NV, Munck S. A workflow for streamlined acquisition and correlation of serial regions of interest in array tomography. BMC Biol 2021; 19:152. [PMID: 34330271 PMCID: PMC8323292 DOI: 10.1186/s12915-021-01072-7] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/04/2020] [Accepted: 06/14/2021] [Indexed: 11/20/2022] Open
Abstract
Background Array tomography (AT) is a high-resolution imaging method to resolve fine details at the organelle level and has the advantage that it can provide 3D volumes to show the tissue context. AT can be carried out in a correlative way, combing light and electron microscopy (LM, EM) techniques. However, the correlation between modalities can be a challenge and delineating specific regions of interest in consecutive sections can be time-consuming. Integrated light and electron microscopes (iLEMs) offer the possibility to provide well-correlated images and may pose an ideal solution for correlative AT. Here, we report a workflow to automate navigation between regions of interest. Results We use a targeted approach that allows imaging specific tissue features, like organelles, cell processes, and nuclei at different scales to enable fast, directly correlated in situ AT using an integrated light and electron microscope (iLEM-AT). Our workflow is based on the detection of section boundaries on an initial transmitted light acquisition that serves as a reference space to compensate for changes in shape between sections, and we apply a stepwise refinement of localizations as the magnification increases from LM to EM. With minimal user interaction, this enables autonomous and speedy acquisition of regions containing cells and cellular organelles of interest correlated across different magnifications for LM and EM modalities, providing a more efficient way to obtain 3D images. We provide a proof of concept of our approach and the developed software tools using both Golgi neuronal impregnation staining and fluorescently labeled protein condensates in cells. Conclusions Our method facilitates tracing and reconstructing cellular structures over multiple sections, is targeted at high resolution ILEMs, and can be integrated into existing devices, both commercial and custom-built systems. Supplementary Information The online version contains supplementary material available at 10.1186/s12915-021-01072-7.
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Affiliation(s)
- Sergio Gabarre
- VIB-KU Leuven Center for Brain & Disease Research, Electron Microscopy Platform & VIB BioImaging Core, O&N5 Herestraat 49 box 602, 3000, Leuven, Belgium.,KU Leuven Department of Neurosciences, Leuven Brain Institute, O&N5 Herestraat 49 box 602, 3000, Leuven, Belgium.,VIB-KU Leuven Center for Brain & Disease Research, Light Microscopy Expertise Unit & VIB BioImaging Core, O&N5 Herestraat 49 box 602, 3000, Leuven, Belgium
| | - Frank Vernaillen
- VIB BioInformatics Core, Technologiepark 75, 9052, Ghent, Belgium
| | - Pieter Baatsen
- VIB-KU Leuven Center for Brain & Disease Research, Electron Microscopy Platform & VIB BioImaging Core, O&N5 Herestraat 49 box 602, 3000, Leuven, Belgium.,KU Leuven Department of Neurosciences, Leuven Brain Institute, O&N5 Herestraat 49 box 602, 3000, Leuven, Belgium
| | - Katlijn Vints
- VIB-KU Leuven Center for Brain & Disease Research, Electron Microscopy Platform & VIB BioImaging Core, O&N5 Herestraat 49 box 602, 3000, Leuven, Belgium.,KU Leuven Department of Neurosciences, Leuven Brain Institute, O&N5 Herestraat 49 box 602, 3000, Leuven, Belgium
| | | | - Steven Boeynaems
- Department of Genetics, Stanford University School of Medicine, Stanford, California, 94305, USA
| | - Emiel Michiels
- VIB Center for Brain and Disease Research, 3000, Leuven, Belgium.,Switch Laboratory, Department of Cellular and Molecular Medicine, KU Leuven, 3000, Leuven, Belgium
| | - Dorien Vandael
- VIB-KU Leuven Center for Brain & Disease Research, Electron Microscopy Platform & VIB BioImaging Core, O&N5 Herestraat 49 box 602, 3000, Leuven, Belgium.,KU Leuven Department of Neurosciences, Leuven Brain Institute, O&N5 Herestraat 49 box 602, 3000, Leuven, Belgium
| | - Natalia V Gounko
- VIB-KU Leuven Center for Brain & Disease Research, Electron Microscopy Platform & VIB BioImaging Core, O&N5 Herestraat 49 box 602, 3000, Leuven, Belgium. .,KU Leuven Department of Neurosciences, Leuven Brain Institute, O&N5 Herestraat 49 box 602, 3000, Leuven, Belgium.
| | - Sebastian Munck
- KU Leuven Department of Neurosciences, Leuven Brain Institute, O&N5 Herestraat 49 box 602, 3000, Leuven, Belgium. .,VIB-KU Leuven Center for Brain & Disease Research, Light Microscopy Expertise Unit & VIB BioImaging Core, O&N5 Herestraat 49 box 602, 3000, Leuven, Belgium.
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22
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Invadopodia Structure in 3D Environment Resolved by Near-Infrared Branding Protocol Combining Correlative Confocal and FIB-SEM Microscopy. Int J Mol Sci 2021; 22:ijms22157805. [PMID: 34360570 PMCID: PMC8346040 DOI: 10.3390/ijms22157805] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Key Words] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/27/2021] [Revised: 07/02/2021] [Accepted: 07/19/2021] [Indexed: 01/18/2023] Open
Abstract
Cancer cell invasion through tissue barriers is the intrinsic feature of metastasis, the most life-threatening aspect of cancer. Detailed observation and analysis of cancer cell behaviour in a 3D environment is essential for a full understanding of the mechanisms of cancer cell invasion. The inherent limits of optical microscopy resolution do not allow to for in-depth observation of intracellular structures, such as invadopodia of invading cancer cells. The required resolution can be achieved using electron microscopy techniques such as FIB-SEM. However, visualising cells in a 3D matrix using FIB-SEM is challenging due to difficulties with localisation of a specific cell deep within the resin block. We have developed a new protocol based on the near-infrared branding (NIRB) procedure that extends the pattern from the surface grid deep inside the resin. This 3D burned pattern allows for precise trimming followed by targeted 3D FIB-SEM. Here we present detailed 3D CLEM results combining confocal and FIB-SEM imaging of cancer cell invadopodia that extend deep into the collagen meshwork.
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23
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Lane R, Vos Y, Wolters AHG, Kessel LV, Chen SE, Liv N, Klumperman J, Giepmans BNG, Hoogenboom JP. Optimization of negative stage bias potential for faster imaging in large-scale electron microscopy. JOURNAL OF STRUCTURAL BIOLOGY-X 2021; 5:100046. [PMID: 33763642 PMCID: PMC7973379 DOI: 10.1016/j.yjsbx.2021.100046] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Subscribe] [Scholar Register] [Received: 09/04/2020] [Revised: 12/17/2020] [Accepted: 01/27/2021] [Indexed: 11/24/2022]
Abstract
The use of a negative bias potential was empirically optimized for tissue imaging with SEM. Optimized bias potential leads to a factor 20 increase in imaging speeds as well as an order of magnitude improvement to SNR. SNR increase results from a combination of BSE acceleration and detector response. Similar increases to SNR can be obtained when a magnetic immersion field is combined with a negative bias potential. Stage bias can be applied within an integrated fluorescence and electron microscope allowing for fast correlative imaging of tissue sections.
Large-scale electron microscopy (EM) allows analysis of both tissues and macromolecules in a semi-automated manner, but acquisition rate forms a bottleneck. We reasoned that a negative bias potential may be used to enhance signal collection, allowing shorter dwell times and thus increasing imaging speed. Negative bias potential has previously been used to tune penetration depth in block-face imaging. However, optimization of negative bias potential for application in thin section imaging will be needed prior to routine use and application in large-scale EM. Here, we present negative bias potential optimized through a combination of simulations and empirical measurements. We find that the use of a negative bias potential generally results in improvement of image quality and signal-to-noise ratio (SNR). The extent of these improvements depends on the presence and strength of a magnetic immersion field. Maintaining other imaging conditions and aiming for the same image quality and SNR, the use of a negative stage bias can allow for a 20-fold decrease in dwell time, thus reducing the time for a week long acquisition to less than 8 h. We further show that negative bias potential can be applied in an integrated correlative light electron microscopy (CLEM) application, allowing fast acquisition of a high precision overlaid LM-EM dataset. Application of negative stage bias potential will thus help to solve the current bottleneck of image acquisition of large fields of view at high resolution in large-scale microscopy.
