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Akkaya M, Al Souz J, Williams D, Kamdar R, Kamenyeva O, Kabat J, Shevach E, Akkaya B. Illuminating T cell-dendritic cell interactions in vivo by FlAsHing antigens. eLife 2024; 12:RP91809. [PMID: 38236633 PMCID: PMC10945603 DOI: 10.7554/elife.91809] [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: 01/19/2024] Open
Abstract
Delineating the complex network of interactions between antigen-specific T cells and antigen presenting cells (APCs) is crucial for effective precision therapies against cancer, chronic infections, and autoimmunity. However, the existing arsenal for examining antigen-specific T cell interactions is restricted to a select few antigen-T cell receptor pairs, with limited in situ utility. This lack of versatility is largely due to the disruptive effects of reagents on the immune synapse, which hinder real-time monitoring of antigen-specific interactions. To address this limitation, we have developed a novel and versatile immune monitoring strategy by adding a short cysteine-rich tag to antigenic peptides that emits fluorescence upon binding to thiol-reactive biarsenical hairpin compounds. Our findings demonstrate the specificity and durability of the novel antigen-targeting probes during dynamic immune monitoring in vitro and in vivo. This strategy opens new avenues for biological validation of T-cell receptors with newly identified epitopes by revealing the behavior of previously unrecognized antigen-receptor pairs, expanding our understanding of T cell responses.
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Affiliation(s)
- Munir Akkaya
- Department of Internal Medicine, Division of Rheumatology and Immunology, The College of Medicine, The Ohio State UniversityColumbusUnited States
- Microbial Infection and Immunity, The Ohio State University Wexner Medical CenterColumbusUnited States
- Pelotonia Institute for Immuno-Oncology, The Ohio State UniversityColumbusUnited States
| | - Jafar Al Souz
- National Institute of Allergy and Infectious DiseasesBethesdaUnited States
| | - Daniel Williams
- National Institute of Allergy and Infectious DiseasesBethesdaUnited States
| | - Rahul Kamdar
- National Institute of Allergy and Infectious DiseasesBethesdaUnited States
| | - Olena Kamenyeva
- National Institute of Allergy and Infectious DiseasesBethesdaUnited States
| | - Juraj Kabat
- National Institute of Allergy and Infectious DiseasesBethesdaUnited States
| | - Ethan Shevach
- National Institute of Allergy and Infectious DiseasesBethesdaUnited States
| | - Billur Akkaya
- Pelotonia Institute for Immuno-Oncology, The Ohio State UniversityColumbusUnited States
- Department of Neurology, The Ohio State University Wexner Medical CenterColumbusUnited States
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2
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Akkaya M, Al Souz J, Williams D, Kamdar R, Kamenyeva O, Kabat J, Shevach EM, Akkaya B. Illuminating T cell-dendritic cell interactions in vivo by FlAsHing antigens. RESEARCH SQUARE 2023:rs.3.rs-3193191. [PMID: 37546912 PMCID: PMC10402196 DOI: 10.21203/rs.3.rs-3193191/v3] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Figures] [Subscribe] [Scholar Register] [Indexed: 01/30/2024]
Abstract
Delineating the complex network of interactions between antigen-specific T cells and antigen presenting cells (APCs) is crucial for effective precision therapies against cancer, chronic infections, and autoimmunity. However, the existing arsenal for examining antigen-specific T cell interactions is restricted to a select few antigen-T cell receptor pairs, with limited in situ utility. This lack of versatility is largely due to the disruptive effects of reagents on the immune synapse, which hinder real-time monitoring of antigen-specific interactions. To address this limitation, we have developed a novel and versatile immune monitoring strategy by adding a short cysteine-rich tag to antigenic peptides that emits fluorescence upon binding to thiol-reactive biarsenical hairpin compounds. Our findings demonstrate the specificity and durability of the novel antigen-targeting probes during dynamic immune monitoring in vitro and in vivo. This strategy opens new avenues for biological validation of T-cell receptors with newly identified epitopes by revealing the behavior of previously unrecognized antigen-receptor pairs, expanding our understanding of T cell responses.
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Affiliation(s)
- Munir Akkaya
- Department of Internal Medicine, Division of Rheumatology and Immunology, The College of Medicine, The Ohio State University, Columbus, OH 43210, USA
- Department of Microbial Infection and Immunity, The College of Medicine, The Ohio State University, Columbus, OH 43210, USA
- Pelotonia Institute for Immuno-Oncology, The James Comprehensive Cancer Center, The Ohio State University, Columbus, OH 43210, USA
| | - Jafar Al Souz
- National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, USA
| | - Daniel Williams
- National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, USA
| | - Rahul Kamdar
- National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, USA
| | - Olena Kamenyeva
- National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, USA
| | - Juraj Kabat
- National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, USA
| | - Ethan M. Shevach
- National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, USA
| | - Billur Akkaya
- Department of Neurology, The College of Medicine, The Ohio State University, Columbus, OH 43210, USA
- Department of Microbial Infection and Immunity, The College of Medicine, The Ohio State University, Columbus, OH 43210, USA
- Pelotonia Institute for Immuno-Oncology, The James Comprehensive Cancer Center, The Ohio State University, Columbus, OH 43210, USA
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3
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Akkaya M, Al Souz J, Williams D, Kamdar R, Kamenyeva O, Kabat J, Shevach EM, Akkaya B. Illuminating T cell-dendritic cell interactions in vivo by FlAsHing antigens. RESEARCH SQUARE 2023:rs.3.rs-3193191. [PMID: 37546912 PMCID: PMC10402196 DOI: 10.21203/rs.3.rs-3193191/v1] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 08/08/2023]
Abstract
Delineating the complex network of interactions between antigen-specific T cells and antigen presenting cells (APCs) is crucial for effective precision therapies against cancer, chronic infections, and autoimmunity. However, the existing arsenal for examining antigen-specific T cell interactions is restricted to a select few antigen-T cell receptor pairs, with limited in situ utility. This lack of versatility is largely due to the disruptive effects of reagents on the immune synapse, which hinder real-time monitoring of antigen-specific interactions. To address this limitation, we have developed a novel and versatile immune monitoring strategy by adding a short cysteine-rich tag to antigenic peptides that emits fluorescence upon binding to thiol-reactive biarsenical hairpin compounds. Our findings demonstrate the specificity and durability of the novel antigen-targeting probes during dynamic immune monitoring in vitro and in vivo. This strategy opens new avenues for biological validation of T-cell receptors with newly identified epitopes by revealing the behavior of previously unrecognized antigen-receptor pairs, expanding our understanding of T cell responses.
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Affiliation(s)
- Munir Akkaya
- Department of Internal Medicine, Division of Rheumatology and Immunology, The College of Medicine, The Ohio State University, Columbus, OH 43210, USA
- Department of Microbial Infection and Immunity, The College of Medicine, The Ohio State University, Columbus, OH 43210, USA
- Pelotonia Institute for Immuno-Oncology, The James Comprehensive Cancer Center, The Ohio State University, Columbus, OH 43210, USA
| | - Jafar Al Souz
- National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, USA
| | - Daniel Williams
- National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, USA
| | - Rahul Kamdar
- National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, USA
| | - Olena Kamenyeva
- National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, USA
| | - Juraj Kabat
- National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, USA
| | - Ethan M. Shevach
- National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, USA
| | - Billur Akkaya
- Department of Neurology, The College of Medicine, The Ohio State University, Columbus, OH 43210, USA
- Department of Microbial Infection and Immunity, The College of Medicine, The Ohio State University, Columbus, OH 43210, USA
- Pelotonia Institute for Immuno-Oncology, The James Comprehensive Cancer Center, The Ohio State University, Columbus, OH 43210, USA
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4
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Maksymenko K, Maurer A, Aghaallaei N, Barry C, Borbarán-Bravo N, Ullrich T, Dijkstra TM, Hernandez Alvarez B, Müller P, Lupas AN, Skokowa J, ElGamacy M. The design of functional proteins using tensorized energy calculations. CELL REPORTS METHODS 2023; 3:100560. [PMID: 37671023 PMCID: PMC10475850 DOI: 10.1016/j.crmeth.2023.100560] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Received: 01/22/2023] [Revised: 05/25/2023] [Accepted: 07/21/2023] [Indexed: 09/07/2023]
Abstract
In protein design, the energy associated with a huge number of sequence-conformer perturbations has to be routinely estimated. Hence, enhancing the throughput and accuracy of these energy calculations can profoundly improve design success rates and enable tackling more complex design problems. In this work, we explore the possibility of tensorizing the energy calculations and apply them in a protein design framework. We use this framework to design enhanced proteins with anti-cancer and radio-tracing functions. Particularly, we designed multispecific binders against ligands of the epidermal growth factor receptor (EGFR), where the tested design could inhibit EGFR activity in vitro and in vivo. We also used this method to design high-affinity Cu2+ binders that were stable in serum and could be readily loaded with copper-64 radionuclide. The resulting molecules show superior functional properties for their respective applications and demonstrate the generalizable potential of the described protein design approach.
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Affiliation(s)
- Kateryna Maksymenko
- Department of Protein Evolution, Max Planck Institute for Biology, 72076 Tübingen, Germany
- Friedrich Miescher Laboratory of the Max Planck Society, 72076 Tübingen, Germany
| | - Andreas Maurer
- Werner Siemens Imaging Center, Department of Preclinical Imaging and Radiopharmacy, Eberhard Karls University, 72076 Tübingen, Germany
- Cluster of Excellence iFIT (EXC 2180) “Image Guided and Functionally Instructed Tumor Therapies,” Eberhard Karls University, 72076 Tübingen, Germany
| | - Narges Aghaallaei
- Division of Translational Oncology, University Hospital Tübingen, 72076 Tübingen, Germany
| | - Caroline Barry
- Department of Protein Evolution, Max Planck Institute for Biology, 72076 Tübingen, Germany
- Krieger School of Arts and Sciences, Johns Hopkins University, Washington, DC 20036, USA
| | - Natalia Borbarán-Bravo
- Division of Translational Oncology, University Hospital Tübingen, 72076 Tübingen, Germany
| | - Timo Ullrich
- Department of Protein Evolution, Max Planck Institute for Biology, 72076 Tübingen, Germany
- Friedrich Miescher Laboratory of the Max Planck Society, 72076 Tübingen, Germany
| | - Tjeerd M.H. Dijkstra
- Department of Protein Evolution, Max Planck Institute for Biology, 72076 Tübingen, Germany
- Department for Women’s Health, University Hospital Tübingen, 72076 Tübingen, Germany
- Translational Bioinformatics, University Hospital Tübingen, 72072 Tübingen, Germany
| | | | - Patrick Müller
- Friedrich Miescher Laboratory of the Max Planck Society, 72076 Tübingen, Germany
| | - Andrei N. Lupas
- Department of Protein Evolution, Max Planck Institute for Biology, 72076 Tübingen, Germany
| | - Julia Skokowa
- Division of Translational Oncology, University Hospital Tübingen, 72076 Tübingen, Germany
| | - Mohammad ElGamacy
- Department of Protein Evolution, Max Planck Institute for Biology, 72076 Tübingen, Germany
- Friedrich Miescher Laboratory of the Max Planck Society, 72076 Tübingen, Germany
- Division of Translational Oncology, University Hospital Tübingen, 72076 Tübingen, Germany
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5
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Identifying Active Progeny Virus Particles in Formalin-Fixed, Paraffin-Embedded Sections Using Correlative Light and Scanning Electron Microscopy. J Transl Med 2023; 103:100020. [PMID: 36748195 DOI: 10.1016/j.labinv.2022.100020] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/07/2022] [Revised: 10/06/2022] [Accepted: 10/10/2022] [Indexed: 01/18/2023] Open
Abstract
Immunohistochemical analysis of formalin-fixed paraffin-embedded (FFPE) tissue blocks is routinely used to identify virus-infected cells. However, detecting virus particles in FFPE sections using light microscopy is difficult because of the light diffraction resolution limitations of an optical microscope. In this study, light microscopy and field emission scanning electron microscopy were performed to observe 3-dimensional virus particles in FFPE sections in a nondestructive manner using NanoSuit or osmium conductive treatment methods. The virus particles in FFPE sections were immunostained with specific antibodies against the surface antigens of the viral particles and stained with 3,3'-diaminobenzidine. A metal solution (0.2% gold chloride or 2% osmium tetroxide) was applied to enhance the 3,3'-diaminobenzidine-stained area. This procedure is nondestructive for FFPE sections and is a simpler method than transmission electron microscopy. To validate the applicability of this technique, we performed 3-dimensional imaging of the virus particles of different sizes, such as human papillomavirus, cytomegalovirus, and varicella-zoster virus. Furthermore, ultrathin sections from the FFPE sections that were observed to harbor viral particles using field emission scanning electron microscopy were prepared and assessed using transmission electron microscopy. In the correlative areas, transmission electron microscopy confirmed the presence of large numbers of virus particles. These results indicated that the combination of marking viral particles with 3,3'-diaminobenzidine/metal staining and conductive treatment can identify active progeny virus particles in FFPE sections using scanning electron microscopy. This easy correlative imaging of field emission scanning electron microscopy of the identical area of FFPE in light microscopy may help elucidate new pathological mechanisms of virus-related diseases.
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6
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Proteomic mapping and optogenetic manipulation of membrane contact sites. Biochem J 2022; 479:1857-1875. [PMID: 36111979 PMCID: PMC9555801 DOI: 10.1042/bcj20220382] [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: 07/18/2022] [Revised: 08/31/2022] [Accepted: 09/01/2022] [Indexed: 11/17/2022]
Abstract
Membrane contact sites (MCSs) mediate crucial physiological processes in eukaryotic cells, including ion signaling, lipid metabolism, and autophagy. Dysregulation of MCSs is closely related to various diseases, such as type 2 diabetes mellitus (T2DM), neurodegenerative diseases, and cancers. Visualization, proteomic mapping and manipulation of MCSs may help the dissection of the physiology and pathology MCSs. Recent technical advances have enabled better understanding of the dynamics and functions of MCSs. Here we present a summary of currently known functions of MCSs, with a focus on optical approaches to visualize and manipulate MCSs, as well as proteomic mapping within MCSs.
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7
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Cheung CY, Huang TT, Chow N, Zhang S, Zhao Y, Chau MP, Chan WC, Wong CCL, Boassa D, Phan S, Ellisman MH, Yates JR, Xu S, Yu Z, Zhang Y, Zhang R, Ng LL, Ko BCB. Unconventional tonicity-regulated nuclear trafficking of NFAT5 mediated by KPNB1, XPOT and RUVBL2. J Cell Sci 2022; 135:275560. [PMID: 35635291 PMCID: PMC9377714 DOI: 10.1242/jcs.259280] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/18/2021] [Accepted: 05/20/2022] [Indexed: 11/20/2022] Open
Abstract
NFAT5 is the only known mammalian tonicity-responsive transcription factor with essential role in cellular adaptation to hypertonic stress. It is also implicated in diverse physiological and pathological processes. NFAT5 activity is tightly regulated by extracellular tonicity, but the underlying mechanisms remain elusive. We demonstrated that NFAT5 enters the nucleus via the nuclear pore complex. We found that NFAT5 utilizes a unique nuclear localization signal (NFAT5-NLS) for nuclear import. siRNA screening revealed that only karyopherin β1 (KPNB1), but not karyopherin alpha, is responsible for the nuclear import of NFAT5 via direct interaction with the NFAT5-NLS. Proteomics analysis and siRNA screening further revealed that nuclear export of NFAT5 under hypotonicity is driven by Exportin-T, where the process requires RuvB-Like AAA type ATPase 2 (RUVBL2) as an indispensable chaperone. Our findings have identified an unconventional tonicity-dependent nucleocytoplasmic trafficking pathway for NFAT5, a critical step in orchestrating rapid cellular adaptation to change in extracellular tonicity. These findings offer an opportunity for the development of novel NFAT5 targeting strategies that are potentially useful for the treatment of diseases associated with NFAT5 dysregulation.
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Affiliation(s)
- Chris Y Cheung
- Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hong Kong, China.,State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hong Kong, China
| | - Ting-Ting Huang
- Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hong Kong, China.,State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hong Kong, China
| | - Ning Chow
- Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hong Kong, China.,State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hong Kong, China
| | - Shuqi Zhang
- Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hong Kong, China.,State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hong Kong, China
| | - Yanxiang Zhao
- Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hong Kong, China.,State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hong Kong, China
| | - Mary P Chau
- Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hong Kong, China.,State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hong Kong, China
| | - Wing Cheung Chan
- Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hong Kong, China.,State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hong Kong, China
| | - Catherine C L Wong
- Center for Precision Medicine Muti-Omics Research, Health Science Center, Peking University, China Clinical Laboratory Department, The Cancer Hospital of the University of Chinese Academy of Sciences, Beijing, China
| | - Daniela Boassa
- Department of Neurosciences, University of California, San Diego, USA.,Center for Research in Biological Systems, National Center for Microscopy and Imaging Research, University of California, San Diego, La Jolla, California, USA
| | - Sebastien Phan
- Department of Neurosciences, University of California, San Diego, USA.,Center for Research in Biological Systems, National Center for Microscopy and Imaging Research, University of California, San Diego, La Jolla, California, USA
| | - Mark H Ellisman
- Department of Neurosciences, University of California, San Diego, USA.,Center for Research in Biological Systems, National Center for Microscopy and Imaging Research, University of California, San Diego, La Jolla, California, USA
| | - John R Yates
- Department of Chemical Physiology, The Scripps Research Institute, La Jolla, California, USA
| | - SongXiao Xu
- The Clinical Laboratory Department, The Cancer Hospital of the University of Chinese Academy of Sciences (Zhejiang Cancer Hospital), Institute of Basic Medicine and Cancer, Chinese Academy of Sciences, Hangzhou, Zhejiang, China
| | - Zicheng Yu
- Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hong Kong, China.,State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hong Kong, China
| | - Yajing Zhang
- Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hong Kong, China.,State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hong Kong, China
| | - Rui Zhang
- Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hong Kong, China.,State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hong Kong, China
| | - Ling Ling Ng
- Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hong Kong, China.,State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hong Kong, China
| | - Ben C B Ko
- Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hong Kong, China.,State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hong Kong, China
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Hübner B, von Otter E, Ahsan B, Wee ML, Henriksson S, Ludwig A, Sandin S. Ultrastructure and nuclear architecture of telomeric chromatin revealed by correlative light and electron microscopy. Nucleic Acids Res 2022; 50:5047-5063. [PMID: 35489064 PMCID: PMC9122609 DOI: 10.1093/nar/gkac309] [Citation(s) in RCA: 6] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/19/2021] [Revised: 04/07/2022] [Accepted: 04/19/2022] [Indexed: 11/25/2022] Open
Abstract
Telomeres, the ends of linear chromosomes, are composed of repetitive DNA sequences, histones and a protein complex called shelterin. How DNA is packaged at telomeres is an outstanding question in the field with significant implications for human health and disease. Here, we studied the architecture of telomeres and their spatial association with other chromatin domains in different cell types using correlative light and electron microscopy. To this end, the shelterin protein TRF1 or TRF2 was fused in tandem to eGFP and the peroxidase APEX2, which provided a selective and electron-dense label to interrogate telomere organization by transmission electron microscopy, electron tomography and scanning electron microscopy. Together, our work reveals, for the first time, ultrastructural insight into telomere architecture. We show that telomeres are composed of a dense and highly compacted mesh of chromatin fibres. In addition, we identify marked differences in telomere size, shape and chromatin compaction between cancer and non-cancer cells and show that telomeres are in direct contact with other heterochromatin regions. Our work resolves the internal architecture of telomeres with unprecedented resolution and advances our understanding of how telomeres are organized in situ.
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Affiliation(s)
- Barbara Hübner
- School of Biological Sciences, Nanyang Technological University, 60 Nanyang Drive, Singapore 637551, Singapore.,NTU Institute of Structural Biology, Nanyang Technological University, 59 Nanyang Drive, Singapore 636921, Singapore
| | - Eric von Otter
- School of Biological Sciences, Nanyang Technological University, 60 Nanyang Drive, Singapore 637551, Singapore.,NTU Institute of Structural Biology, Nanyang Technological University, 59 Nanyang Drive, Singapore 636921, Singapore
| | - Bilal Ahsan
- School of Biological Sciences, Nanyang Technological University, 60 Nanyang Drive, Singapore 637551, Singapore.,NTU Institute of Structural Biology, Nanyang Technological University, 59 Nanyang Drive, Singapore 636921, Singapore
| | - Mei Ling Wee
- School of Biological Sciences, Nanyang Technological University, 60 Nanyang Drive, Singapore 637551, Singapore.,NTU Institute of Structural Biology, Nanyang Technological University, 59 Nanyang Drive, Singapore 636921, Singapore
| | - Sara Henriksson
- Umeå Centre for Electron Microscopy, Umeå University, Chemical Biological Centre (KBC) Building, Linnaeus väg 6, SE-90736 Umeå, Sweden
| | - Alexander Ludwig
- School of Biological Sciences, Nanyang Technological University, 60 Nanyang Drive, Singapore 637551, Singapore.,NTU Institute of Structural Biology, Nanyang Technological University, 59 Nanyang Drive, Singapore 636921, Singapore
| | - Sara Sandin
- School of Biological Sciences, Nanyang Technological University, 60 Nanyang Drive, Singapore 637551, Singapore.,NTU Institute of Structural Biology, Nanyang Technological University, 59 Nanyang Drive, Singapore 636921, Singapore
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9
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Kimmel J, Kehrer J, Frischknecht F, Spielmann T. Proximity-dependent biotinylation approaches to study apicomplexan biology. Mol Microbiol 2021; 117:553-568. [PMID: 34587292 DOI: 10.1111/mmi.14815] [Citation(s) in RCA: 7] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/20/2021] [Accepted: 09/20/2021] [Indexed: 11/28/2022]
Abstract
In the last 10 years, proximity-dependent biotinylation (PDB) techniques greatly expanded the ability to study protein environments in the living cell that range from specific protein complexes to entire compartments. This is achieved by using enzymes such as BirA* and APEX that are fused to proteins of interest and biotinylate proteins in their proximity. PDB techniques are now also increasingly used in apicomplexan parasites. In this review, we first give an overview of the main PDB approaches and how they compare with other techniques that address similar questions. PDB is particularly valuable to detect weak or transient protein associations under physiological conditions and to study cellular structures that are difficult to purify or have a poorly understood protein composition. We also highlight new developments such as novel smaller or faster-acting enzyme variants and conditional PDB approaches, providing improvements in both temporal and spatial resolution which may offer broader application possibilities useful in apicomplexan research. In the second part, we review work using PDB techniques in apicomplexan parasites and how this expanded our knowledge about these medically important parasites.
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Affiliation(s)
- Jessica Kimmel
- Bernhard Nocht Institute for Tropical Medicine, Hamburg, Germany
| | - Jessica Kehrer
- Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany.,German Center for Infectious Disease Research, DZIF, Heidelberg, Germany
| | - Friedrich Frischknecht
- Integrative Parasitology, Center for Infectious Diseases, Heidelberg University Medical School, Heidelberg, Germany.,German Center for Infectious Disease Research, DZIF, Heidelberg, Germany
| | - Tobias Spielmann
- Bernhard Nocht Institute for Tropical Medicine, Hamburg, Germany
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10
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Stelate A, Tihlaříková E, Schwarzerová K, Neděla V, Petrášek J. Correlative Light-Environmental Scanning Electron Microscopy of Plasma Membrane Efflux Carriers of Plant Hormone Auxin. Biomolecules 2021; 11:1407. [PMID: 34680040 PMCID: PMC8533460 DOI: 10.3390/biom11101407] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.7] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/31/2021] [Revised: 09/21/2021] [Accepted: 09/22/2021] [Indexed: 12/17/2022] Open
Abstract
Fluorescence light microscopy provided convincing evidence for the domain organization of plant plasma membrane (PM) proteins. Both peripheral and integral PM proteins show an inhomogeneous distribution within the PM. However, the size of PM nanodomains and protein clusters is too small to accurately determine their dimensions and nano-organization using routine confocal fluorescence microscopy and super-resolution methods. To overcome this limitation, we have developed a novel correlative light electron microscopy method (CLEM) using total internal reflection fluorescence microscopy (TIRFM) and advanced environmental scanning electron microscopy (A-ESEM). Using this technique, we determined the number of auxin efflux carriers from the PINFORMED (PIN) family (NtPIN3b-GFP) within PM nanodomains of tobacco cell PM ghosts. Protoplasts were attached to coverslips and immunostained with anti-GFP primary antibody and secondary antibody conjugated to fluorochrome and gold nanoparticles. After imaging the nanodomains within the PM with TIRFM, the samples were imaged with A-ESEM without further processing, and quantification of the average number of molecules within the nanodomain was performed. Without requiring any post-fixation and coating procedures, this method allows to study details of the organization of auxin carriers and other plant PM proteins.