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Affiliation(s)
- Ryan Lane
- Imaging Physics, Delft University of Technology, The Netherlands
| | - Yoram Vos
- Imaging Physics, Delft University of Technology, The Netherlands
| | - Anouk H G Wolters
- Department of Biomedical Sciences of Cells and Systems, University Groningen, University Medical Center Groningen, The Netherlands
| | - Luc van Kessel
- Imaging Physics, Delft University of Technology, The Netherlands
| | - S Elisa Chen
- Cell Biology, Center for Molecular Medicine, University Medical Center Utrecht, The Netherlands
| | - Nalan Liv
- Cell Biology, Center for Molecular Medicine, University Medical Center Utrecht, The Netherlands
| | - Judith Klumperman
- Cell Biology, Center for Molecular Medicine, University Medical Center Utrecht, The Netherlands
| | - Ben N G Giepmans
- Department of Biomedical Sciences of Cells and Systems, University Groningen, University Medical Center Groningen, The Netherlands
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24
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Heiligenstein X, de Beer M, Heiligenstein J, Eyraud F, Manet L, Schmitt F, Lamers E, Lindenau J, Kea-Te Lindert M, Salamero J, Raposo G, Sommerdijk N, Belle M, Akiva A. HPM live μ for a full CLEM workflow. Methods Cell Biol 2021; 162:115-149. [PMID: 33707009 DOI: 10.1016/bs.mcb.2020.10.022] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.7] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 02/08/2023]
Abstract
With the development of advanced imaging methods that took place in the last decade, the spatial correlation of microscopic and spectroscopic information-known as multimodal imaging or correlative microscopy (CM)-has become a broadly applied technique to explore biological and biomedical materials at different length scales. Among the many different combinations of techniques, Correlative Light and Electron Microscopy (CLEM) has become the flagship of this revolution. Where light (mainly fluorescence) microscopy can be used directly for the live imaging of cells and tissues, for almost all applications, electron microscopy (EM) requires fixation of the biological materials. Although sample preparation for EM is traditionally done by chemical fixation and embedding in a resin, rapid cryogenic fixation (vitrification) has become a popular way to avoid the formation of artifacts related to the chemical fixation/embedding procedures. During vitrification, the water in the sample transforms into an amorphous ice, keeping the ultrastructure of the biological sample as close as possible to the native state. One immediate benefit of this cryo-arrest is the preservation of protein fluorescence, allowing multi-step multi-modal imaging techniques for CLEM. To minimize the delay separating live imaging from cryo-arrest, we developed a high-pressure freezing (HPF) system directly coupled to a light microscope. We address the optimization of sample preservation and the time needed to capture a biological event, going from live imaging to cryo-arrest using HPF. To further explore the potential of cryo-fixation related to the forthcoming transition from imaging 2D (cell monolayers) to imaging 3D samples (tissue) and the associated importance of homogeneous deep vitrification, the HPF core technology has been revisited to allow easy modification of the environmental parameters during vitrification. Lastly, we will discuss the potential of our HPM within CLEM protocols especially for correlating live imaging using the Zeiss LSM900 with electron microscopy.
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Affiliation(s)
| | - Marit de Beer
- Electron Microscopy Center, Radboudumc Technology Center Microscopy, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands; Department of Cell Biology, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands; Department of Biochemistry, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands
| | | | | | | | | | | | | | - Mariska Kea-Te Lindert
- Electron Microscopy Center, Radboudumc Technology Center Microscopy, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands; Department of Cell Biology, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands
| | - Jean Salamero
- SERPICO Inria Team/UMR 144 CNRS & National Biology and Health Infrastructure "France Bioimaging", Institut Curie, Paris, France
| | - Graça Raposo
- Institut Curie, PSL Research University, CNRS, UMR144, Cell and Tissue Imaging Facility (PICT-IBiSA), Paris, France; Institut Curie, PSL Research University, CNRS, UMR144, Structure and Membrane Compartments, Paris, France
| | - Nico Sommerdijk
- Electron Microscopy Center, Radboudumc Technology Center Microscopy, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands; Department of Biochemistry, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands
| | | | - Anat Akiva
- Electron Microscopy Center, Radboudumc Technology Center Microscopy, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands; Department of Cell Biology, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands.
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25
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Yoon A, Herzog A, Grosse P, Alsem DH, Chee SW, Roldán Cuenya B. Dynamic Imaging of Nanostructures in an Electrolyte with a Scanning Electron Microscope. MICROSCOPY AND MICROANALYSIS : THE OFFICIAL JOURNAL OF MICROSCOPY SOCIETY OF AMERICA, MICROBEAM ANALYSIS SOCIETY, MICROSCOPICAL SOCIETY OF CANADA 2021; 27:121-128. [PMID: 33403947 DOI: 10.1017/s1431927620024769] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/12/2023]
Abstract
The development of microfabricated liquid cells has enabled dynamic studies of nanostructures within a liquid environment with electron microscopy. While such setups are most commonly found in transmission electron microscope (TEM) holders, their implementation in a scanning electron microscope (SEM) offers intriguing potential for multi-modal studies where the large chamber volume allows for the integration of multiple detectors. Here, we describe an electrochemical liquid cell SEM platform that employs the same cells enclosed by silicon nitride membrane windows found in liquid cell TEM holders and demonstrate the imaging of copper oxide nanoparticles in solution using both backscattered and transmitted electrons. In particular, the transmitted electron images collected at high scattering angles show contrast inversion at liquid layer thicknesses of several hundred nanometers, which can be used to determine the presence of liquid in the cell, while maintaining enough resolution to image nanoparticles that are tens of nanometers in size. Using Monte Carlo simulations, we show that both imaging modes have their advantages for liquid phase imaging and rationalize the contrast inversion observed in the transmitted electron image.
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Affiliation(s)
- Aram Yoon
- Department of Interface Science, Fritz-Haber-Institute of the Max-Planck Society, Berlin, Germany
| | - Antonia Herzog
- Department of Interface Science, Fritz-Haber-Institute of the Max-Planck Society, Berlin, Germany
| | - Philipp Grosse
- Department of Interface Science, Fritz-Haber-Institute of the Max-Planck Society, Berlin, Germany
| | | | - See Wee Chee
- Department of Interface Science, Fritz-Haber-Institute of the Max-Planck Society, Berlin, Germany
| | - Beatriz Roldán Cuenya
- Department of Interface Science, Fritz-Haber-Institute of the Max-Planck Society, Berlin, Germany
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26
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Vos Y, Lane RI, Peddie CJ, Wolters AHG, Hoogenboom JP. Retarding Field Integrated Fluorescence and Electron Microscope. MICROSCOPY AND MICROANALYSIS : THE OFFICIAL JOURNAL OF MICROSCOPY SOCIETY OF AMERICA, MICROBEAM ANALYSIS SOCIETY, MICROSCOPICAL SOCIETY OF CANADA 2021; 27:109-120. [PMID: 33349285 DOI: 10.1017/s1431927620024745] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/12/2023]
Abstract
The authors present the application of a retarding field between the electron objective lens and sample in an integrated fluorescence and electron microscope. The retarding field enhances signal collection and signal strength in the electron microscope. This is beneficial for samples prepared for integrated fluorescence and electron microscopy as the amount of staining material added to enhance electron microscopy signal is typically lower compared to conventional samples in order to preserve fluorescence. We demonstrate signal enhancement through the applied retarding field for both 80-nm post-embedding immunolabeled sections and 100-nm in-resin preserved fluorescence sections. Moreover, we show that tuning the electron landing energy particularly improves imaging conditions for ultra-thin (50 nm) sections, where optimization of both retarding field and interaction volume contribute to the signal improvement. Finally, we show that our integrated retarding field setup allows landing energies down to a few electron volts with 0.3 eV dispersion, which opens new prospects for assessing electron beam induced damage by in situ quantification of the observed bleaching of the fluorescence following irradiation.
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Affiliation(s)
- Yoram Vos
- Faculty of Applied Sciences, Delft University of Technology, Lorentzweg 1, Delft2628CJ, The Netherlands
| | - Ryan I Lane
- Faculty of Applied Sciences, Delft University of Technology, Lorentzweg 1, Delft2628CJ, The Netherlands
| | - Chris J Peddie
- Electron Microscopy STP, The Francis Crick Institute, 1 Midland Road, LondonNW1 1AT, UK
| | - Anouk H G Wolters
- Department of Cell Biology, University Medical Center Groningen, University of Groningen, A. Deusinglaan 1, Groningen9713 AV, The Netherlands
| | - Jacob P Hoogenboom
- Faculty of Applied Sciences, Delft University of Technology, Lorentzweg 1, Delft2628CJ, The Netherlands
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27
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Thomas CI, Ryan MA, Scholl B, Guerrero-Given D, Fitzpatrick D, Kamasawa N. Targeting Functionally Characterized Synaptic Architecture Using Inherent Fiducials and 3D Correlative Microscopy. MICROSCOPY AND MICROANALYSIS : THE OFFICIAL JOURNAL OF MICROSCOPY SOCIETY OF AMERICA, MICROBEAM ANALYSIS SOCIETY, MICROSCOPICAL SOCIETY OF CANADA 2021; 27:156-169. [PMID: 33303051 DOI: 10.1017/s1431927620024757] [Citation(s) in RCA: 13] [Impact Index Per Article: 4.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/12/2023]
Abstract
Brain circuits are highly interconnected three-dimensional structures fabricated from components ranging vastly in size; from cell bodies to individual synapses. While neuronal activity can be visualized with advanced light microscopy (LM) techniques, the resolution of electron microscopy (EM) is critical for identifying synaptic connections between neurons. Here, we combine these two techniques, affording the advantage of each and allowing for measurements to be made of the same neural features across imaging platforms. We established an EM-label-free workflow utilizing inherent structural features to correlate in vivo two-photon LM and volumetric scanning EM (SEM) in the ferret visual cortex. By optimizing the volume SEM sample preparation protocol, imaging with the OnPoint detector, and utilizing the focal charge compensation device during serial block-face imaging, we achieved sufficient resolution and signal-to-noise ratio to analyze synaptic ultrastructure for hundreds of synapses within sample volumes. Our novel workflow provides a reliable method for quantitatively characterizing synaptic ultrastructure in functionally imaged neurons, providing new insights into neuronal circuit organization.