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Affiliation(s)
- Ayoub Stelate
- Department of Experimental Plant Biology, Faculty of Science, Charles University, Viničná 5, 128 44 Prague 2, Czech Republic; (A.S.); (K.S.)
| | - Eva Tihlaříková
- Institute of Scientific Instruments, Academy of Sciences of the Czech Republic, Královopolská 147, 612 64 Brno, Czech Republic; (E.T.); (V.N.)
| | - Kateřina Schwarzerová
- Department of Experimental Plant Biology, Faculty of Science, Charles University, Viničná 5, 128 44 Prague 2, Czech Republic; (A.S.); (K.S.)
| | - Vilém Neděla
- Institute of Scientific Instruments, Academy of Sciences of the Czech Republic, Královopolská 147, 612 64 Brno, Czech Republic; (E.T.); (V.N.)
| | - Jan Petrášek
- Department of Experimental Plant Biology, Faculty of Science, Charles University, Viničná 5, 128 44 Prague 2, Czech Republic; (A.S.); (K.S.)
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11
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Harper CB, Smillie KJ. Current molecular approaches to investigate pre-synaptic dysfunction. J Neurochem 2021; 157:107-129. [PMID: 33544872 DOI: 10.1111/jnc.15316] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/06/2020] [Revised: 01/29/2021] [Accepted: 02/01/2021] [Indexed: 12/19/2022]
Abstract
Over the course of the last few decades it has become clear that many neurodevelopmental and neurodegenerative disorders have a synaptic defect, which contributes to pathogenicity. A rise in new techniques, and in particular '-omics'-based methods providing large datasets, has led to an increase in potential proteins and pathways implicated in synaptic function and related disorders. Additionally, advancements in imaging techniques have led to the recent discovery of alternative modes of synaptic vesicle recycling. This has resulted in a lack of clarity over the precise role of different pathways in maintaining synaptic function and whether these new pathways are dysfunctional in neurodevelopmental and neurodegenerative disorders. A greater understanding of the molecular detail of pre-synaptic function in health and disease is key to targeting new proteins and pathways for novel treatments and the variety of new techniques currently available provides an ideal opportunity to investigate these functions. This review focuses on techniques to interrogate pre-synaptic function, concentrating mainly on synaptic vesicle recycling. It further examines techniques to determine the underlying molecular mechanism of pre-synaptic dysfunction and discusses methods to identify molecular targets, along with protein-protein interactions and cellular localization. In combination, these techniques will provide an expanding and more complete picture of pre-synaptic function. With the application of recent technological advances, we are able to resolve events with higher spatial and temporal resolution, leading research towards a greater understanding of dysfunction at the presynapse and the role it plays in pathogenicity.
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Affiliation(s)
- Callista B Harper
- Centre for Discovery Brain Sciences, University of Edinburgh, Scotland, UK
| | - Karen J Smillie
- Centre for Discovery Brain Sciences, University of Edinburgh, Scotland, UK
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12
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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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13
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Bayguinov PO, Fisher MR, Fitzpatrick JAJ. Assaying three-dimensional cellular architecture using X-ray tomographic and correlated imaging approaches. J Biol Chem 2020; 295:15782-15793. [PMID: 32938716 PMCID: PMC7667966 DOI: 10.1074/jbc.rev120.009633] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/01/2020] [Revised: 09/15/2020] [Indexed: 12/16/2022] Open
Abstract
Much of our understanding of the spatial organization of and interactions between cellular organelles and macromolecular complexes has been the result of imaging studies utilizing either light- or electron-based microscopic analyses. These classical approaches, while insightful, are nonetheless limited either by restrictions in resolution or by the sheer complexity of generating multidimensional data. Recent advances in the use and application of X-rays to acquire micro- and nanotomographic data sets offer an alternative methodology to visualize cellular architecture at the nanoscale. These new approaches allow for the subcellular analyses of unstained vitrified cells and three-dimensional localization of specific protein targets and have served as an essential tool in bridging light and electron correlative microscopy experiments. Here, we review the theory, instrumentation details, acquisition principles, and applications of both soft X-ray tomography and X-ray microscopy and how the use of these techniques offers a succinct means of analyzing three-dimensional cellular architecture. We discuss some of the recent work that has taken advantage of these approaches and detail how they have become integral in correlative microscopy workflows.
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Affiliation(s)
- Peter O Bayguinov
- Washington University Center for Cellular Imaging, Washington University School of Medicine, Saint Louis, Missouri, USA
| | - Max R Fisher
- Washington University Center for Cellular Imaging, Washington University School of Medicine, Saint Louis, Missouri, USA
| | - James A J Fitzpatrick
- Washington University Center for Cellular Imaging, Washington University School of Medicine, Saint Louis, Missouri, USA; Departments of Cell Biology and Physiology and Neuroscience, Washington University School of Medicine, Saint Louis, Missouri, USA; Department of Biomedical Engineering, Washington University in Saint Louis, Saint Louis, Missouri, USA.
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14
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Kremer A, VAN Hamme E, Bonnardel J, Borghgraef P, GuÉrin CJ, Guilliams M, Lippens S. A workflow for 3D-CLEM investigating liver tissue. J Microsc 2020; 281:231-242. [PMID: 33034376 DOI: 10.1111/jmi.12967] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/06/2020] [Revised: 09/22/2020] [Accepted: 10/07/2020] [Indexed: 12/31/2022]
Abstract
Correlative light and electron microscopy (CLEM) is a method used to investigate the exact same region in both light and electron microscopy (EM) in order to add ultrastructural information to a light microscopic (usually fluorescent) signal. Workflows combining optical or fluorescent data with electron microscopic images are complex, hence there is a need to communicate detailed protocols and share tips & tricks for successful application of these methods. With the development of volume-EM techniques such as serial blockface scanning electron microscopy (SBF-SEM) and Focussed Ion Beam-SEM, correlation in three dimensions has become more efficient. Volume electron microscopy allows automated acquisition of serial section imaging data that can be reconstructed in three dimensions (3D) to provide a detailed, geometrically accurate view of cellular ultrastructure. In addition, combining volume-EM with high-resolution light microscopy (LM) techniques decreases the resolution gap between LM and EM, making retracing of a region of interest and eventual overlays more straightforward. Here, we present a workflow for 3D CLEM on mouse liver, combining high-resolution confocal microscopy with SBF-SEM. In this workflow, we have made use of two types of landmarks: (1) near infrared laser branding marks to find back the region imaged in LM in the electron microscope and (2) landmarks present in the tissue but independent of the cell or structure of interest to make overlay images of LM and EM data. Using this approach, we were able to make accurate 3D-CLEM overlays of liver tissue and correlate the fluorescent signal to the ultrastructural detail provided by the electron microscope. This workflow can be adapted for other dense cellular tissues and thus act as a guide for other three-dimensional correlative studies. LAY DESCRIPTION: As cells and tissues exist in three dimensions, microscopy techniques have been developed to image samples, in 3D, at the highest possible detail. In light microscopy, fluorescent probes are used to identify specific proteins or structures either in live samples, (providing dynamic information), or in fixed slices of tissue. A disadvantage of fluorescence microscopy is that only the labeled proteins/structures are visible, while their cellular context remains hidden. Electron microscopy is able to image biological samples at high resolution and has the advantage that all structures in the tissue are visible at nanometer (10-9 m) resolution. Disadvantages of this technique are that it is more difficult to label a single structure and that the samples must be imaged under high vacuum, so biological samples need to be fixed and embedded in a plastic resin to stay as close to their natural state as possible inside the microscope. Correlative Light and Electron Microscopy aims to combine the advantages of both light and electron microscopy on the same sample. This results in datasets where fluorescent labels can be combined with the high-resolution contextual information provided by the electron microscope. In this study we present a workflow to guide a tissue sample from the light microscope to the electron microscope and image the ultra-structure of a specific cell type in the liver. In particular we focus on the incorporation of fiducial markers during the sample preparation to help navigate through the tissue in 3D in both microscopes. One sample is followed throughout the workflow to visualize the important steps in the process, showing the final result; a dataset combining fluorescent labels with ultra-structural detail.
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Affiliation(s)
- A Kremer
- VIB BioImaging Core, Technologiepark 71, Ghent, 9052, Belgium.,VIB Center for Inflammation Research, Technologiepark 71, Ghent, 9052, Belgium.,Department of Biomedical Molecular Biology, Ghent University, Technologiepark 71, Ghent, 9052, Belgium
| | - E VAN Hamme
- VIB BioImaging Core, Technologiepark 71, Ghent, 9052, Belgium.,VIB Center for Inflammation Research, Technologiepark 71, Ghent, 9052, Belgium.,Department of Biomedical Molecular Biology, Ghent University, Technologiepark 71, Ghent, 9052, Belgium
| | - J Bonnardel
- VIB BioImaging Core, Technologiepark 71, Ghent, 9052, Belgium.,VIB Center for Inflammation Research, Technologiepark 71, Ghent, 9052, Belgium.,Department of Biomedical Molecular Biology, Ghent University, Technologiepark 71, Ghent, 9052, Belgium
| | - P Borghgraef
- VIB BioImaging Core, Technologiepark 71, Ghent, 9052, Belgium.,VIB Center for Inflammation Research, Technologiepark 71, Ghent, 9052, Belgium.,Department of Biomedical Molecular Biology, Ghent University, Technologiepark 71, Ghent, 9052, Belgium
| | - C J GuÉrin
- VIB BioImaging Core, Technologiepark 71, Ghent, 9052, Belgium.,VIB Center for Inflammation Research, Technologiepark 71, Ghent, 9052, Belgium.,Department of Biomedical Molecular Biology, Ghent University, Technologiepark 71, Ghent, 9052, Belgium
| | - M Guilliams
- VIB BioImaging Core, Technologiepark 71, Ghent, 9052, Belgium.,VIB Center for Inflammation Research, Technologiepark 71, Ghent, 9052, Belgium.,Department of Biomedical Molecular Biology, Ghent University, Technologiepark 71, Ghent, 9052, Belgium
| | - S Lippens
- VIB BioImaging Core, Technologiepark 71, Ghent, 9052, Belgium.,VIB Center for Inflammation Research, Technologiepark 71, Ghent, 9052, Belgium.,Department of Biomedical Molecular Biology, Ghent University, Technologiepark 71, Ghent, 9052, Belgium
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15
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Bosch JA, Chen CL, Perrimon N. Proximity-dependent labeling methods for proteomic profiling in living cells: An update. WILEY INTERDISCIPLINARY REVIEWS-DEVELOPMENTAL BIOLOGY 2020; 10:e392. [PMID: 32909689 DOI: 10.1002/wdev.392] [Citation(s) in RCA: 45] [Impact Index Per Article: 11.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Subscribe] [Scholar Register] [Received: 02/13/2020] [Revised: 06/11/2020] [Accepted: 07/01/2020] [Indexed: 12/14/2022]
Abstract
Characterizing the proteome composition of organelles and subcellular regions of living cells can facilitate the understanding of cellular organization as well as protein interactome networks. Proximity labeling-based methods coupled with mass spectrometry (MS) offer a high-throughput approach for systematic analysis of spatially restricted proteomes. Proximity labeling utilizes enzymes that generate reactive radicals to covalently tag neighboring proteins. The tagged endogenous proteins can then be isolated for further analysis by MS. To analyze protein-protein interactions or identify components that localize to discrete subcellular compartments, spatial expression is achieved by fusing the enzyme to specific proteins or signal peptides that target to particular subcellular regions. Although these technologies have only been introduced recently, they have already provided deep insights into a wide range of biological processes. Here, we provide an updated description and comparison of proximity labeling methods, as well as their applications and improvements. As each method has its own unique features, the goal of this review is to describe how different proximity labeling methods can be used to answer different biological questions. This article is categorized under: Technologies > Analysis of Proteins.
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Affiliation(s)
- Justin A Bosch
- Department of Genetics, Blavatnik Institute, Harvard Medical School, Boston, Massachusetts, USA
| | - Chiao-Lin Chen
- Department of Genetics, Blavatnik Institute, Harvard Medical School, Boston, Massachusetts, USA
| | - Norbert Perrimon
- Department of Genetics, Blavatnik Institute, Harvard Medical School, Boston, Massachusetts, USA.,Howard Hughes Medical Institute, Boston, Massachusetts, USA
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16
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Hansel CS, Holme MN, Gopal S, Stevens MM. Advances in high-resolution microscopy for the study of intracellular interactions with biomaterials. Biomaterials 2020; 226:119406. [DOI: 10.1016/j.biomaterials.2019.119406] [Citation(s) in RCA: 22] [Impact Index Per Article: 5.5] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/11/2019] [Revised: 07/16/2019] [Accepted: 08/01/2019] [Indexed: 12/15/2022]
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17
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Doh JK, Tobin SJ, Beatty KE. Generation of CoilR Probe Peptides for VIPER-labeling of Cellular Proteins. Bio Protoc 2019; 9:e3412. [PMID: 33654912 PMCID: PMC7853930 DOI: 10.21769/bioprotoc.3412] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.6] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/24/2019] [Revised: 09/24/2019] [Accepted: 09/30/2019] [Indexed: 11/02/2022] Open
Abstract
Versatile Interacting Peptide (VIP) tags are a new class of genetically-encoded tag designed for imaging cellular proteins by fluorescence and electron microscopy. In 2018, we reported the VIPER tag ( Doh et al., 2018 ), which contains two elements: a genetically-encoded peptide tag (i.e., CoilE) and a probe peptide (i.e., CoilR). These two peptides deliver contrast to a protein of interest by forming a specific, high-affinity heterodimer. The probe peptide was designed with a single cysteine residue for site-specific modification via thiol-maleimide chemistry. This feature can be used to attach a variety of biophysical reporters to the peptide, including bright fluorophores for fluorescence microscopy or electron-dense nanoparticles for electron microscopy. In this Bio-Protocol, we describe our methods for expressing and purifying recombinant CoilR. Additionally, we describe protocols for making fluorescent or biotinylated probe peptides for labeling CoilE-tagged cellular proteins. This protocol is complemented by two other Bio-Protocols outlining the use of VIPER ( Doh et al., 2019a and 2019b).
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Affiliation(s)
- Julia K. Doh
- Department of Biomedical Engineering, Oregon Health & Science University, Portland, Oregon 97239, USA
| | - Savannah J. Tobin
- Department of Biomedical Engineering, Oregon Health & Science University, Portland, Oregon 97239, USA
| | - Kimberly E. Beatty
- Department of Biomedical Engineering, Oregon Health & Science University, Portland, Oregon 97239, USA
- OHSU Center for Spatial Systems Biomedicine, Oregon Health & Science University, Portland, Oregon 97239, USA
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18
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Doh JK, Chang YH, Enns CA, Lόpez CS, Beatty KE. Imaging VIPER-labeled Cellular Proteins by Correlative Light and Electron Microscopy. Bio Protoc 2019; 9:e3414. [PMID: 33654913 PMCID: PMC7853974 DOI: 10.21769/bioprotoc.3414] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.4] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/04/2019] [Revised: 09/24/2019] [Accepted: 09/30/2019] [Indexed: 01/02/2023] Open
Abstract
Advances in fluorescence microscopy (FM), electron microscopy (EM), and correlative light and EM (CLEM) offer unprecedented opportunities for studying diverse proteins and nanostructures involved in fundamental cell biology. It is now possible to visualize and quantify the spatial organization of cellular proteins and other macromolecules by FM, EM, and CLEM. However, tagging and tracking cellular proteins across size scales is restricted by the scarcity of methods for attaching appropriate reporter chemistries to target proteins. Namely, there are few genetic tags compatible with EM. To overcome these issues we developed Versatile Interacting Peptide (VIP) tags, genetically-encoded peptide tags that can be used to image proteins by fluorescence and EM. VIPER, a VIP tag, can be used to label cellular proteins with bright, photo-stable fluorophores for FM or electron-dense nanoparticles for EM. In this Bio-Protocol, we provide an instructional guide for implementing VIPER for imaging a cell-surface receptor by CLEM. This protocol is complemented by two other Bio-Protocols outlining the use of VIPER ( Doh et al., 2019a and 2019b).
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Affiliation(s)
- Julia K. Doh
- Department of Biomedical Engineering, Oregon Health & Science University, Portland, Oregon 97239, USA
| | - Young Hwan Chang
- Department of Biomedical Engineering, Oregon Health & Science University, Portland, Oregon 97239, USA
- OHSU Center for Spatial Systems Biomedicine, Oregon Health & Science University, Portland, Oregon 97239, USA
| | - Caroline A. Enns
- Department of Cell, Developmental, and Cancer Biology, Oregon Health & Science University, Portland, Oregon 97239, USA
| | - Claudia S. Lόpez
- Department of Biomedical Engineering, Oregon Health & Science University, Portland, Oregon 97239, USA
- OHSU Center for Spatial Systems Biomedicine, Oregon Health & Science University, Portland, Oregon 97239, USA
- Multiscale Microscopy Core, Oregon Health & Science University, Portland, Oregon 97239, USA
| | - Kimberly E. Beatty
- Department of Biomedical Engineering, Oregon Health & Science University, Portland, Oregon 97239, USA
- OHSU Center for Spatial Systems Biomedicine, Oregon Health & Science University, Portland, Oregon 97239, USA
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19
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Shahmoradian SH, Lewis AJ, Genoud C, Hench J, Moors TE, Navarro PP, Castaño-Díez D, Schweighauser G, Graff-Meyer A, Goldie KN, Sütterlin R, Huisman E, Ingrassia A, Gier YD, Rozemuller AJM, Wang J, Paepe AD, Erny J, Staempfli A, Hoernschemeyer J, Großerüschkamp F, Niedieker D, El-Mashtoly SF, Quadri M, Van IJcken WFJ, Bonifati V, Gerwert K, Bohrmann B, Frank S, Britschgi M, Stahlberg H, Van de Berg WDJ, Lauer ME. Lewy pathology in Parkinson's disease consists of crowded organelles and lipid membranes. Nat Neurosci 2019; 22:1099-1109. [PMID: 31235907 DOI: 10.1038/s41593-019-0423-2] [Citation(s) in RCA: 521] [Impact Index Per Article: 104.2] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/26/2018] [Accepted: 05/09/2019] [Indexed: 12/17/2022]
Abstract
Parkinson's disease, the most common age-related movement disorder, is a progressive neurodegenerative disease with unclear etiology. Key neuropathological hallmarks are Lewy bodies and Lewy neurites: neuronal inclusions immunopositive for the protein α-synuclein. In-depth ultrastructural analysis of Lewy pathology is crucial to understanding pathogenesis of this disease. Using correlative light and electron microscopy and tomography on postmortem human brain tissue from Parkinson's disease brain donors, we identified α-synuclein immunopositive Lewy pathology and show a crowded environment of membranes therein, including vesicular structures and dysmorphic organelles. Filaments interspersed between the membranes and organelles were identifiable in many but not all α-synuclein inclusions. Crowding of organellar components was confirmed by stimulated emission depletion (STED)-based super-resolution microscopy, and high lipid content within α-synuclein immunopositive inclusions was corroborated by confocal imaging, Fourier-transform coherent anti-Stokes Raman scattering infrared imaging and lipidomics. Applying such correlative high-resolution imaging and biophysical approaches, we discovered an aggregated protein-lipid compartmentalization not previously described in the Parkinsons' disease brain.
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Affiliation(s)
- Sarah H Shahmoradian
- Center for Cellular Imaging and NanoAnalytics, Biozentrum, University of Basel, Basel, Switzerland.,Department of Biology and Chemistry, Paul Scherrer Institute, Villigen, Switzerland
| | - Amanda J Lewis
- Center for Cellular Imaging and NanoAnalytics, Biozentrum, University of Basel, Basel, Switzerland
| | - Christel Genoud
- Friedrich Miescher Institute for Biomedical Research, Basel, Switzerland
| | - Jürgen Hench
- Division of Neuropathology, Institute of Pathology, University Hospital Basel, Basel, Switzerland
| | - Tim E Moors
- Amsterdam Neuroscience, VU University Medical Center, Department of Anatomy and Neurosciences, Section Clinical Neuroanatomy, Amsterdam, The Netherlands
| | - Paula P Navarro
- Center for Cellular Imaging and NanoAnalytics, Biozentrum, University of Basel, Basel, Switzerland
| | - Daniel Castaño-Díez
- Center for Cellular Imaging and NanoAnalytics, Biozentrum, University of Basel, Basel, Switzerland
| | - Gabriel Schweighauser
- Division of Neuropathology, Institute of Pathology, University Hospital Basel, Basel, Switzerland
| | | | - Kenneth N Goldie
- Center for Cellular Imaging and NanoAnalytics, Biozentrum, University of Basel, Basel, Switzerland
| | - Rosmarie Sütterlin
- Center for Cellular Imaging and NanoAnalytics, Biozentrum, University of Basel, Basel, Switzerland
| | - Evelien Huisman
- Amsterdam Neuroscience, VU University Medical Center, Department of Anatomy and Neurosciences, Section Clinical Neuroanatomy, Amsterdam, The Netherlands
| | - Angela Ingrassia
- Amsterdam Neuroscience, VU University Medical Center, Department of Anatomy and Neurosciences, Section Clinical Neuroanatomy, Amsterdam, The Netherlands
| | - Yvonne de Gier
- Amsterdam Neuroscience, VU University Medical Center, Department of Anatomy and Neurosciences, Section Clinical Neuroanatomy, Amsterdam, The Netherlands
| | - Annemieke J M Rozemuller
- Amsterdam Neuroscience, VU University Medical Center, Department of Pathology, Amsterdam, The Netherlands
| | - Jing Wang
- Center for Cellular Imaging and NanoAnalytics, Biozentrum, University of Basel, Basel, Switzerland
| | - Anne De Paepe
- Roche Pharma Research and Early Development, Lead Discovery, Roche Innovation Center Basel, Basel, Switzerland
| | - Johannes Erny
- Roche Pharma Research and Early Development, Preclinical CMC, Roche Innovation Center Basel, Basel, Switzerland
| | - Andreas Staempfli
- Roche Pharma Research and Early Development, Preclinical CMC, Roche Innovation Center Basel, Basel, Switzerland
| | - Joerg Hoernschemeyer
- Roche Pharma Research and Early Development, Preclinical CMC, Roche Innovation Center Basel, Basel, Switzerland
| | | | | | | | - Marialuisa Quadri
- Department of Clinical Genetics, Erasmus Medical Center, Rotterdam, The Netherlands
| | | | - Vincenzo Bonifati
- Department of Clinical Genetics, Erasmus Medical Center, Rotterdam, The Netherlands
| | - Klaus Gerwert
- Department of Biophysics, Ruhr University, Bochum, Germany
| | - Bernd Bohrmann
- Roche Pharma Research and Early Development, Neuroscience, Ophthalmology, and Rare Diseases Discovery and Translational Area/Neuroscience Discovery, Roche Innovation Center Basel, Basel, Switzerland
| | - Stephan Frank
- Division of Neuropathology, Institute of Pathology, University Hospital Basel, Basel, Switzerland
| | - Markus Britschgi
- Roche Pharma Research and Early Development, Neuroscience, Ophthalmology, and Rare Diseases Discovery and Translational Area/Neuroscience Discovery, Roche Innovation Center Basel, Basel, Switzerland
| | - Henning Stahlberg
- Center for Cellular Imaging and NanoAnalytics, Biozentrum, University of Basel, Basel, Switzerland.
| | - Wilma D J Van de Berg
- Amsterdam Neuroscience, VU University Medical Center, Department of Anatomy and Neurosciences, Section Clinical Neuroanatomy, Amsterdam, The Netherlands.
| | - Matthias E Lauer
- Roche Pharma Research and Early Development, Lead Discovery, Roche Innovation Center Basel, Basel, Switzerland.
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20
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Weinhardt V, Chen JH, Ekman A, McDermott G, Le Gros MA, Larabell C. Imaging cell morphology and physiology using X-rays. Biochem Soc Trans 2019; 47:489-508. [PMID: 30952801 PMCID: PMC6716605 DOI: 10.1042/bst20180036] [Citation(s) in RCA: 16] [Impact Index Per Article: 3.2] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/17/2018] [Revised: 01/02/2019] [Accepted: 01/09/2019] [Indexed: 02/07/2023]
Abstract
Morphometric measurements, such as quantifying cell shape, characterizing sub-cellular organization, and probing cell-cell interactions, are fundamental in cell biology and clinical medicine. Until quite recently, the main source of morphometric data on cells has been light- and electron-based microscope images. However, many technological advances have propelled X-ray microscopy into becoming another source of high-quality morphometric information. Here, we review the status of X-ray microscopy as a quantitative biological imaging modality. We also describe the combination of X-ray microscopy data with information from other modalities to generate polychromatic views of biological systems. For example, the amalgamation of molecular localization data, from fluorescence microscopy or spectromicroscopy, with structural information from X-ray tomography. This combination of data from the same specimen generates a more complete picture of the system than that can be obtained by a single microscopy method. Such multimodal combinations greatly enhance our understanding of biology by combining physiological and morphological data to create models that more accurately reflect the complexities of life.
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Affiliation(s)
- Venera Weinhardt
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, U.S.A
- Department of Anatomy, University of California San Francisco, San Francisco, California, U.S.A
| | - Jian-Hua Chen
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, U.S.A
| | - Axel Ekman
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, U.S.A
| | - Gerry McDermott
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, U.S.A
| | - Mark A Le Gros
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, U.S.A
- Department of Anatomy, University of California San Francisco, San Francisco, California, U.S.A
| | - Carolyn Larabell
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA, U.S.A.