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Affiliation(s)
- Connon I Thomas
- Electron Microscopy Core Facility, Imaging Center, Max Planck Florida Institute for Neuroscience, 1 Max Planck Way, Jupiter, FL33458, USA
| | - Melissa A Ryan
- Electron Microscopy Core Facility, Imaging Center, Max Planck Florida Institute for Neuroscience, 1 Max Planck Way, Jupiter, FL33458, USA
| | - Benjamin Scholl
- Functional Architecture and Development of Cerebral Cortex, Max Planck Florida Institute for Neuroscience, 1 Max Planck Way, Jupiter, FL33458, USA
| | - Debbie Guerrero-Given
- Electron Microscopy Core Facility, Imaging Center, Max Planck Florida Institute for Neuroscience, 1 Max Planck Way, Jupiter, FL33458, USA
| | - David Fitzpatrick
- Functional Architecture and Development of Cerebral Cortex, Max Planck Florida Institute for Neuroscience, 1 Max Planck Way, Jupiter, FL33458, USA
| | - Naomi Kamasawa
- Electron Microscopy Core Facility, Imaging Center, Max Planck Florida Institute for Neuroscience, 1 Max Planck Way, Jupiter, FL33458, USA
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28
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Naya M, Sato C. Pyrene Excimer-Based Fluorescent Labeling of Cysteines Brought into Close Proximity by Protein Dynamics: ASEM-Induced Thiol-Ene Click Reaction for High Spatial Resolution CLEM. Int J Mol Sci 2020; 21:E7550. [PMID: 33066147 PMCID: PMC7589919 DOI: 10.3390/ijms21207550] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/04/2020] [Revised: 10/02/2020] [Accepted: 10/08/2020] [Indexed: 12/16/2022] Open
Abstract
Fluorescence microscopy (FM) has revealed vital molecular mechanisms of life. Mainly, molecules labeled by fluorescent probes are imaged. However, the diversity of labeling probes and their functions remain limited. We synthesized a pyrene-based fluorescent probe targeting SH groups, which are important for protein folding and oxidative stress sensing in cells. The labeling achieved employs thiol-ene click reactions between the probes and SH groups and is triggered by irradiation by UV light or an electron beam. When two tagged pyrene groups were close enough to be excited as a dimer (excimer), they showed red-shifted fluorescence; theoretically, the proximity of two SH residues within ~30 Å can thus be monitored. Moreover, correlative light/electron microscopy (CLEM) was achieved using our atmospheric scanning electron microscope (ASEM); radicals formed in liquid by the electron beam caused the thiol-ene click reactions, and excimer fluorescence of the labeled proteins in cells and tissues was visualized by FM. Since the fluorescent labeling is induced by a narrow electron beam, high spatial resolution labeling is expected. The method can be widely applied to biological fields, for example, to study protein dynamics with or without cysteine mutagenesis, and to beam-induced micro-fabrication and the precise post-modification of materials.
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Affiliation(s)
- Masami Naya
- Health and Medical Research Institute, National Institute of Advanced Industrial Science and Technology (AIST), Tsukuba 305-8566, Japan;
| | - Chikara Sato
- Health and Medical Research Institute, National Institute of Advanced Industrial Science and Technology (AIST), Tsukuba 305-8566, Japan;
- Master’s and Doctoral Programs in Neuroscience, Graduate School of Comprehensive Human Sciences, University of Tsukuba, Tsukuba 305-8574, Japan
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29
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Tian X, De Pace C, Ruiz-Perez L, Chen B, Su R, Zhang M, Zhang R, Zhang Q, Wang Q, Zhou H, Wu J, Zhang Z, Tian Y, Battaglia G. A Cyclometalated Iridium (III) Complex as a Microtubule Probe for Correlative Super-Resolution Fluorescence and Electron Microscopy. ADVANCED MATERIALS (DEERFIELD BEACH, FLA.) 2020; 32:e2003901. [PMID: 32815192 DOI: 10.1002/adma.202003901] [Citation(s) in RCA: 10] [Impact Index Per Article: 2.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Received: 06/08/2020] [Revised: 07/12/2020] [Indexed: 06/11/2023]
Abstract
The visualization of microtubules by combining optical and electron microscopy techniques provides valuable information to understand correlated intracellular activities. However, the lack of appropriate probes to bridge both microscopic resolutions restricts the areas and structures that can be comprehended within such highly assembled structures. Here, a versatile cyclometalated iridium (III) complex is designed that achieves synchronous fluorescence-electron microscopy correlation. The selective insertion of the probe into a microtubule triggers remarkable fluorescence enhancement and promising electron contrast. The long-life, highly photostable probe allows live-cell super-resolution imaging of tubulin localization and motion with a resolution of ≈30 nm. Furthermore, correlative light-electron microscopy and energy-filtered transmission electron microscopy reveal the well-associated optical and electron signal at a high specificity, with an interspace of ≈41 Å of microtubule monomer in cells.
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Affiliation(s)
- Xiaohe Tian
- School of Life Science, Anhui University, Hefei, 230000, P. R. China
- Department of Chemistry, Anhui University, Hefei, 230000, P. R. China
- Institute of Physical Science and Information Technology, Anhui University, Hefei, 230000, P. R. China
- Department of Chemistry, University College London, London, WC1H 0AJ, UK
| | - Cesare De Pace
- Department of Chemistry, University College London, London, WC1H 0AJ, UK
- Institute for the Physics of Living Systems, University College London, London, WC1E 6BT, UK
- EPSRC/JEOL Centre for Liquid Phase Electron Microscopy, University College London, London, WC1H 0AJ, UK
| | - Lorena Ruiz-Perez
- Department of Chemistry, University College London, London, WC1H 0AJ, UK
- Institute for the Physics of Living Systems, University College London, London, WC1E 6BT, UK
- EPSRC/JEOL Centre for Liquid Phase Electron Microscopy, University College London, London, WC1H 0AJ, UK
| | - Bo Chen
- School of Materials Science and Engineering, Tongji University, Shanghai, 201804, China
| | - Rina Su
- School of Life Science, Anhui University, Hefei, 230000, P. R. China
| | - Mingzhu Zhang
- Department of Chemistry, Anhui University, Hefei, 230000, P. R. China
| | - Ruilong Zhang
- Department of Chemistry, Anhui University, Hefei, 230000, P. R. China
| | - Qiong Zhang
- Department of Chemistry, Anhui University, Hefei, 230000, P. R. China
| | - Qin Wang
- Biotechnology Centre, Anhui Agriculture University, Hefei, 230036, China
| | - Hongping Zhou
- Department of Chemistry, Anhui University, Hefei, 230000, P. R. China
| | - Jieying Wu
- Department of Chemistry, Anhui University, Hefei, 230000, P. R. China
| | - Zhongping Zhang
- Department of Chemistry, Anhui University, Hefei, 230000, P. R. China
- Institute of Physical Science and Information Technology, Anhui University, Hefei, 230000, P. R. China
- CAS Center for Excellence in Nanoscience, Institute of Intelligent Machines, Chinese Academy of Science, Hefei, 230031, China
| | - Yupeng Tian
- Department of Chemistry, Anhui University, Hefei, 230000, P. R. China
| | - Giuseppe Battaglia
- Department of Chemistry, Anhui University, Hefei, 230000, P. R. China
- Institute of Physical Science and Information Technology, Anhui University, Hefei, 230000, P. R. China
- Institute for the Physics of Living Systems, University College London, London, WC1E 6BT, UK
- EPSRC/JEOL Centre for Liquid Phase Electron Microscopy, University College London, London, WC1H 0AJ, UK
- Institute for Bioengineering of Catalonia, The Barcelona Institute for Science and Technology (BIST), Barcelona, 08007, Spain
- Catalan Institution for Research and Advanced Studies, Barcelona, 08010, Spain
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30
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An accelerated procedure for approaching and imaging of optically branded region of interest in tissue. Methods Cell Biol 2020; 162:205-221. [PMID: 33707013 DOI: 10.1016/bs.mcb.2020.08.002] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/04/2023]
Abstract
Many areas of biology have benefited from advances in light microscopy (LM). However, one limitation of the LM approach is that numerous critically important aspects of subcellular machineries are well beyond the resolution of conventional LM. For studying these, electron microscopy (EM) remains the technique of choice to visualize and identify macromolecules at the ultrastructural level. The most powerful approach is combining both techniques, LM and EM (i.e., to apply correlative light/electron microscopy, CLEM) to image exactly the same region of interest. This combination allows, for example, to immuno-localize proteins by LM and then to visualize the ultrastructural context of the same region of the sample. However, the identification and correlation of the regions of interest (ROIs) at the levels of LM and EM remains a major challenge, mostly due to the difficulties with correlation along the Z-axis for both modalities. In this chapter, we address this difficulty and describe an approach for performing CLEM in tissue samples using marks from near-infrared branding as indicators of a ROI, and then using serial block face-scanning electron microscopy (SBF-SEM) to identify and approach this ROI. Once a ROI has been approached, serial sections are collected on grids for high-resolution imaging by transmission EM, and subsequent correlation with LM images showing labeled proteins.
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31
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Keevend K, Coenen T, Herrmann IK. Correlative cathodoluminescence electron microscopy bioimaging: towards single protein labelling with ultrastructural context. NANOSCALE 2020; 12:15588-15603. [PMID: 32677648 DOI: 10.1039/d0nr02563a] [Citation(s) in RCA: 6] [Impact Index Per Article: 1.5] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/11/2023]
Abstract
The understanding of living systems and their building blocks relies heavily on the assessment of structure-function relationships at the nanoscale. Ever since the development of the first optical microscope, the reliance of scientists across disciplines on microscopy has increased. The development of the first electron microscope and with it the access to information at the nanoscale has prompted numerous disruptive discoveries. While fluorescence imaging allows identification of specific entities based on the labelling with fluorophores, the unlabelled constituents of the samples remain invisible. In electron microscopy on the other hand, structures can be comprehensively visualized based on their distinct electron density and geometry. Although electron microscopy is a powerful tool, it does not implicitly provide information on the location and activity of specific organic molecules. While correlative light and electron microscopy techniques have attempted to unify the two modalities, the resolution mismatch between the two data sets poses major challenges. Recent developments in optical super resolution microscopy enable high resolution correlative light and electron microscopy, however, with considerable constraints due to sample preparation requirements. Labelling of specific structures directly for electron microscopy using small gold nanoparticles (i.e. immunogold) has been used extensively. However, identification of specific entities solely based on electron contrast, and the differentiation from endogenous dense granules, remains challenging. Recently, the use of correlative cathodoluminescence electron microscopy (CCLEM) imaging based on luminescent inorganic nanocrystals has been proposed. While nanometric resolution can be reached for both the electron and the optical signal, high energy electron beams are potentially damaging to the sample. In this review, we discuss the opportunities of (volumetric) multi-color single protein labelling based on correlative cathodoluminescence electron microscopy, and its prospective impact on biomedical research in general. We elaborate on the potential challenges of correlative cathodoluminescence electron microscopy-based bioimaging and benchmark CCLEM against alternative high-resolution correlative imaging techniques.