- Department of Anatomy, University of California San Francisco, San Francisco, California, U.S.A
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21
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VIPER is a genetically encoded peptide tag for fluorescence and electron microscopy. Proc Natl Acad Sci U S A 2018; 115:12961-12966. [PMID: 30518560 DOI: 10.1073/pnas.1808626115] [Citation(s) in RCA: 22] [Impact Index Per Article: 3.7] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/18/2022] Open
Abstract
Many discoveries in cell biology rely on making specific proteins visible within their native cellular environment. There are various genetically encoded tags, such as fluorescent proteins, developed for fluorescence microscopy (FM). However, there are almost no genetically encoded tags that enable cellular proteins to be observed by both FM and electron microscopy (EM). Herein, we describe a technology for labeling proteins with diverse chemical reporters, including bright organic fluorophores for FM and electron-dense nanoparticles for EM. Our technology uses versatile interacting peptide (VIP) tags, a class of genetically encoded tag. We present VIPER, which consists of a coiled-coil heterodimer formed between the genetic tag, CoilE, and a probe-labeled peptide, CoilR. Using confocal FM, we demonstrate that VIPER can be used to highlight subcellular structures or to image receptor-mediated iron uptake. Additionally, we used VIPER to image the iron uptake machinery by correlative light and EM (CLEM). VIPER compared favorably with immunolabeling for imaging proteins by CLEM, and is an enabling technology for protein targets that cannot be immunolabeled. VIPER is a versatile peptide tag that can be used to label and track proteins with diverse chemical reporters observable by both FM and EM instrumentation.
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22
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Abstract
Sensory photoreceptors underpin light-dependent adaptations of organismal physiology, development, and behavior in nature. Adapted for optogenetics, sensory photoreceptors become genetically encoded actuators and reporters to enable the noninvasive, spatiotemporally accurate and reversible control by light of cellular processes. Rooted in a mechanistic understanding of natural photoreceptors, artificial photoreceptors with customized light-gated function have been engineered that greatly expand the scope of optogenetics beyond the original application of light-controlled ion flow. As we survey presently, UV/blue-light-sensitive photoreceptors have particularly allowed optogenetics to transcend its initial neuroscience applications by unlocking numerous additional cellular processes and parameters for optogenetic intervention, including gene expression, DNA recombination, subcellular localization, cytoskeleton dynamics, intracellular protein stability, signal transduction cascades, apoptosis, and enzyme activity. The engineering of novel photoreceptors benefits from powerful and reusable design strategies, most importantly light-dependent protein association and (un)folding reactions. Additionally, modified versions of these same sensory photoreceptors serve as fluorescent proteins and generators of singlet oxygen, thereby further enriching the optogenetic toolkit. The available and upcoming UV/blue-light-sensitive actuators and reporters enable the detailed and quantitative interrogation of cellular signal networks and processes in increasingly more precise and illuminating manners.
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Affiliation(s)
- Aba Losi
- Department of Mathematical, Physical and Computer Sciences , University of Parma , Parco Area delle Scienze 7/A-43124 Parma , Italy
| | - Kevin H Gardner
- Structural Biology Initiative, CUNY Advanced Science Research Center , New York , New York 10031 , United States.,Department of Chemistry and Biochemistry, City College of New York , New York , New York 10031 , United States.,Ph.D. Programs in Biochemistry, Chemistry, and Biology , The Graduate Center of the City University of New York , New York , New York 10016 , United States
| | - Andreas Möglich
- Lehrstuhl für Biochemie , Universität Bayreuth , 95447 Bayreuth , Germany.,Research Center for Bio-Macromolecules , Universität Bayreuth , 95447 Bayreuth , Germany.,Bayreuth Center for Biochemistry & Molecular Biology , Universität Bayreuth , 95447 Bayreuth , Germany
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23
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Clarke NI, Royle SJ. Correlating light microscopy with serial block face scanning electron microscopy to study mitotic spindle architecture. Methods Cell Biol 2018; 145:29-43. [PMID: 29957210 DOI: 10.1016/bs.mcb.2018.03.010] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/16/2022]
Abstract
The mitotic spindle is a complex structure that coordinates the accurate segregation of chromosomes during cell division. To understand how the mitotic spindle operates at the molecular level, high resolution imaging is needed. Serial block face-scanning electron microscopy (SBF-SEM) is a technique that can be used to visualize the ultrastructure of entire cells, including components of the mitotic spindle such as microtubules, kinetochores, centrosomes, and chromosomes. Although transmission electron microscopy (TEM) has higher resolution, the reconstruction of large volumes using TEM and tomography is labor intensive, whereas SBF-SEM takes only days to process, image, and segment samples. SBF-SEM fills the resolution gap between light microscopy (LM) and TEM. When combined with LM, SBF-SEM provides a platform where dynamic cellular events can be selected and imaged at high resolution. Here we outline methods to use correlation and SBF-SEM to study mitotic spindle architecture in 3D with high resolution.
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Affiliation(s)
- Nicholas I Clarke
- Centre for Mechanochemical Cell Biology, Warwick Medical School, Coventry, United Kingdom
| | - Stephen J Royle
- Centre for Mechanochemical Cell Biology, Warwick Medical School, Coventry, United Kingdom.
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24
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Maleckar MM, Edwards AG, Louch WE, Lines GT. Studying dyadic structure-function relationships: a review of current modeling approaches and new insights into Ca 2+ (mis)handling. CLINICAL MEDICINE INSIGHTS-CARDIOLOGY 2017; 11:1179546817698602. [PMID: 28469494 PMCID: PMC5392018 DOI: 10.1177/1179546817698602] [Citation(s) in RCA: 8] [Impact Index Per Article: 1.1] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Submit a Manuscript] [Subscribe] [Scholar Register] [Received: 06/30/2016] [Accepted: 12/19/2016] [Indexed: 11/25/2022]
Abstract
Excitation–contraction coupling in cardiac myocytes requires calcium influx through L-type calcium channels in the sarcolemma, which gates calcium release through sarcoplasmic reticulum ryanodine receptors in a process known as calcium-induced calcium release, producing a myoplasmic calcium transient and enabling cardiomyocyte contraction. The spatio-temporal dynamics of calcium release, buffering, and reuptake into the sarcoplasmic reticulum play a central role in excitation–contraction coupling in both normal and diseased cardiac myocytes. However, further quantitative understanding of these cells’ calcium machinery and the study of mechanisms that underlie both normal cardiac function and calcium-dependent etiologies in heart disease requires accurate knowledge of cardiac ultrastructure, protein distribution and subcellular function. As current imaging techniques are limited in spatial resolution, limiting insight into changes in calcium handling, computational models of excitation–contraction coupling have been increasingly employed to probe these structure–function relationships. This review will focus on the development of structural models of cardiac calcium dynamics at the subcellular level, orienting the reader broadly towards the development of models of subcellular calcium handling in cardiomyocytes. Specific focus will be given to progress in recent years in terms of multi-scale modeling employing resolved spatial models of subcellular calcium machinery. A review of the state-of-the-art will be followed by a review of emergent insights into calcium-dependent etiologies in heart disease and, finally, we will offer a perspective on future directions for related computational modeling and simulation efforts.
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Affiliation(s)
- Mary M Maleckar
- Simula Research Laboratory, Center for Cardiological Innovation and Center for Biomedical Computing, Lysaker, Norway
| | - Andrew G Edwards
- Simula Research Laboratory, Center for Cardiological Innovation and Center for Biomedical Computing, Lysaker, Norway.,University of Oslo, Oslo, Norway
| | - William E Louch
- Institute for Experimental Medical Research (IEMR), Oslo University Hospital and the University of Oslo, Oslo, Norway
| | - Glenn T Lines
- Simula Research Laboratory, Center for Cardiological Innovation and Center for Biomedical Computing, Lysaker, Norway
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25
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Chen CL, Perrimon N. Proximity-dependent labeling methods for proteomic profiling in living cells. WILEY INTERDISCIPLINARY REVIEWS-DEVELOPMENTAL BIOLOGY 2017; 6. [PMID: 28387482 DOI: 10.1002/wdev.272] [Citation(s) in RCA: 38] [Impact Index Per Article: 5.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Received: 01/03/2017] [Revised: 02/03/2017] [Accepted: 02/10/2017] [Indexed: 02/05/2023]
Abstract
Characterizing the proteome composition of organelles and subcellular regions of living cells can facilitate the understanding of cellular organization as well as protein interactome networks. Proximity labeling-based methods coupled with mass spectrometry (MS) offer a high-throughput approach for systematic analysis of spatially restricted proteomes. Proximity labeling utilizes enzymes that generate reactive radicals to covalently tag neighboring proteins with biotin. The biotinylated endogenous proteins can then be isolated for further analysis by MS. To analyze protein-protein interactions or identify components that localize to discrete subcellular compartments, spatial expression is achieved by fusing the enzyme to specific proteins or signal peptides that target to particular subcellular regions. Although these technologies have only been introduced recently, they have already provided deep insights into a wide range of biological processes. Here, we describe and compare current methods of proximity labeling as well as their applications. As each method has its own unique features, the goal of this review is to describe how different proximity labeling methods can be used to answer different biological questions. WIREs Dev Biol 2017, 6:e272. doi: 10.1002/wdev.272 For further resources related to this article, please visit the WIREs website.
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Affiliation(s)
- Chiao-Lin Chen
- Department of Genetics, Harvard Medical School, Boston, MA, USA
| | - Norbert Perrimon
- Department of Genetics, Harvard Medical School, Boston, MA, USA.,Howard Hughes Medical Institute, Boston, MA, USA
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26
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Eid L, Parent M. Preparation of Non-human Primate Brain Tissue for Pre-embedding Immunohistochemistry and Electron Microscopy. J Vis Exp 2017. [PMID: 28448038 DOI: 10.3791/55397] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/19/2023] Open
Abstract
Despite all the technological advances at the light microscopy level, electron microscopy remains the only tool in neuroscience to examine and characterize ultrastructural and morphological details of neurons, such as synaptic contacts. Good preservation of brain tissue for electron microscopy can be obtained by rigorous cryo-fixation methods, but these techniques are rather costly and limit the use of immunolabeling, which is crucial to understand the connectivity of identified neuronal systems. Freeze-substitution methods have been developed to allow the combination of cryo-fixation with immunolabeling. However, the reproducibility of these methodological approaches usually relies on costly freezing devices. Moreover, achieving reliable results with this technique is very time-consuming and skill-challenging. Hence, the traditional chemically fixed brain, particularly with acrolein fixative, remains a time-efficient and low-cost method to combine electron microscopy with immunohistochemistry. Here, we provide a reliable experimental protocol using chemical acrolein fixation that leads to the preservation of primate brain tissue and is compatible with pre-embedding immunohistochemistry and transmission electron microscopic examination.
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Affiliation(s)
- Lara Eid
- Centre de recherche de l'Institut universitaire en santé mentale de Québec, Department of Psychiatry and Neuroscience, Université Laval; Centre de recherche du CHU Sainte-Justine;
| | - Martin Parent
- Centre de recherche de l'Institut universitaire en santé mentale de Québec, Department of Psychiatry and Neuroscience, Université Laval
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27
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McIntosh JR, Morphew MK, Giddings TH. Electron Microscopy of Fission Yeast. Cold Spring Harb Protoc 2017; 2017:2017/1/pdb.top079822. [PMID: 28049809 DOI: 10.1101/pdb.top079822] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 06/06/2023]
Abstract
Electron microscopy (EM) can provide images of cells with a spatial resolution that significantly surpasses that available from light microscopy (LM), even with modern methods that give LM "super resolution." However, EM resolution comes with costs in time spent with sample preparation, expense of instrumentation, and concerns regarding sample preparation artifacts. It is therefore important to know the limitations of EM as well as its strengths. Here we describe the most reliable methods for the preservation of fission yeast cells currently available. We describe the properties of images obtained by transmission EM (TEM) and contrast them with images from scanning EM (SEM). We also show how one can make three-dimensional TEM images and discuss several approaches to address the problem of localizing specific proteins within cells. We give references to work by others who have pursued similar goals with different methods, and we discuss briefly the complex subject of image interpretation.
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Affiliation(s)
- J Richard McIntosh
- Department of Molecular, Cellular, and Developmental Biology, University of Colorado, Boulder, Colorado 80309-0347
- Laboratory for 3D Electron Microscopy, University of Colorado, Boulder, Colorado 80309-0347
| | - Mary K Morphew
- Laboratory for 3D Electron Microscopy, University of Colorado, Boulder, Colorado 80309-0347
| | - Thomas H Giddings
- Department of Molecular, Cellular, and Developmental Biology, University of Colorado, Boulder, Colorado 80309-0347
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28
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Hauser M, Wojcik M, Kim D, Mahmoudi M, Li W, Xu K. Correlative Super-Resolution Microscopy: New Dimensions and New Opportunities. Chem Rev 2017; 117:7428-7456. [PMID: 28045508 DOI: 10.1021/acs.chemrev.6b00604] [Citation(s) in RCA: 101] [Impact Index Per Article: 14.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 02/08/2023]
Abstract
Correlative microscopy, the integration of two or more microscopy techniques performed on the same sample, produces results that emphasize the strengths of each technique while offsetting their individual weaknesses. Light microscopy has historically been a central method in correlative microscopy due to its widespread availability, compatibility with hydrated and live biological samples, and excellent molecular specificity through fluorescence labeling. However, conventional light microscopy can only achieve a resolution of ∼300 nm, undercutting its advantages in correlations with higher-resolution methods. The rise of super-resolution microscopy (SRM) over the past decade has drastically improved the resolution of light microscopy to ∼10 nm, thus creating exciting new opportunities and challenges for correlative microscopy. Here we review how these challenges are addressed to effectively correlate SRM with other microscopy techniques, including light microscopy, electron microscopy, cryomicroscopy, atomic force microscopy, and various forms of spectroscopy. Though we emphasize biological studies, we also discuss the application of correlative SRM to materials characterization and single-molecule reactions. Finally, we point out current limitations and discuss possible future improvements and advances. We thus demonstrate how a correlative approach adds new dimensions of information and provides new opportunities in the fast-growing field of SRM.
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Affiliation(s)
- Meghan Hauser
- Department of Chemistry, University of California , Berkeley, California 94720, United States
| | - Michal Wojcik
- Department of Chemistry, University of California , Berkeley, California 94720, United States
| | - Doory Kim
- Department of Chemistry, University of California , Berkeley, California 94720, United States
| | - Morteza Mahmoudi
- Center for Nanomedicine and Department of Anesthesiology, Brigham and Women's Hospital, Harvard Medical School , Boston, Massachusetts 02115, United States
| | - Wan Li
- Department of Chemistry, University of California , Berkeley, California 94720, United States
| | - Ke Xu
- Department of Chemistry, University of California , Berkeley, California 94720, United States.,Division of Molecular Biophysics and Integrated Bioimaging, Lawrence Berkeley National Laboratory , Berkeley, California 94720, United States
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29
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Ekman AA, Chen JH, Guo J, McDermott G, Le Gros MA, Larabell CA. Mesoscale imaging with cryo-light and X-rays: Larger than molecular machines, smaller than a cell. Biol Cell 2017; 109:24-38. [PMID: 27690365 PMCID: PMC5261833 DOI: 10.1111/boc.201600044] [Citation(s) in RCA: 25] [Impact Index Per Article: 3.6] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/28/2016] [Revised: 09/27/2016] [Accepted: 09/28/2016] [Indexed: 12/11/2022]
Abstract
In the context of cell biology, the term mesoscale describes length scales ranging from that of an individual cell, down to the size of the molecular machines. In this spatial regime, small building blocks self-organise to form large, functional structures. A comprehensive set of rules governing mesoscale self-organisation has not been established, making the prediction of many cell behaviours difficult, if not impossible. Our knowledge of mesoscale biology comes from experimental data, in particular, imaging. Here, we explore the application of soft X-ray tomography (SXT) to imaging the mesoscale, and describe the structural insights this technology can generate. We also discuss how SXT imaging is complemented by the addition of correlative fluorescence data measured from the same cell. This combination of two discrete imaging modalities produces a 3D view of the cell that blends high-resolution structural information with precise molecular localisation data.
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Affiliation(s)
- Axel A. Ekman
- Department of Anatomy, School of Medicine, University of California San Francisco, San Francisco, CA 94158, USA
| | - Jian-Hua Chen
- Department of Anatomy, School of Medicine, University of California San Francisco, San Francisco, CA 94158, USA
| | - Jessica Guo
- Department of Anatomy, School of Medicine, University of California San Francisco, San Francisco, CA 94158, USA
| | - Gerry McDermott
- Department of Anatomy, School of Medicine, University of California San Francisco, San Francisco, CA 94158, USA
| | - Mark A. Le Gros
- Department of Anatomy, School of Medicine, University of California San Francisco, San Francisco, CA 94158, USA
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, USA
| | - Carolyn A. Larabell
- Department of Anatomy, School of Medicine, University of California San Francisco, San Francisco, CA 94158, USA
- Molecular Biophysics and Integrated Bioimaging Division, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, USA
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30
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Martynenko IV, Litvin AP, Purcell-Milton F, Baranov AV, Fedorov AV, Gun'ko YK. Application of semiconductor quantum dots in bioimaging and biosensing. J Mater Chem B 2017; 5:6701-6727. [DOI: 10.1039/c7tb01425b] [Citation(s) in RCA: 200] [Impact Index Per Article: 28.6] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/15/2022]
Abstract
In this review we present new concepts and recent progress in the application of semiconductor quantum dots (QD) as labels in two important areas of biology, bioimaging and biosensing.
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Affiliation(s)
- I. V. Martynenko
- BAM Federal Institute for Materials Research and Testing
- 12489 Berlin
- Germany
- ITMO University
- St. Petersburg
| | | | | | | | | | - Y. K. Gun'ko
- ITMO University
- St. Petersburg
- Russia
- School of Chemistry and CRANN
- Trinity College Dublin
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31
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Spatially resolved proteomic mapping in living cells with the engineered peroxidase APEX2. Nat Protoc 2016; 11:456-75. [PMID: 26866790 DOI: 10.1038/nprot.2016.018] [Citation(s) in RCA: 347] [Impact Index Per Article: 43.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/25/2022]
Abstract
This protocol describes a method to obtain spatially resolved proteomic maps of specific compartments within living mammalian cells. An engineered peroxidase, APEX2, is genetically targeted to a cellular region of interest. Upon the addition of hydrogen peroxide for 1 min to cells preloaded with a biotin-phenol substrate, APEX2 generates biotin-phenoxyl radicals that covalently tag proximal endogenous proteins. Cells are then lysed, and biotinylated proteins are enriched with streptavidin beads and identified by mass spectrometry. We describe the generation of an appropriate APEX2 fusion construct, proteomic sample preparation, and mass spectrometric data acquisition and analysis. A two-state stable isotope labeling by amino acids in cell culture (SILAC) protocol is used for proteomic mapping of membrane-enclosed cellular compartments from which APEX2-generated biotin-phenoxyl radicals cannot escape. For mapping of open cellular regions, we instead use a 'ratiometric' three-state SILAC protocol for high spatial specificity. Isotopic labeling of proteins takes 5-7 cell doublings. Generation of the biotinylated proteomic sample takes 1 d, acquiring the mass spectrometric data takes 2-5 d and analysis of the data to obtain the final proteomic list takes 1 week.
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32
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Vectors for Genetically-Encoded Tags for Electron Microscopy Contrast in Drosophila. Biol Proced Online 2016; 18:5. [PMID: 26839516 PMCID: PMC4736618 DOI: 10.1186/s12575-016-0034-1] [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: 07/07/2015] [Accepted: 01/25/2016] [Indexed: 11/10/2022] Open
Abstract
BACKGROUND One of the most notable recent advances in electron microscopy (EM) was the development of genetically-encoded EM tags, including the fluorescent flavoprotein Mini-SOG (Mini-Singlet Oxygen Generator). Mini-SOG generates good EM contrast, thus providing a viable alternative to technically-demanding methods such as immuno-electron microcopy (immuno-EM). Based on the Mini-SOG technology, in this paper, we describe the construction, validation and optimization of a series of vectors which allow expression of Mini-SOG in the Drosophila melanogaster genetic model system. FINDINGS We constructed a Mini-SOG tag that has been codon-optimized for expression in Drosophila (DMS tag) using PCR-mediated gene assembly. The photo-oxidation reaction triggered by DMS was then tested using these vectors in Drosophila cell lines. DMS tag did not affect the subcellular localization of the proteins we tested. More importantly, we demonstrated the utility of the DMS tag for EM in Drosophila by showing that it can produce robust photo-oxidation reactions in the presence of blue light and the substrate DAB; the resultant electron micrographs contain electron-dense regions corresponding to the protein of interest. The vectors we generated allow protein tagging at both termini, for constitutive and inducible protein expression, as well as the generation of transgenic lines by P-element transformation. CONCLUSIONS We demonstrated the feasibility of Mini-SOG tagging in Drosophila. The constructed vectors will no doubt be a useful molecular tool for genetic tagging to facilitate high-resolution localization of proteins in Drosophila by electron microscopy.
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33
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Self-labelling enzymes as universal tags for fluorescence microscopy, super-resolution microscopy and electron microscopy. Sci Rep 2015; 5:17740. [PMID: 26643905 PMCID: PMC4672345 DOI: 10.1038/srep17740] [Citation(s) in RCA: 66] [Impact Index Per Article: 7.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/16/2015] [Accepted: 11/04/2015] [Indexed: 11/17/2022] Open
Abstract
Research in cell biology demands advanced microscopy techniques such as confocal fluorescence microscopy (FM), super-resolution microscopy (SRM) and transmission electron microscopy (TEM). Correlative light and electron microscopy (CLEM) is an approach to combine data on the dynamics of proteins or protein complexes in living cells with the ultrastructural details in the low nanometre scale. To correlate both data sets, markers functional in FM, SRM and TEM are required. Genetically encoded markers such as fluorescent proteins or self-labelling enzyme tags allow observations in living cells. Various genetically encoded tags are available for FM and SRM, but only few tags are suitable for CLEM. Here, we describe the red fluorescent dye tetramethylrhodamine (TMR) as a multimodal marker for CLEM. TMR is used as fluorochrome coupled to ligands of genetically encoded self-labelling enzyme tags HaloTag, SNAP-tag and CLIP-tag in FM and SRM. We demonstrate that TMR can additionally photooxidize diaminobenzidine (DAB) to an osmiophilic polymer visible on TEM sections, thus being a marker suitable for FM, SRM and TEM. We evaluated various organelle markers with enzymatic tags in mammalian cells labelled with TMR-coupled ligands and demonstrate the use as efficient and versatile DAB photooxidizer for CLEM approaches.
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34
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Morphew MK, O'Toole ET, Page CL, Pagratis M, Meehl J, Giddings T, Gardner JM, Ackerson C, Jaspersen SL, Winey M, Hoenger A, McIntosh JR. Metallothionein as a clonable tag for protein localization by electron microscopy of cells. J Microsc 2015; 260:20-9. [PMID: 25974385 PMCID: PMC4573841 DOI: 10.1111/jmi.12262] [Citation(s) in RCA: 24] [Impact Index Per Article: 2.7] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/10/2015] [Accepted: 04/07/2015] [Indexed: 11/28/2022]
Abstract
A benign, clonable tag for the localization of proteins by electron microscopy of cells would be valuable, especially if it provided labelling with high signal-to-noise ratio and good spatial resolution. Here we explore the use of metallothionein as such a localization marker. We have achieved good success with desmin labelled in vitro and with a component of the yeast spindle pole body labelled in cells. Heavy metals added after fixation and embedding or during the process of freeze-substitution fixation provide readily visible signals with no concern that the heavy atoms are affecting the behaviour of the protein in its physiological environment. However, our methods did not work with protein components of the nuclear pore complex, suggesting that this approach is not yet universally applicable. We provide a full description of our optimal labelling conditions and other conditions tried, hoping that our work will allow others to label their own proteins of interest and/or improve on the methods we have defined.