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Affiliation(s)
- Kerda Keevend
- Laboratory for Particles Biology Interactions, Swiss Federal Laboratories for Materials Science and Technology (Empa), Lerchenfeldstrasse 5, CH-9014, St Gallen, Switzerland.
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32
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Huang R, Feng FP, Huang CH, Mao L, Tang M, Yan ZY, Shao B, Qin L, Xu T, Xue YH, Zhu BZ. Chiral Os(II) Polypyridyl Complexes as Enantioselective Nuclear DNA Imaging Agents Especially Suitable for Correlative High-Resolution Light and Electron Microscopy Studies. ACS APPLIED MATERIALS & INTERFACES 2020; 12:3465-3473. [PMID: 31913004 DOI: 10.1021/acsami.9b19776] [Citation(s) in RCA: 11] [Impact Index Per Article: 2.8] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 05/22/2023]
Abstract
The high-resolution technique transmission electron microscopy (TEM), with OsO4 as the traditional fixative, is an essential tool for cell biology and medicine. Although OsO4 has been extensively used, it is far from perfect because of its high volatility and toxicity. Os(II) polypyridyl complexes like [Os(phen)2(dppz)]2+ (phen = 1,10-phenanthroline; dppz = dipyridophenazine) are not only the well-known molecular DNA "light-switches" but also the potential ideal candidates for TEM studies. Here, we report that the cell-impermeable cationic [Os(phen)2(dppz)]2+ can be preferentially delivered into the live-cell nucleus through ion-pairing with chlorophenolate counter-anions, where it functions as an unparalleled enantioselective nuclear DNA imaging reagent especially suitable for correlative light and electron microscopy (CLEM) studies in both living and fixed cells, which can clearly visualize chromosome aggregation and decondensation during mitosis simultaneously. We propose that the chiral Os(II) polypyridyl complexes can be used as a distinctive group of enantioselective high-resolution CLEM imaging probes for live-cell nuclear DNA studies.
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Affiliation(s)
- Rong Huang
- State Key Laboratory of Environmental Chemistry and Ecotoxicology, Research Center for Eco-Environmental Sciences , Chinese Academy of Sciences , Beijing 100085 , P. R. China
- University of Chinese Academy of Sciences , Beijing 100049 , P. R. China
| | - Feng-Ping Feng
- National Laboratory of Biomacromolecules, CAS Center for Excellence in Biomacromolecules, Institute of Biophysics , Chinese Academy of Sciences , Beijing 100101 , China
| | - Chun-Hua Huang
- State Key Laboratory of Environmental Chemistry and Ecotoxicology, Research Center for Eco-Environmental Sciences , Chinese Academy of Sciences , Beijing 100085 , P. R. China
- University of Chinese Academy of Sciences , Beijing 100049 , P. R. China
| | - Li Mao
- State Key Laboratory of Environmental Chemistry and Ecotoxicology, Research Center for Eco-Environmental Sciences , Chinese Academy of Sciences , Beijing 100085 , P. R. China
- University of Chinese Academy of Sciences , Beijing 100049 , P. R. China
| | - Miao Tang
- State Key Laboratory of Environmental Chemistry and Ecotoxicology, Research Center for Eco-Environmental Sciences , Chinese Academy of Sciences , Beijing 100085 , P. R. China
- University of Chinese Academy of Sciences , Beijing 100049 , P. R. China
| | - Zhu-Ying Yan
- State Key Laboratory of Environmental Chemistry and Ecotoxicology, Research Center for Eco-Environmental Sciences , Chinese Academy of Sciences , Beijing 100085 , P. R. China
- University of Chinese Academy of Sciences , Beijing 100049 , P. R. China
| | - Bo Shao
- State Key Laboratory of Environmental Chemistry and Ecotoxicology, Research Center for Eco-Environmental Sciences , Chinese Academy of Sciences , Beijing 100085 , P. R. China
- University of Chinese Academy of Sciences , Beijing 100049 , P. R. China
| | - Li Qin
- State Key Laboratory of Environmental Chemistry and Ecotoxicology, Research Center for Eco-Environmental Sciences , Chinese Academy of Sciences , Beijing 100085 , P. R. China
- University of Chinese Academy of Sciences , Beijing 100049 , P. R. China
| | - Tao Xu
- University of Chinese Academy of Sciences , Beijing 100049 , P. R. China
- National Laboratory of Biomacromolecules, CAS Center for Excellence in Biomacromolecules, Institute of Biophysics , Chinese Academy of Sciences , Beijing 100101 , China
| | - Yan-Hong Xue
- National Laboratory of Biomacromolecules, CAS Center for Excellence in Biomacromolecules, Institute of Biophysics , Chinese Academy of Sciences , Beijing 100101 , China
| | - Ben-Zhan Zhu
- State Key Laboratory of Environmental Chemistry and Ecotoxicology, Research Center for Eco-Environmental Sciences , Chinese Academy of Sciences , Beijing 100085 , P. R. China
- University of Chinese Academy of Sciences , Beijing 100049 , P. R. China
- Joint Institute for Environmental Science , Research Center for Eco-Environmental Sciences and Hong Kong Baptist University , Kowloon 999077 , Hong Kong
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33
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Arlauckas SP, Browning EA, Poptani H, Delikatny EJ. Imaging of cancer lipid metabolism in response to therapy. NMR IN BIOMEDICINE 2019; 32:e4070. [PMID: 31107583 DOI: 10.1002/nbm.4070] [Citation(s) in RCA: 9] [Impact Index Per Article: 1.8] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 07/28/2018] [Revised: 12/21/2018] [Accepted: 12/21/2018] [Indexed: 06/09/2023]
Abstract
Lipids represent a diverse array of molecules essential to the cell's structure, defense, energy, and communication. Lipid metabolism can often become dysregulated during tumor development. During cancer therapy, targeted inhibition of cell proliferation can likewise cause widespread and drastic changes in lipid composition. Molecular imaging techniques have been developed to monitor altered lipid profiles as a biomarker for cancer diagnosis and treatment response. For decades, MRS has been the dominant non-invasive technique for studying lipid metabolite levels. Recent insights into the oncogenic transformations driving changes in lipid metabolism have revealed new mechanisms and signaling molecules that can be exploited using optical imaging, mass spectrometry imaging, and positron emission tomography. These novel imaging modalities have provided researchers with a diverse toolbox to examine changes in lipids in response to a wide array of anticancer strategies including chemotherapy, radiation therapy, signal transduction inhibitors, gene therapy, immunotherapy, or a combination of these strategies. The understanding of lipid metabolism in response to cancer therapy continues to evolve as each therapeutic method emerges, and this review seeks to summarize the current field and areas of unmet needs.
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Affiliation(s)
- Sean Philip Arlauckas
- Department of Radiology, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA
- Center for Systems Biology, Mass General Hospital, Boston, MA, USA
| | - Elizabeth Anne Browning
- Department of Radiology, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA
| | - Harish Poptani
- Department of Cellular and Molecular Physiology, Institute of Regenerative Medicine, University of Liverpool, Liverpool, UK
| | - Edward James Delikatny
- Department of Radiology, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA
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34
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Smith D, Starborg T. Serial block face scanning electron microscopy in cell biology: Applications and technology. Tissue Cell 2019; 57:111-122. [DOI: 10.1016/j.tice.2018.08.011] [Citation(s) in RCA: 21] [Impact Index Per Article: 4.2] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/31/2018] [Revised: 08/22/2018] [Accepted: 08/26/2018] [Indexed: 10/28/2022]
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35
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Mohammadian S, Fokkema J, Agronskaia AV, Liv N, de Heus C, van Donselaar E, Blab GA, Klumperman J, Gerritsen HC. High accuracy, fiducial marker-based image registration of correlative microscopy images. Sci Rep 2019; 9:3211. [PMID: 30824844 PMCID: PMC6397213 DOI: 10.1038/s41598-019-40098-4] [Citation(s) in RCA: 22] [Impact Index Per Article: 4.4] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/02/2018] [Accepted: 01/28/2019] [Indexed: 12/03/2022] Open
Abstract
Fluorescence microscopy (FM) and electron microscopy (EM) are complementary techniques. FM affords examination of large fields of view and identifying regions of interest but has a low resolution. EM exhibits excellent resolution over a limited field of view. The combination of these two techniques, correlative microscopy, received considerable interest in the past years and has proven its potential in biology and material science. Accurate correlation of FM and EM images is, however, challenging due to the differences in contrast mechanism, size of field of view and resolution. We report an accurate, fast and robust method to correlate FM and EM images using low densities of fiducial markers. Here, 120 nm diameter fiducial markers consisting of fluorescently labelled silica coated gold nanoparticles are used. The method relies on recording FM, low magnification EM and high magnification EM images. Two linear transformation matrices are constructed, FM to low magnification EM and low magnification EM to high magnification EM. Combination of these matrices results in a high accuracy transformation of FM to high magnification EM coordinates. The method was tested using two different transmission electron microscopes and different Tokuyasu and Lowicryl sections. The overall accuracy of the correlation method is high, 5-30 nm.
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Affiliation(s)
- Sajjad Mohammadian
- Molecular Biophysics, Debye Institute for Nanomaterials Science, Utrecht University, Utrecht, The Netherlands
| | - Jantina Fokkema
- Molecular Biophysics, Debye Institute for Nanomaterials Science, Utrecht University, Utrecht, The Netherlands
| | - Alexandra V Agronskaia
- Molecular Biophysics, Debye Institute for Nanomaterials Science, Utrecht University, Utrecht, The Netherlands
| | - Nalan Liv
- Section Cell Biology, Center for Molecular Medicine, University Medical Center Utrecht, Utrecht, The Netherlands
| | - Cecilia de Heus
- Section Cell Biology, Center for Molecular Medicine, University Medical Center Utrecht, Utrecht, The Netherlands
| | - Elly van Donselaar
- Section Cell Biology, Center for Molecular Medicine, University Medical Center Utrecht, Utrecht, The Netherlands
| | - Gerhard A Blab
- Molecular Biophysics, Debye Institute for Nanomaterials Science, Utrecht University, Utrecht, The Netherlands
| | - Judith Klumperman
- Section Cell Biology, Center for Molecular Medicine, University Medical Center Utrecht, Utrecht, The Netherlands
| | - Hans C Gerritsen
- Molecular Biophysics, Debye Institute for Nanomaterials Science, Utrecht University, Utrecht, The Netherlands.