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Affiliation(s)
- M K Morphew
- Department of Molecular, Cellular and Developmental, Biology University of Colorado, Boulder, Colorado, 80309-0347, U.S.A
| | - E T O'Toole
- Department of Molecular, Cellular and Developmental, Biology University of Colorado, Boulder, Colorado, 80309-0347, U.S.A
| | - C L Page
- Department of Molecular, Cellular and Developmental, Biology University of Colorado, Boulder, Colorado, 80309-0347, U.S.A
| | - M Pagratis
- Department of Molecular, Cellular and Developmental, Biology University of Colorado, Boulder, Colorado, 80309-0347, U.S.A
| | - J Meehl
- Department of Molecular, Cellular and Developmental, Biology University of Colorado, Boulder, Colorado, 80309-0347, U.S.A
| | - T Giddings
- Department of Molecular, Cellular and Developmental, Biology University of Colorado, Boulder, Colorado, 80309-0347, U.S.A
| | - J M Gardner
- The Stowers Institute for Medical Research, Kansas City, Missouri, 64110 and Department of Molecular and Integrative Physiology, University of Kansas Medical Center, Kansas City, Kansas, 66160, U.S.A
| | - C Ackerson
- Department of Chemistry, Colorado State University, Fort Collins, Colorado, U.S.A
| | - S L Jaspersen
- The Stowers Institute for Medical Research, Kansas City, Missouri, 64110 and Department of Molecular and Integrative Physiology, University of Kansas Medical Center, Kansas City, Kansas, 66160, U.S.A
| | - M Winey
- Department of Molecular, Cellular and Developmental, Biology University of Colorado, Boulder, Colorado, 80309-0347, U.S.A
| | - A Hoenger
- Department of Molecular, Cellular and Developmental, Biology University of Colorado, Boulder, Colorado, 80309-0347, U.S.A
| | - J R McIntosh
- Department of Molecular, Cellular and Developmental, Biology University of Colorado, Boulder, Colorado, 80309-0347, U.S.A
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35
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Rajagopal V, Bass G, Walker CG, Crossman DJ, Petzer A, Hickey A, Siekmann I, Hoshijima M, Ellisman MH, Crampin EJ, Soeller C. Examination of the Effects of Heterogeneous Organization of RyR Clusters, Myofibrils and Mitochondria on Ca2+ Release Patterns in Cardiomyocytes. PLoS Comput Biol 2015; 11:e1004417. [PMID: 26335304 PMCID: PMC4559435 DOI: 10.1371/journal.pcbi.1004417] [Citation(s) in RCA: 30] [Impact Index Per Article: 3.3] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/07/2014] [Accepted: 06/26/2015] [Indexed: 11/18/2022] Open
Abstract
Spatio-temporal dynamics of intracellular calcium, [Ca2+]i, regulate the contractile function of cardiac muscle cells. Measuring [Ca2+]i flux is central to the study of mechanisms that underlie both normal cardiac function and calcium-dependent etiologies in heart disease. However, current imaging techniques are limited in the spatial resolution to which changes in [Ca2+]i can be detected. Using spatial point process statistics techniques we developed a novel method to simulate the spatial distribution of RyR clusters, which act as the major mediators of contractile Ca2+ release, upon a physiologically-realistic cellular landscape composed of tightly-packed mitochondria and myofibrils. We applied this method to computationally combine confocal-scale (~ 200 nm) data of RyR clusters with 3D electron microscopy data (~ 30 nm) of myofibrils and mitochondria, both collected from adult rat left ventricular myocytes. Using this hybrid-scale spatial model, we simulated reaction-diffusion of [Ca2+]i during the rising phase of the transient (first 30 ms after initiation). At 30 ms, the average peak of the simulated [Ca2+]i transient and of the simulated fluorescence intensity signal, F/F0, reached values similar to that found in the literature ([Ca2+]i ≈1 μM; F/F0≈5.5). However, our model predicted the variation in [Ca2+]i to be between 0.3 and 12.7 μM (~3 to 100 fold from resting value of 0.1 μM) and the corresponding F/F0 signal ranging from 3 to 9.5. We demonstrate in this study that: (i) heterogeneities in the [Ca2+]i transient are due not only to heterogeneous distribution and clustering of mitochondria; (ii) but also to heterogeneous local densities of RyR clusters. Further, we show that: (iii) these structure-induced heterogeneities in [Ca2+]i can appear in line scan data. Finally, using our unique method for generating RyR cluster distributions, we demonstrate the robustness in the [Ca2+]i transient to differences in RyR cluster distributions measured between rat and human cardiomyocytes. Calcium (Ca2+) acts as a signal for many functions in the heart cell, from its primary role in triggering contractions during the heartbeat to acting as a signal for cell growth. Cellular function is tightly coupled to its ultra-structural organization. Spatially-realistic and biophysics-based computational models can provide quantitative insights into structure-function relationships in Ca2+ signaling. We developed a novel computational model of a rat ventricular myocyte that integrates structural information from confocal and electron microscopy datasets that were independently acquired and includes: myofibrils (protein complexes that contract during the heartbeat), mitochondria (organelles that provide energy for contraction), and ryanodine receptors (RyR, ion channels that release the Ca2+ required to trigger myofibril contraction from intracellular stores). Using this model, we examined [Ca2+]i dynamics throughout the cell cross-section at a much higher resolution than previously possible. We estimated the size of structural maladaptation that would cause disease-related alterations in [Ca2+]i dynamics. Using our methods for data integration, we also tested whether reducing the density of RyRs in human cardiomyocytes (~40% relative to rat) would have a significant effect on [Ca2+]i. We found that Ca2+ release patterns between the two species are similar, suggesting Ca2+ dynamics are robust to variations in cell ultrastructure.
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Affiliation(s)
- Vijay Rajagopal
- Department of Mechanical Engineering, University of Melbourne, Melbourne, Australia
- Auckland Bioengineering Institute, University of Auckland, Auckland, New Zealand
- Systems Biology Laboratory, Melbourne School of Engineering, University of Melbourne, Melbourne, Australia
- * E-mail:
| | - Gregory Bass
- Systems Biology Laboratory, Melbourne School of Engineering, University of Melbourne, Melbourne, Australia
| | - Cameron G. Walker
- Department of Engineering Science, University of Auckland, Auckland, New Zealand
| | - David J. Crossman
- Department of Physiology, University of Auckland, Auckland, New Zealand
| | - Amorita Petzer
- School of Biological Sciences, University of Auckland, Auckland. New Zealand
| | - Anthony Hickey
- School of Biological Sciences, University of Auckland, Auckland. New Zealand
| | - Ivo Siekmann
- Department of Mechanical Engineering, University of Melbourne, Melbourne, Australia
| | - Masahiko Hoshijima
- Department of Medicine, University of California San Diego, San Diego, United States of America
- National Center for Microscopy and Imaging Research, University of California San Diego, San Diego, United States of America
| | - Mark H. Ellisman
- National Center for Microscopy and Imaging Research, University of California San Diego, San Diego, United States of America
| | - Edmund J. Crampin
- Systems Biology Laboratory, Melbourne School of Engineering, University of Melbourne, Melbourne, Australia
- School of Mathematics and Statistics, Faculty of Science, University of Melbourne, Melbourne, Australia
- School of Medicine, Faculty of Medicine, Dentistry and Health Sciences, University of Melbourne, Melbourne, Australia
- ARC Centre of Excellence in Convergent Bio-Nano Science and Technology, University of Melbourne, Melbourne, Australia
| | - Christian Soeller
- Department of Physiology, University of Auckland, Auckland, New Zealand
- Biomedical Physics, University of Exeter, Exeter, United Kingdom
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Do M, Isaacson SA, McDermott G, Le Gros MA, Larabell CA. Imaging and characterizing cells using tomography. Arch Biochem Biophys 2015; 581:111-21. [PMID: 25602704 PMCID: PMC4506273 DOI: 10.1016/j.abb.2015.01.011] [Citation(s) in RCA: 24] [Impact Index Per Article: 2.7] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/01/2014] [Revised: 12/29/2014] [Accepted: 01/11/2015] [Indexed: 12/11/2022]
Abstract
We can learn much about cell function by imaging and quantifying sub-cellular structures, especially if this is done non-destructively without altering said structures. Soft X-ray tomography (SXT) is a high-resolution imaging technique for visualizing cells and their interior structure in 3D. A tomogram of the cell, reconstructed from a series of 2D projection images, can be easily segmented and analyzed. SXT has a very high specimen throughput compared to other high-resolution structure imaging modalities; for example, tomographic data for reconstructing an entire eukaryotic cell is acquired in a matter of minutes. SXT visualizes cells without the need for chemical fixation, dehydration, or staining of the specimen. As a result, the SXT reconstructions are close representations of cells in their native state. SXT is applicable to most cell types. The deep penetration of soft X-rays allows cells, even mammalian cells, to be imaged without being sectioned. Image contrast in SXT is generated by the differential attenuation soft X-ray illumination as it passes through the specimen. Accordingly, each voxel in the tomographic reconstruction has a measured linear absorption coefficient (LAC) value. LAC values are quantitative and give rise to each sub-cellular component having a characteristic LAC profile, allowing organelles to be identified and segmented from the milieu of other cell contents. In this chapter, we describe the fundamentals of SXT imaging and how this technique can answer real world questions in the study of the nucleus. We also describe the development of correlative methods for the localization of specific molecules in a SXT reconstruction. The combination of fluorescence and SXT data acquired from the same specimen produces composite 3D images, rich with detailed information on the inner workings of cells.
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Affiliation(s)
- Myan Do
- Department of Anatomy, University of California San Francisco, San Francisco, CA 94143, United States; National Center for X-ray Tomography, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, United States; Physical Biosciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, United States
| | - Samuel A Isaacson
- Department of Mathematics and Statistics, Boston University, Boston, MA 02215, United States
| | - Gerry McDermott
- Department of Anatomy, University of California San Francisco, San Francisco, CA 94143, United States; National Center for X-ray Tomography, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, United States; Physical Biosciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, United States
| | - Mark A Le Gros
- Department of Anatomy, University of California San Francisco, San Francisco, CA 94143, United States; National Center for X-ray Tomography, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, United States; Physical Biosciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, United States
| | - Carolyn A Larabell
- Department of Anatomy, University of California San Francisco, San Francisco, CA 94143, United States; National Center for X-ray Tomography, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, United States; Physical Biosciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, United States.
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37
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Quantitatively imaging chromosomes by correlated cryo-fluorescence and soft x-ray tomographies. Biophys J 2015; 107:1988-1996. [PMID: 25418180 DOI: 10.1016/j.bpj.2014.09.011] [Citation(s) in RCA: 41] [Impact Index Per Article: 4.6] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/01/2014] [Revised: 08/28/2014] [Accepted: 09/09/2014] [Indexed: 11/24/2022] Open
Abstract
Soft x-ray tomography (SXT) is increasingly being recognized as a valuable method for visualizing and quantifying the ultrastructure of cryopreserved cells. Here, we describe the combination of SXT with cryogenic confocal fluorescence tomography (CFT). This correlative approach allows the incorporation of molecular localization data, with isotropic precision, into high-resolution three-dimensional (3-D) SXT reconstructions of the cell. CFT data are acquired first using a cryogenically adapted confocal light microscope in which the specimen is coupled to a high numerical aperture objective lens by an immersion fluid. The specimen is then cryo-transferred to a soft x-ray microscope (SXM) for SXT data acquisition. Fiducial markers visible in both types of data act as common landmarks, enabling accurate coalignment of the two complementary tomographic reconstructions. We used this method to identify the inactive X chromosome (Xi) in female v-abl transformed thymic lymphoma cells by localizing enhanced green fluorescent protein-labeled macroH2A with CFT. The molecular localization data were used to guide segmentation of Xi in the SXT reconstructions, allowing characterization of the Xi topological arrangement in near-native state cells. Xi was seen to adopt a number of different topologies with no particular arrangement being dominant.
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38
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The potential of bioorthogonal chemistry for correlative light and electron microscopy: a call to arms. J Chem Biol 2015. [DOI: 10.1007/s12154-015-0134-4] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.2] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 10/23/2022] Open
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39
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Advances in molecular probe-based labeling tools and their application to multiscale multimodal correlated microscopies. J Chem Biol 2015. [DOI: 10.1007/s12154-015-0132-6] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.1] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 02/05/2023] Open
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40
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Abstract
Correlative fluorescence and electron microscopy (CFEM) is a multimodal technique that combines dynamic and localization information from fluorescence methods with ultrastructural data from electron microscopy, to give new information about how cellular components change relative to the spatiotemporal dynamics within their environment. In this review, we will discuss some of the basic techniques and tools of the trade for utilizing this attractive research method, which is becoming a very powerful tool for biology labs. The information obtained from correlative methods has proven to be invaluable in creating consensus between the two types of microscopy, extending the capability of each, and cutting the time and expense associated with using each method separately for comparative analysis. The realization of the advantages of these methods in cell biology has led to rapid improvement in the protocols and has ushered in a new generation of instruments to reach the next level of correlation--integration.