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Staszowska AD, Fox-Roberts P, Hirvonen LM, Peddie CJ, Collinson LM, Jones GE, Cox S. The Rényi divergence enables accurate and precise cluster analysis for localization microscopy. Bioinformatics 2018; 34:4102-4111. [PMID: 29868717 PMCID: PMC6247934 DOI: 10.1093/bioinformatics/bty403] [Citation(s) in RCA: 4] [Impact Index Per Article: 0.7] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/07/2017] [Accepted: 05/30/2018] [Indexed: 11/18/2022] Open
Abstract
Motivation Clustering analysis is a key technique for quantitatively characterizing structures in localization microscopy images. To build up accurate information about biological structures, it is critical that the quantification is both accurate (close to the ground truth) and precise (has small scatter and is reproducible). Results Here, we describe how the Rényi divergence can be used for cluster radius measurements in localization microscopy data. We demonstrate that the Rényi divergence can operate with high levels of background and provides results which are more accurate than Ripley’s functions, Voronoi tesselation or DBSCAN. Availability and implementation The data supporting this research and the software described are accessible at the following site: https://dx.doi.org/10.18742/RDM01-316. Correspondence and requests for materials should be addressed to the corresponding author. Supplementary information Supplementary data are available at Bioinformatics online.
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Affiliation(s)
- Adela D Staszowska
- Randall Centre for Cell and Molecular Biophysics, King's College London, London, UK
| | - Patrick Fox-Roberts
- Randall Centre for Cell and Molecular Biophysics, King's College London, London, UK
| | - Liisa M Hirvonen
- Randall Centre for Cell and Molecular Biophysics, King's College London, London, UK
| | - Christopher J Peddie
- Electron Microscopy Science Technology Platform, The Francis Crick Institute, London, UK
| | - Lucy M Collinson
- Electron Microscopy Science Technology Platform, The Francis Crick Institute, London, UK
| | - Gareth E Jones
- Randall Centre for Cell and Molecular Biophysics, King's College London, London, UK
| | - Susan Cox
- Randall Centre for Cell and Molecular Biophysics, King's College London, London, UK
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37
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Porrati F, Grewe D, Seybert A, Frangakis AS, Eltsov M. FIB-SEM imaging properties of Drosophila melanogaster tissues embedded in Lowicryl HM20. J Microsc 2018; 273:91-104. [PMID: 30417390 DOI: 10.1111/jmi.12764] [Citation(s) in RCA: 5] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/13/2017] [Revised: 09/17/2018] [Accepted: 10/12/2018] [Indexed: 11/27/2022]
Abstract
Lowicryl resins enable processing of biological material for electron microscopy at the lowest temperatures compatible with resin embedding. When combined with high-pressure freezing and freeze-substitution, Lowicryl embedding supports preservation of fine structural details and fluorescent markers. Here, we analysed the applicability of Lowicryl HM20 embedding for focused ion beam (FIB) scanning electron microscopy (SEM) tomography of Drosophila melanogaster embryonic and larval model systems. We show that the freeze-substitution with per-mill concentrations of uranyl acetate provided sufficient contrast and an image quality of SEM imaging in the range of similar samples analysed by transmission electron microscopy (TEM). Preservation of genetically encoded fluorescent proteins allowed correlative localization of regions of interest (ROI) within the embedded tissue block. TEM on sections cut from the block face enabled evaluation of structural preservation to allow ROI ranking and thus targeted, time-efficient FIB-SEM tomography data collection. The versatility of Lowicryl embedding opens new perspectives for designing hybrid SEM-TEM workflows to comprehensively analyse biological structures. LAY DESCRIPTION: Focused ion beam scanning electron microscopy is becoming a widely used technique for the three-dimensional analysis of biological samples at fine structural details beyond levels feasible for light microscopy. To withstand the abrasion of material by the ion beam and the imaging by the scanning electron beam, biological samples have to be embedded into resins, most commonly these are very dense epoxy-based plastics. However, dense resins generate electron scattering which interferes with the signal from the biological specimen. Furthermore, to improve the imaging contrast, epoxy embedding requires chemical treatments with e.g. heavy metals, which deteriorate the ultrastructure of the biological specimen. In this study we explored the applicability of an electron lucent resin, Lowicryl HM 20, for focused ion beam scanning electron microscopy. The Lowicryl embedding workflow operates at milder chemical treatments and lower temperatures, thus preserving the sub-cellular and sub-organellar organization, as well as fluorescent markers visible by light microscopy. Here we show that focus ion beam scanning electron microscopy of Lowicryl-embedded fruit flies tissues provides reliable imaging revealing fine structural details. Our workflow benefited from use of transmission electron microscopy for the quality control of the ultrastructural preservation and fluorescent light microscopy for localization of regions of interest. The versatility of Lowicryl embedding opens up new perspectives for designing hybrid workflows combining fluorescent light, scanning, and transmission electron microscopy techniques to comprehensively analyze biological structures.
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Affiliation(s)
- F Porrati
- Buchmann Institute for Molecular Life Sciences and Institute for Biophysics, Goethe-University, Frankfurt am Main, Germany
| | - D Grewe
- Buchmann Institute for Molecular Life Sciences and Institute for Biophysics, Goethe-University, Frankfurt am Main, Germany
| | - A Seybert
- Buchmann Institute for Molecular Life Sciences and Institute for Biophysics, Goethe-University, Frankfurt am Main, Germany
| | - A S Frangakis
- Buchmann Institute for Molecular Life Sciences and Institute for Biophysics, Goethe-University, Frankfurt am Main, Germany
| | - M Eltsov
- Buchmann Institute for Molecular Life Sciences and Institute for Biophysics, Goethe-University, Frankfurt am Main, Germany
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38
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Ando T, Bhamidimarri SP, Brending N, Colin-York H, Collinson L, De Jonge N, de Pablo PJ, Debroye E, Eggeling C, Franck C, Fritzsche M, Gerritsen H, Giepmans BNG, Grunewald K, Hofkens J, Hoogenboom JP, Janssen KPF, Kaufman R, Klumpermann J, Kurniawan N, Kusch J, Liv N, Parekh V, Peckys DB, Rehfeldt F, Reutens DC, Roeffaers MBJ, Salditt T, Schaap IAT, Schwarz US, Verkade P, Vogel MW, Wagner R, Winterhalter M, Yuan H, Zifarelli G. The 2018 correlative microscopy techniques roadmap. JOURNAL OF PHYSICS D: APPLIED PHYSICS 2018; 51:443001. [PMID: 30799880 PMCID: PMC6372154 DOI: 10.1088/1361-6463/aad055] [Citation(s) in RCA: 75] [Impact Index Per Article: 12.5] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 02/09/2018] [Revised: 06/14/2018] [Accepted: 07/01/2018] [Indexed: 05/19/2023]
Abstract
Developments in microscopy have been instrumental to progress in the life sciences, and many new techniques have been introduced and led to new discoveries throughout the last century. A wide and diverse range of methodologies is now available, including electron microscopy, atomic force microscopy, magnetic resonance imaging, small-angle x-ray scattering and multiple super-resolution fluorescence techniques, and each of these methods provides valuable read-outs to meet the demands set by the samples under study. Yet, the investigation of cell development requires a multi-parametric approach to address both the structure and spatio-temporal organization of organelles, and also the transduction of chemical signals and forces involved in cell-cell interactions. Although the microscopy technologies for observing each of these characteristics are well developed, none of them can offer read-out of all characteristics simultaneously, which limits the information content of a measurement. For example, while electron microscopy is able to disclose the structural layout of cells and the macromolecular arrangement of proteins, it cannot directly follow dynamics in living cells. The latter can be achieved with fluorescence microscopy which, however, requires labelling and lacks spatial resolution. A remedy is to combine and correlate different readouts from the same specimen, which opens new avenues to understand structure-function relations in biomedical research. At the same time, such correlative approaches pose new challenges concerning sample preparation, instrument stability, region of interest retrieval, and data analysis. Because the field of correlative microscopy is relatively young, the capabilities of the various approaches have yet to be fully explored, and uncertainties remain when considering the best choice of strategy and workflow for the correlative experiment. With this in mind, the Journal of Physics D: Applied Physics presents a special roadmap on the correlative microscopy techniques, giving a comprehensive overview from various leading scientists in this field, via a collection of multiple short viewpoints.
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Affiliation(s)
- Toshio Ando
- Nano Life Science Institute (WPI-NanoLSI), Kanazawa University, Kanazawa, Japan
| | | | | | - H Colin-York
- MRC Human Immunology Unit, Weatherall Institute of Molecular Medicine, University of Oxford, Headley Way, OX3 9DS Oxford, United Kingdom
| | | | - Niels De Jonge
- INM-Leibniz Institute for New Materials, 66123 Saarbrücken, Germany
- Saarland University, 66123 Saarbrücken, Germany
| | - P J de Pablo
- Dpto. Física de la Materia Condensada Universidad Autónoma de Madrid 28049, Madrid, Spain
- Instituto de Física de la Materia Condensada IFIMAC, Universidad Autónoma de Madrid 28049, Madrid, Spain
| | - Elke Debroye
- KU Leuven, Department of Chemistry, B-3001 Heverlee, Belgium
| | - Christian Eggeling
- MRC Human Immunology Unit, Weatherall Institute of Molecular Medicine, University of Oxford, Headley Way, OX3 9DS Oxford, United Kingdom
- Institute of Applied Optics, Friedrich-Schiller University, Jena, Germany
- Leibniz Institute of Photonic Technology (IPHT), Jena, Germany
| | - Christian Franck
- Department of Mechanical Engineering, University of Wisconsin-Madison, 1513 University Ave, Madison, WI 53706, United States of America
| | - Marco Fritzsche
- MRC Human Immunology Unit, Weatherall Institute of Molecular Medicine, University of Oxford, Headley Way, OX3 9DS Oxford, United Kingdom
- Kennedy Institute for Rheumatology, University of Oxford, Oxford, United Kingdom
| | - Hans Gerritsen
- Debye Institute, Utrecht University, Utrecht, Netherlands
| | - Ben N G Giepmans
- Department of Cell Biology, University of Groningen, University Medical Center Groningen, Groningen, Netherlands
| | - Kay Grunewald
- Division of Structural Biology, Wellcome Trust Centre for Human Genetics, University of Oxford, Oxford, United Kingdom
- Centre of Structural Systems Biology Hamburg and University of Hamburg, Hamburg, Germany
- Heinrich-Pette-Institute, Leibniz Institute of Virology, Hamburg, Germany
| | - Johan Hofkens
- KU Leuven, Department of Chemistry, B-3001 Heverlee, Belgium
| | | | | | - Rainer Kaufman
- Division of Structural Biology, Wellcome Trust Centre for Human Genetics, University of Oxford, Oxford, United Kingdom
- Centre of Structural Systems Biology Hamburg and University of Hamburg, Hamburg, Germany
- Department of Biochemistry, University of Oxford, Oxford, United Kingdom
| | - Judith Klumpermann
- Section Cell Biology, Center for Molecular Medicine, University Medical Center Utrecht, Utrecht University, Heidelberglaan 100, 3584CX Utrecht, Netherlands
| | - Nyoman Kurniawan
- Centre for Advanced Imaging, The University of Queensland, Brisbane, QLD 4072, Australia
| | | | - Nalan Liv
- Section Cell Biology, Center for Molecular Medicine, University Medical Center Utrecht, Utrecht University, Heidelberglaan 100, 3584CX Utrecht, Netherlands
| | - Viha Parekh
- Centre for Advanced Imaging, The University of Queensland, Brisbane, QLD 4072, Australia
| | - Diana B Peckys
- Faculty of Medicine, Saarland University, 66421 Homburg, Germany
| | - Florian Rehfeldt
- University of Göttingen, Third Institute of Physics-Biophysics, 37077 Göttingen, Germany
| | - David C Reutens
- Centre for Advanced Imaging, The University of Queensland, Brisbane, QLD 4072, Australia
| | | | - Tim Salditt
- University of Göttingen, Institute for X-Ray Physics, 37077 Göttingen, Germany
| | - Iwan A T Schaap
- SmarAct GmbH, Schütte-Lanz-Str. 9, D-26135 Oldenburg, Germany
| | - Ulrich S Schwarz
- Institute for Theoretical Physics and BioQuant, Heidelberg University, Heidelberg, Germany
| | - Paul Verkade
- School of Biochemistry, University of Bristol, Bristol, United Kingdom
| | - Michael W Vogel
- Centre for Advanced Imaging, The University of Queensland, Brisbane, QLD 4072, Australia
| | - Richard Wagner
- Department of Life Sciences & Chemistry, Jacobs University, Bremen, Germany
| | | | - Haifeng Yuan
- KU Leuven, Department of Chemistry, B-3001 Heverlee, Belgium
| | - Giovanni Zifarelli
- Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom
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Ultra-stable super-resolution fluorescence cryo-microscopy for correlative light and electron cryo-microscopy. SCIENCE CHINA-LIFE SCIENCES 2018; 61:1312-1319. [PMID: 30426455 DOI: 10.1007/s11427-018-9380-3] [Citation(s) in RCA: 11] [Impact Index Per Article: 1.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Subscribe] [Scholar Register] [Received: 08/14/2018] [Accepted: 09/17/2018] [Indexed: 11/27/2022]
Abstract
Remarkable progress in correlative light and electron cryo-microscopy (cryo-CLEM) has been made in the past decade. A crucial component for cryo-CLEM is a dedicated cryo-fluorescence microscope (cryo-FM). Here, we describe an ultra-stable super-resolution cryo-FM that exhibits excellent thermal and mechanical stability. The temperature fluctuations in 10 h are less than 0.06 K, and the mechanical drift over 5 h is less than 200 nm in three dimensions. We have demonstrated the super-resolution imaging capability of this system (average single molecule localization accuracy of ∼13.0 nm). The results suggest that our system is particularly suitable for long-term observations, such as single molecule localization microscopy (SMLM) and cryogenic super-resolution correlative light and electron microscopy (csCLEM).
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40
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Santarella-Mellwig R, Haselmann U, Schieber NL, Walther P, Schwab Y, Antony C, Bartenschlager R, Romero-Brey I. Correlative Light Electron Microscopy (CLEM) for Tracking and Imaging Viral Protein Associated Structures in Cryo-immobilized Cells. J Vis Exp 2018. [PMID: 30247481 PMCID: PMC6235138 DOI: 10.3791/58154] [Citation(s) in RCA: 8] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/21/2022] Open
Abstract
Due to its high resolution, electron microscopy (EM) is an indispensable tool for virologists. However, one of the main difficulties when analyzing virus-infected or transfected cells via EM are the low efficiencies of infection or transfection, hindering the examination of these cells. In order to overcome this difficulty, light microscopy (LM) can be performed first to allocate the subpopulation of infected or transfected cells. Thus, taking advantage of the use of fluorescent proteins (FPs) fused to viral proteins, LM is used here to record the positions of the "positive-transfected" cells, expressing a FP and growing on a support with an alphanumeric pattern. Subsequently, cells are further processed for EM via high pressure freezing (HPF), freeze substitution (FS) and resin embedding. The ultra-rapid freezing step ensures excellent membrane preservation of the selected cells that can then be analyzed at the ultrastructural level by transmission electron microscopy (TEM). Here, a step-by-step correlative light electron microscopy (CLEM) workflow is provided, describing sample preparation, imaging and correlation in detail. The experimental design can be also applied to address many cell biology questions.
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Affiliation(s)
| | - Uta Haselmann
- Department of Infectious Diseases, Molecular Virology, Heidelberg University
| | | | - Paul Walther
- Central Facility for Electron Microscopy, Ulm University
| | | | | | - Ralf Bartenschlager
- Department of Infectious Diseases, Molecular Virology, Heidelberg University; Heidelberg Partner Site, German Center for Infection Research;
| | - Inés Romero-Brey
- Department of Infectious Diseases, Molecular Virology, Heidelberg University;
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41
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Chung GHC, Domart MC, Peddie C, Mantell J, Mclaverty K, Arabiotorre A, Hodgson L, Byrne RD, Verkade P, Arkill K, Collinson LM, Larijani B. Acute depletion of diacylglycerol from the cis-Golgi affects localized nuclear envelope morphology during mitosis. J Lipid Res 2018; 59:1402-1413. [PMID: 29895700 PMCID: PMC6071775 DOI: 10.1194/jlr.m083899] [Citation(s) in RCA: 4] [Impact Index Per Article: 0.7] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/15/2018] [Revised: 06/12/2018] [Indexed: 12/28/2022] Open
Abstract
Dysregulation of nuclear envelope (NE) assembly results in various cancers; for example, renal and some lung carcinomas ensue due to NE malformation. The NE is a dynamic membrane compartment and its completion during mitosis is a highly regulated process, but the detailed mechanism still remains incompletely understood. Previous studies have found that isolated diacylglycerol (DAG)-containing vesicles are essential for completing the fusion of the NE in nonsomatic cells. We investigated the impact of DAG depletion from the cis-Golgi in mammalian cells on NE reassembly. Using advanced electron microscopy, we observed an enriched DAG population of vesicles at the vicinity of the NE gaps of telophase mammalian cells. We applied a mini singlet oxygen generator-C1-domain tag that localized DAG-enriched vesicles at the perinuclear region, which suggested the existence of NE fusogenic vesicles. We quantified the impact of Golgi-DAG depletion by measuring the in situ NE rim curvature of the reforming NE. The rim curvature in these cells was significantly reduced compared with controls, which indicated a localized defect in NE morphology. Our novel results demonstrate the significance of the role of DAG from the cis-Golgi for the regulation of NE assembly.
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Affiliation(s)
- Gary Hong Chun Chung
- Cell Biophysics Laboratory, Ikerbasque Basque Foundation for Science, Research Centre for Experimental Marine Biology and Biotechnology (PiE) and Biofísika Institute (UPV/EHU, CSIC), University of the Basque Country, Barrio Sarriena s/n 48940, Leioa, Spain
| | - Marie-Charlotte Domart
- Electron Microscopy Science Technology Platform, Francis Crick Institute, London, United Kingdom
| | - Christopher Peddie
- Electron Microscopy Science Technology Platform, Francis Crick Institute, London, United Kingdom
| | - Judith Mantell
- School of Biochemistry, Faculty of Biomedical Sciences, University of Bristol, Bristol, United Kingdom
| | - Kieran Mclaverty
- Cell Biophysics Laboratory, Ikerbasque Basque Foundation for Science, Research Centre for Experimental Marine Biology and Biotechnology (PiE) and Biofísika Institute (UPV/EHU, CSIC), University of the Basque Country, Barrio Sarriena s/n 48940, Leioa, Spain
- School of Biochemistry, Faculty of Biomedical Sciences, University of Bristol, Bristol, United Kingdom
| | - Angela Arabiotorre
- Cell Biophysics Laboratory, Ikerbasque Basque Foundation for Science, Research Centre for Experimental Marine Biology and Biotechnology (PiE) and Biofísika Institute (UPV/EHU, CSIC), University of the Basque Country, Barrio Sarriena s/n 48940, Leioa, Spain
| | - Lorna Hodgson
- School of Biochemistry, Faculty of Biomedical Sciences, University of Bristol, Bristol, United Kingdom
| | - Richard D Byrne
- Cell Biophysics Laboratory, Ikerbasque Basque Foundation for Science, Research Centre for Experimental Marine Biology and Biotechnology (PiE) and Biofísika Institute (UPV/EHU, CSIC), University of the Basque Country, Barrio Sarriena s/n 48940, Leioa, Spain
| | - Paul Verkade
- School of Biochemistry, Faculty of Biomedical Sciences, University of Bristol, Bristol, United Kingdom
- Wolfson Bioimaging Facility, University of Bristol, Bristol, United Kingdom
| | - Kenton Arkill
- Cell Biophysics Laboratory, Ikerbasque Basque Foundation for Science, Research Centre for Experimental Marine Biology and Biotechnology (PiE) and Biofísika Institute (UPV/EHU, CSIC), University of the Basque Country, Barrio Sarriena s/n 48940, Leioa, Spain
- Division of Cancer and Stem Cells, The Queen's Medical Centre, University of Nottingham, Nottingham, United Kingdom
| | - Lucy M Collinson
- Electron Microscopy Science Technology Platform, Francis Crick Institute, London, United Kingdom
| | - Banafshé Larijani
- Cell Biophysics Laboratory, Ikerbasque Basque Foundation for Science, Research Centre for Experimental Marine Biology and Biotechnology (PiE) and Biofísika Institute (UPV/EHU, CSIC), University of the Basque Country, Barrio Sarriena s/n 48940, Leioa, Spain
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42
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Clarke NI, Royle SJ. FerriTag is a new genetically-encoded inducible tag for correlative light-electron microscopy. Nat Commun 2018; 9:2604. [PMID: 29973588 PMCID: PMC6031641 DOI: 10.1038/s41467-018-04993-0] [Citation(s) in RCA: 35] [Impact Index Per Article: 5.8] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/30/2017] [Accepted: 05/24/2018] [Indexed: 01/20/2023] Open
Abstract
A current challenge is to develop tags to precisely visualize proteins in cells by light and electron microscopy. Here, we introduce FerriTag, a genetically-encoded chemically-inducible tag for correlative light-electron microscopy. FerriTag is a fluorescent recombinant electron-dense ferritin particle that can be attached to a protein-of-interest using rapamycin-induced heterodimerization. We demonstrate the utility of FerriTag for correlative light-electron microscopy by labeling proteins associated with various intracellular structures including mitochondria, plasma membrane, and clathrin-coated pits and vesicles. FerriTagging has a good signal-to-noise ratio and a labeling resolution of approximately 10 nm. We demonstrate how FerriTagging allows nanoscale mapping of protein location relative to a subcellular structure, and use it to detail the distribution and conformation of huntingtin-interacting protein 1 related (HIP1R) in and around clathrin-coated pits.
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Affiliation(s)
- Nicholas I Clarke
- Centre for Mechanochemical Cell Biology, Warwick Medical School, Gibbet Hill Road, Coventry, CV4 7AL, UK
| | - Stephen J Royle
- Centre for Mechanochemical Cell Biology, Warwick Medical School, Gibbet Hill Road, Coventry, CV4 7AL, UK.
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43
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DELPIANO J, PIZARRO L, PEDDIE C, JONES M, GRIFFIN L, COLLINSON L. Automated detection of fluorescent cells in in-resin fluorescence sections for integrated light and electron microscopy. J Microsc 2018; 271:109-119. [PMID: 29698565 PMCID: PMC6032852 DOI: 10.1111/jmi.12700] [Citation(s) in RCA: 12] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/21/2017] [Accepted: 03/13/2018] [Indexed: 12/25/2022]
Abstract
Integrated array tomography combines fluorescence and electron imaging of ultrathin sections in one microscope, and enables accurate high-resolution correlation of fluorescent proteins to cell organelles and membranes. Large numbers of serial sections can be imaged sequentially to produce aligned volumes from both imaging modalities, thus producing enormous amounts of data that must be handled and processed using novel techniques. Here, we present a scheme for automated detection of fluorescent cells within thin resin sections, which could then be used to drive automated electron image acquisition from target regions via 'smart tracking'. The aim of this work is to aid in optimization of the data acquisition process through automation, freeing the operator to work on other tasks and speeding up the process, while reducing data rates by only acquiring images from regions of interest. This new method is shown to be robust against noise and able to deal with regions of low fluorescence.
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Affiliation(s)
- J. DELPIANO
- School of Engineering and Applied SciencesUniversidad de los AndesSantiagoChile
| | - L. PIZARRO
- Department of Computer ScienceUniversity College LondonLondonUnited Kingdom
| | - C.J. PEDDIE
- Electron MicroscopyThe Francis Crick InstituteLondonUnited Kingdom
| | - M.L. JONES
- Electron MicroscopyThe Francis Crick InstituteLondonUnited Kingdom
| | - L.D. GRIFFIN
- Department of Computer ScienceUniversity College LondonLondonUnited Kingdom
| | - L.M. COLLINSON
- Electron MicroscopyThe Francis Crick InstituteLondonUnited Kingdom
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Burel A, Lavault MT, Chevalier C, Gnaegi H, Prigent S, Mucciolo A, Dutertre S, Humbel BM, Guillaudeux T, Kolotuev I. A targeted 3D EM and correlative microscopy method using SEM array tomography. Development 2018; 145:dev.160879. [PMID: 29802150 DOI: 10.1242/dev.160879] [Citation(s) in RCA: 37] [Impact Index Per Article: 6.2] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/25/2017] [Accepted: 05/16/2018] [Indexed: 12/15/2022]
Abstract
Using electron microscopy to localize rare cellular events or structures in complex tissue is challenging. Correlative light and electron microscopy procedures have been developed to link fluorescent protein expression with ultrastructural resolution. Here, we present an optimized scanning electron microscopy (SEM) workflow for volumetric array tomography for asymmetric samples and model organisms (Caenorhabditis elegans, Drosophila melanogaster, Danio rerio). We modified a diamond knife to simplify serial section array acquisition with minimal artifacts. After array acquisition, the arrays were transferred to a glass coverslip or silicon wafer support. Using light microscopy, the arrays were screened rapidly for initial recognition of global anatomical features (organs or body traits). Then, using SEM, an in-depth study of the cells and/or organs of interest was performed. Our manual and automatic data acquisition strategies make 3D data acquisition and correlation simpler and more precise than alternative methods. This method can be used to address questions in cell and developmental biology that require the efficient identification of a labeled cell or organelle.
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Affiliation(s)
- Agnes Burel
- University of Rennes 1, UMS Biosit, MRic, 35043 Rennes, France
| | | | | | | | - Sylvain Prigent
- University of Rennes 1, UMS Biosit, MRic, 35043 Rennes, France
| | - Antonio Mucciolo
- University of Lausanne, Faculté de biologie et de médecine, Electron Microscopy Facility, CH-1015 Lausanne, Switzerland
| | | | - Bruno M Humbel
- University of Lausanne, Faculté de biologie et de médecine, Electron Microscopy Facility, CH-1015 Lausanne, Switzerland
| | | | - Irina Kolotuev
- University of Rennes 1, UMS Biosit, MRic, 35043 Rennes, France .,University of Lausanne, Faculté de biologie et de médecine, Electron Microscopy Facility, CH-1015 Lausanne, Switzerland
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Wood PL, Cebak JE, Woltjer RL. Diacylglycerols as biomarkers of sustained immune activation in Proteinopathies associated with dementia. Clin Chim Acta 2018; 476:107-110. [DOI: 10.1016/j.cca.2017.11.009] [Citation(s) in RCA: 7] [Impact Index Per Article: 1.2] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/10/2017] [Revised: 11/10/2017] [Accepted: 11/12/2017] [Indexed: 12/12/2022]
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46
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Hodgson L, Verkade P, Yamauchi Y. Correlative Light and Electron Microscopy of Influenza Virus Entry and Budding. Methods Mol Biol 2018; 1836:237-260. [PMID: 30151577 DOI: 10.1007/978-1-4939-8678-1_12] [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: 12/13/2022]
Abstract
Influenza A virus (IAV) entry is a stepwise process regulated by viral and cellular cues, facilitating cellular functions. Virus entry begins by attachment of hemagglutinin to cell surface sialic acids, followed by endocytic uptake, vesicular transport along microtubules, low-pH-mediated viral membrane fusion with the late endosomal membrane, capsid uncoating, viral ribonucleoprotein (vRNP) release, and nuclear import of vRNPs. Here we show a basic methodology to visualize incoming and egressing IAV particles by correlative light and electron microscopy (CLEM). We combine fluorescence microscopy of virus-infected human lung carcinoma A549 cells with high-pressure freezing (HPF) and in-resin fluorescence CLEM and the Tokuyasu CLEM method. This approach forms a basis to study the virus life cycle and virus-host interactions at the ultrastructural level.
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Affiliation(s)
- Lorna Hodgson
- School of Biochemistry, University of Bristol, Bristol, UK
| | - Paul Verkade
- School of Biochemistry, University of Bristol, Bristol, UK.,Wolfson Bioimaging Facility, University of Bristol, Bristol, UK
| | - Yohei Yamauchi
- School of Cellular and Molecular Medicine, University of Bristol, Bristol, UK.
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47
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Lemercier N, Middel V, Hentsch D, Taubert S, Takamiya M, Beil T, Vonesch JL, Baumbach T, Schultz P, Antony C, Strähle U. Microtome-integrated microscope system for high sensitivity tracking of in-resin fluorescence in blocks and ultrathin sections for correlative microscopy. Sci Rep 2017; 7:13583. [PMID: 29051533 PMCID: PMC5648784 DOI: 10.1038/s41598-017-13348-6] [Citation(s) in RCA: 4] [Impact Index Per Article: 0.6] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/22/2017] [Accepted: 09/22/2017] [Indexed: 12/02/2022] Open
Abstract
Many areas of biological research demand the combined use of different imaging modalities to cover a wide range of magnifications and measurements or to place fluorescent patterns into an ultrastructural context. A technically difficult problem is the efficient specimen transfer between different imaging modalities without losing the coordinates of the regions-of-interest (ROI). Here, we report a new and highly sensitive integrated system that combines a custom designed microscope with an ultramicrotome for in-resin-fluorescence detection in blocks, ribbons and sections on EM-grids. Although operating with long-distance lenses, this system achieves a very high light sensitivity. Our instrumental set-up and operating workflow are designed to investigate rare events in large tissue volumes. Applications range from studies of individual immune, stem and cancer cells to the investigation of non-uniform subcellular processes. As a use case, we present the ultrastructure of a single membrane repair patch on a muscle fiber in intact muscle in a whole animal context.
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Affiliation(s)
- Nicolas Lemercier
- Institut de Génétique et de Biologie Moléculaire et Cellulaire, 1, rue Laurent Fries, 67404, Illkirch, France.,Centre National de la Recherche Scientifique, UMR7104, 1, rue Laurent Fries, 67404, Illkirch, France.,Institut National de la Santé et de la Recherche Médicale, U964, 1, rue Laurent Fries, 67404, Illkirch, France.,Université de Strasbourg, 1, rue Laurent Fries, 67404, Illkirch, France
| | - Volker Middel
- Institute of Toxicology and Genetics, Karlsruhe Institute of Technology, Hermann-von-Helmholtz-Platz 1, 76344, Eggenstein-Leopoldshafen, Germany
| | - Didier Hentsch
- Institut de Génétique et de Biologie Moléculaire et Cellulaire, 1, rue Laurent Fries, 67404, Illkirch, France.,Centre National de la Recherche Scientifique, UMR7104, 1, rue Laurent Fries, 67404, Illkirch, France.,Institut National de la Santé et de la Recherche Médicale, U964, 1, rue Laurent Fries, 67404, Illkirch, France.,Université de Strasbourg, 1, rue Laurent Fries, 67404, Illkirch, France
| | - Serge Taubert
- Institut de Génétique et de Biologie Moléculaire et Cellulaire, 1, rue Laurent Fries, 67404, Illkirch, France.,Centre National de la Recherche Scientifique, UMR7104, 1, rue Laurent Fries, 67404, Illkirch, France.,Institut National de la Santé et de la Recherche Médicale, U964, 1, rue Laurent Fries, 67404, Illkirch, France.,Université de Strasbourg, 1, rue Laurent Fries, 67404, Illkirch, France
| | - Masanari Takamiya
- Institute of Toxicology and Genetics, Karlsruhe Institute of Technology, Hermann-von-Helmholtz-Platz 1, 76344, Eggenstein-Leopoldshafen, Germany
| | - Tanja Beil
- Institute of Toxicology and Genetics, Karlsruhe Institute of Technology, Hermann-von-Helmholtz-Platz 1, 76344, Eggenstein-Leopoldshafen, Germany
| | - Jean-Luc Vonesch
- Institut de Génétique et de Biologie Moléculaire et Cellulaire, 1, rue Laurent Fries, 67404, Illkirch, France.,Centre National de la Recherche Scientifique, UMR7104, 1, rue Laurent Fries, 67404, Illkirch, France.,Institut National de la Santé et de la Recherche Médicale, U964, 1, rue Laurent Fries, 67404, Illkirch, France.,Université de Strasbourg, 1, rue Laurent Fries, 67404, Illkirch, France
| | - Tilo Baumbach
- Laboratory for Applications of Synchrotron Radiation, Karlsruhe Institute of Technology, Kaiserstr. 12, 76131, Karlsruhe, Germany
| | - Patrick Schultz
- Institut de Génétique et de Biologie Moléculaire et Cellulaire, 1, rue Laurent Fries, 67404, Illkirch, France. .,Centre National de la Recherche Scientifique, UMR7104, 1, rue Laurent Fries, 67404, Illkirch, France. .,Institut National de la Santé et de la Recherche Médicale, U964, 1, rue Laurent Fries, 67404, Illkirch, France. .,Université de Strasbourg, 1, rue Laurent Fries, 67404, Illkirch, France.
| | - Claude Antony
- Institut de Génétique et de Biologie Moléculaire et Cellulaire, 1, rue Laurent Fries, 67404, Illkirch, France.,Centre National de la Recherche Scientifique, UMR7104, 1, rue Laurent Fries, 67404, Illkirch, France.,Institut National de la Santé et de la Recherche Médicale, U964, 1, rue Laurent Fries, 67404, Illkirch, France.,Université de Strasbourg, 1, rue Laurent Fries, 67404, Illkirch, France.,Institute of Toxicology and Genetics, Karlsruhe Institute of Technology, Hermann-von-Helmholtz-Platz 1, 76344, Eggenstein-Leopoldshafen, Germany
| | - Uwe Strähle
- Institute of Toxicology and Genetics, Karlsruhe Institute of Technology, Hermann-von-Helmholtz-Platz 1, 76344, Eggenstein-Leopoldshafen, Germany.
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48
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Inducing fluorescence of uranyl acetate as a dual-purpose contrast agent for correlative light-electron microscopy with nanometre precision. Sci Rep 2017; 7:10442. [PMID: 28874723 PMCID: PMC5585351 DOI: 10.1038/s41598-017-10905-x] [Citation(s) in RCA: 10] [Impact Index Per Article: 1.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/13/2017] [Accepted: 08/16/2017] [Indexed: 01/24/2023] Open
Abstract
Correlative light-electron microscopy (CLEM) combines the high spatial resolution of transmission electron microscopy (TEM) with the capability of fluorescence light microscopy (FLM) to locate rare or transient cellular events within a large field of view. CLEM is therefore a powerful technique to study cellular processes. Aligning images derived from both imaging modalities is a prerequisite to correlate the two microscopy data sets, and poor alignment can limit interpretability of the data. Here, we describe how uranyl acetate, a commonly-used contrast agent for TEM, can be induced to fluoresce brightly at cryogenic temperatures (−195 °C) and imaged by cryoFLM using standard filter sets. This dual-purpose contrast agent can be used as a general tool for CLEM, whereby the equivalent staining allows direct correlation between fluorescence and TEM images. We demonstrate the potential of this approach by performing multi-colour CLEM of cells containing equine arteritis virus proteins tagged with either green- or red-fluorescent protein, and achieve high-precision localization of virus-induced intracellular membrane modifications. Using uranyl acetate as a dual-purpose contrast agent, we achieve an image alignment precision of ~30 nm, twice as accurate as when using fiducial beads, which will be essential for combining TEM with the evolving field of super-resolution light microscopy.
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Correlative super-resolution fluorescence and electron microscopy using conventional fluorescent proteins in vacuo. J Struct Biol 2017; 199:120-131. [PMID: 28576556 PMCID: PMC5531056 DOI: 10.1016/j.jsb.2017.05.013] [Citation(s) in RCA: 47] [Impact Index Per Article: 6.7] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/15/2016] [Revised: 04/27/2017] [Accepted: 05/29/2017] [Indexed: 12/12/2022]
Abstract
Super-resolution light microscopy, correlative light and electron microscopy, and volume electron microscopy are revolutionising the way in which biological samples are examined and understood. Here, we combine these approaches to deliver super-accurate correlation of fluorescent proteins to cellular structures. We show that YFP and GFP have enhanced blinking properties when embedded in acrylic resin and imaged under partial vacuum, enabling in vacuo single molecule localisation microscopy. In conventional section-based correlative microscopy experiments, the specimen must be moved between imaging systems and/or further manipulated for optimal viewing. These steps can introduce undesirable alterations in the specimen, and complicate correlation between imaging modalities. We avoided these issues by using a scanning electron microscope with integrated optical microscope to acquire both localisation and electron microscopy images, which could then be precisely correlated. Collecting data from ultrathin sections also improved the axial resolution and signal-to-noise ratio of the raw localisation microscopy data. Expanding data collection across an array of sections will allow 3-dimensional correlation over unprecedented volumes. The performance of this technique is demonstrated on vaccinia virus (with YFP) and diacylglycerol in cellular membranes (with GFP).
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50
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Relocation is the key to successful correlative fluorescence and scanning electron microscopy. Methods Cell Biol 2017; 140:215-244. [PMID: 28528635 DOI: 10.1016/bs.mcb.2017.03.013] [Citation(s) in RCA: 4] [Impact Index Per Article: 0.6] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 02/07/2023]
Abstract
In this chapter the authors report on an automated hardware and software solution enabling swift correlative sample array mapping of fluorescently stained molecules within cells and tissues across length scales. Samples are first observed utilizing wide-field optical and fluorescence microscopy, followed by scanning electron microscopy, using calibration points on a dedicated sample-relocation holder. We investigated HeLa cells in vitro, fluorescently labeled for monosialoganglioside one (GM-1), across both imaging platforms within tens of minutes of initial sample preparation. This resulted in a high-throughput and high spatially resolved correlative fluorescence and electron microscopy analysis and allowed us to collect complementary nanoscopic information on the molecular and structural composition of two differently distinct HeLa cell populations expressing different levels of GM-1. Furthermore, using the small zebrafish animal model Danio rerio, we showed the versatility and relocation accuracy of the sample-relocation holder to locate fluo-tagged macromolecular complexes within large volumes using long ribbons of serial tissue sections. The subsequent electron microscopy imaging of the tissue arrays of interest enabled the generation of correlated information on the fine distribution of albumin within hepatic and kidney tissue. Our approach underpins the merits that an automated sample-relocation holder solution brings in support of results-driven research, where relevant biological questions can be answered, and high-throughput data can be generated in a rigorous statistical manner.
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