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Affiliation(s)
- Randall T Schirra
- Department of Structural Biology, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania
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41
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Reorganization of the endosomal system in Salmonella-infected cells: the ultrastructure of Salmonella-induced tubular compartments. PLoS Pathog 2014; 10:e1004374. [PMID: 25254663 PMCID: PMC4177991 DOI: 10.1371/journal.ppat.1004374] [Citation(s) in RCA: 55] [Impact Index Per Article: 5.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/10/2014] [Accepted: 08/03/2014] [Indexed: 12/03/2022] Open
Abstract
During the intracellular life of Salmonella enterica, a unique membrane-bound compartment termed Salmonella-containing vacuole, or SCV, is formed. By means of translocated effector proteins, intracellular Salmonella also induce the formation of extensive, highly dynamic membrane tubules termed Salmonella-induced filaments or SIF. Here we report the first detailed ultrastructural analyses of the SCV and SIF by electron microscopy (EM), EM tomography and live cell correlative light and electron microscopy (CLEM). We found that a subset of SIF is composed of double membranes that enclose portions of host cell cytosol and cytoskeletal filaments within its inner lumen. Despite some morphological similarities, we found that the formation of SIF double membranes is independent from autophagy and requires the function of the effector proteins SseF and SseG. The lumen of SIF network is accessible to various types of endocytosed material and our CLEM analysis of double membrane SIF demonstrated that fluid phase markers accumulate only between the inner and outer membrane of these structures, a space continual with endosomal lumen. Our work reveals how manipulation of the endosomal membrane system by an intracellular pathogen results in a unique tubular membrane compartmentalization of the host cell, generating a shielded niche permissive for intracellular proliferation of Salmonella. Salmonella enterica is an invasive, facultative intracellular bacterial pathogen. Within mammalian host cells, Salmonella inhabits a specialized membrane-bound compartment, the Salmonella-containing vacuole (SCV), redirects host cell vesicular transport and massively remodels the endosomal system. These activities depend on the function of a type III secretion system and its translocated effector proteins. Intracellular Salmonella induces several types of tubular compartments termed Salmonella-induced tubules (SIT), but the biogenesis and biological function of SIT is only partially understood. Our work combines live cell imaging with correlative light and electron microscopy to provide ultrastructural insight into SIT. We report that SIT emerge as single membrane tubules that convert into double membrane tubules entrapping cytosol and cytoskeletal filaments. Labeling of the endosomal compartment and cytochemistry demonstrate that the space between inner and outer SIT membrane is composed of internalized material and connected to Salmonella within the SCV. The effector proteins SseF and SseG translocated by intracellular Salmonella are essential for the conversion of single to double membrane SIT. These findings challenge current models for the intracellular lifestyle of Salmonella and the composition of its intracellular habitat.
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42
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Cinquin BP, Do M, McDermott G, Walters AD, Myllys M, Smith EA, Cohen-Fix O, Le Gros MA, Larabell CA. Putting molecules in their place. J Cell Biochem 2014; 115:209-16. [PMID: 23966233 DOI: 10.1002/jcb.24658] [Citation(s) in RCA: 32] [Impact Index Per Article: 3.2] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/10/2013] [Accepted: 08/14/2013] [Indexed: 12/18/2022]
Abstract
Each class of microscope is limited to imaging specific aspects of cell structure and/or molecular organization. However, imaging the specimen by complementary microscopes and correlating the data can overcome this limitation. Whilst not a new approach, the field of correlative imaging is currently benefitting from the emergence of new microscope techniques. Here we describe the correlation of cryogenic fluorescence tomography (CFT) with soft X-ray tomography (SXT). This amalgamation of techniques integrates 3D molecular localization data (CFT) with a high-resolution, 3D cell reconstruction of the cell (SXT). Cells are imaged in both modalities in a near-native, cryopreserved state. Here we describe the current state of the art in correlative CFT-SXT, and discuss the future outlook for this method.
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Affiliation(s)
- Bertrand P Cinquin
- Department of Anatomy, University of California San Francisco, San Francisco, California
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43
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Bos E, Hussaarts L, van Weering JRT, Ellisman MH, de Wit H, Koster AJ. Vitrification of Tokuyasu-style immuno-labelled sections for correlative cryo light microscopy and cryo electron tomography. J Struct Biol 2014; 186:273-82. [PMID: 24704216 DOI: 10.1016/j.jsb.2014.03.021] [Citation(s) in RCA: 24] [Impact Index Per Article: 2.4] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/11/2013] [Revised: 03/24/2014] [Accepted: 03/25/2014] [Indexed: 10/25/2022]
Abstract
We present an approach for the preparation of immuno-labelled ultrathin sections from cells or tissue that are compatible with both fluorescence and transmission electron microscopy. Our approach is inspired by a method of Sabanay et al. (1991) that is based on the Tokuyasu technique for immunogold labelling of sections from aldehyde-fixed samples. The difference of this method with the original Tokuyasu technique is that the immuno-labelled sections are stabilized in a thin layer of vitreous water by plunge-freezing prior to electron microscopical observation. The vitrification step allows for phase contrast-based imaging at cryogenic conditions. We show that this immuno-labelling method is well-suited for imaging cellular ultrastructure in three dimensions (tomography) at cryogenic conditions, and that fluorescence associated with the sections is retained. This method is a valuable tool for Correlative Light and Electron Microscopy (CLEM), and we refer to this method in combination with CLEM as VOS (vitrification of sections). We provide examples for the application of VOS using dendritic cells and neurons, and show specifically that this method enables the researcher to navigate to lysosomes and synapses.
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Affiliation(s)
- Erik Bos
- Department of Molecular Cell Biology, Section Electron Microscopy, Leiden University Medical Center, P.O. Box 9600, 2300 RC, Leiden, The Netherlands
| | - Leonie Hussaarts
- Department of Parasitology, Leiden University Medical Center, Leiden, The Netherlands
| | - Jan R T van Weering
- Department of Functional Genomics and Clinical Genetics, Center for Neurogenomics and Cognitive Research, Neuroscience Campus Amsterdam, VU University and VU Medical Center, De Boelelaan 1085, 1081 HV Amsterdam, The Netherlands
| | - Mark H Ellisman
- National Center for Microscopy and Imaging Research (NCMIR), Department of Neurosciences, University of California San Diego, 9500 Gilman Drive MC0608, La Jolla, CA 92093-0608, United States
| | - Heidi de Wit
- Department of Functional Genomics and Clinical Genetics, Center for Neurogenomics and Cognitive Research, Neuroscience Campus Amsterdam, VU University and VU Medical Center, De Boelelaan 1085, 1081 HV Amsterdam, The Netherlands
| | - Abraham J Koster
- Department of Molecular Cell Biology, Section Electron Microscopy, Leiden University Medical Center, P.O. Box 9600, 2300 RC, Leiden, The Netherlands.
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44
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Perkovic M, Kunz M, Endesfelder U, Bunse S, Wigge C, Yu Z, Hodirnau VV, Scheffer MP, Seybert A, Malkusch S, Schuman EM, Heilemann M, Frangakis AS. Correlative light- and electron microscopy with chemical tags. J Struct Biol 2014; 186:205-13. [PMID: 24698954 DOI: 10.1016/j.jsb.2014.03.018] [Citation(s) in RCA: 52] [Impact Index Per Article: 5.2] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/19/2014] [Revised: 03/20/2014] [Accepted: 03/24/2014] [Indexed: 10/25/2022]
Abstract
Correlative microscopy incorporates the specificity of fluorescent protein labeling into high-resolution electron micrographs. Several approaches exist for correlative microscopy, most of which have used the green fluorescent protein (GFP) as the label for light microscopy. Here we use chemical tagging and synthetic fluorophores instead, in order to achieve protein-specific labeling, and to perform multicolor imaging. We show that synthetic fluorophores preserve their post-embedding fluorescence in the presence of uranyl acetate. Post-embedding fluorescence is of such quality that the specimen can be prepared with identical protocols for scanning electron microscopy (SEM) and transmission electron microscopy (TEM); this is particularly valuable when singular or otherwise difficult samples are examined. We show that synthetic fluorophores give bright, well-resolved signals in super-resolution light microscopy, enabling us to superimpose light microscopic images with a precision of up to 25 nm in the x-y plane on electron micrographs. To exemplify the preservation quality of our new method we visualize the molecular arrangement of cadherins in adherens junctions of mouse epithelial cells.
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Affiliation(s)
- Mario Perkovic
- Goethe University Frankfurt, Buchmann Institute for Molecular Life Sciences and Institute for Biophysics, Max-von-Laue Str. 15, 60438 Frankfurt am Main, Germany
| | - Michael Kunz
- Goethe University Frankfurt, Buchmann Institute for Molecular Life Sciences and Institute for Biophysics, Max-von-Laue Str. 15, 60438 Frankfurt am Main, Germany
| | - Ulrike Endesfelder
- Goethe University Frankfurt, Institute for Physical and Theoretical Chemistry, Max-von-Laue Str. 13, 60438 Frankfurt am Main, Germany
| | - Stefanie Bunse
- Max-Planck-Institute for Brain Research, Max-von-Laue Str. 4, 60438 Frankfurt am Main, Germany
| | - Christoph Wigge
- Goethe University Frankfurt, Buchmann Institute for Molecular Life Sciences and Institute for Biophysics, Max-von-Laue Str. 15, 60438 Frankfurt am Main, Germany
| | - Zhou Yu
- Goethe University Frankfurt, Buchmann Institute for Molecular Life Sciences and Institute for Biophysics, Max-von-Laue Str. 15, 60438 Frankfurt am Main, Germany
| | - Victor-Valentin Hodirnau
- Goethe University Frankfurt, Buchmann Institute for Molecular Life Sciences and Institute for Biophysics, Max-von-Laue Str. 15, 60438 Frankfurt am Main, Germany
| | - Margot P Scheffer
- Goethe University Frankfurt, Buchmann Institute for Molecular Life Sciences and Institute for Biophysics, Max-von-Laue Str. 15, 60438 Frankfurt am Main, Germany
| | - Anja Seybert
- Goethe University Frankfurt, Buchmann Institute for Molecular Life Sciences and Institute for Biophysics, Max-von-Laue Str. 15, 60438 Frankfurt am Main, Germany
| | - Sebastian Malkusch
- Goethe University Frankfurt, Institute for Physical and Theoretical Chemistry, Max-von-Laue Str. 13, 60438 Frankfurt am Main, Germany
| | - Erin M Schuman
- Max-Planck-Institute for Brain Research, Max-von-Laue Str. 4, 60438 Frankfurt am Main, Germany
| | - Mike Heilemann
- Goethe University Frankfurt, Institute for Physical and Theoretical Chemistry, Max-von-Laue Str. 13, 60438 Frankfurt am Main, Germany
| | - Achilleas S Frangakis
- Goethe University Frankfurt, Buchmann Institute for Molecular Life Sciences and Institute for Biophysics, Max-von-Laue Str. 15, 60438 Frankfurt am Main, Germany.
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45
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Hughes L, Hawes C, Monteith S, Vaughan S. Serial block face scanning electron microscopy--the future of cell ultrastructure imaging. PROTOPLASMA 2014; 251:395-401. [PMID: 24240569 DOI: 10.1007/s00709-013-0580-1] [Citation(s) in RCA: 65] [Impact Index Per Article: 6.5] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 10/25/2013] [Accepted: 10/29/2013] [Indexed: 06/02/2023]
Abstract
One of the major drawbacks in transmission electron microscopy has been the production of three-dimensional views of cells and tissues. Currently, there is no one suitable 3D microscopy technique that answers all questions and serial block face scanning electron microscopy (SEM) fills the gap between 3D imaging using high-end fluorescence microscopy and the high resolution offered by electron tomography. In this review, we discuss the potential of the serial block face SEM technique for studying the three-dimensional organisation of animal, plant and microbial cells.
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MESH Headings
- Animals
- Cells/ultrastructure
- Humans
- Image Processing, Computer-Assisted/instrumentation
- Image Processing, Computer-Assisted/methods
- Microscopy, Electron, Scanning/instrumentation
- Microscopy, Electron, Scanning/methods
- Microscopy, Electron, Transmission/instrumentation
- Microscopy, Electron, Transmission/methods
- Microscopy, Fluorescence/instrumentation
- Microscopy, Fluorescence/methods
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Affiliation(s)
- Louise Hughes
- Department of Biological & Medical Sciences, Oxford Brookes University, Headington, Oxford, OX3 0BP, UK
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46
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McDonald KL. Rapid embedding methods into epoxy and LR White resins for morphological and immunological analysis of cryofixed biological specimens. MICROSCOPY AND MICROANALYSIS : THE OFFICIAL JOURNAL OF MICROSCOPY SOCIETY OF AMERICA, MICROBEAM ANALYSIS SOCIETY, MICROSCOPICAL SOCIETY OF CANADA 2014; 20:152-163. [PMID: 24252586 DOI: 10.1017/s1431927613013846] [Citation(s) in RCA: 38] [Impact Index Per Article: 3.8] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/02/2023]
Abstract
A variety of specimens including bacteria, ciliates, choanoflagellates (Salpingoeca rosetta), zebrafish (Danio rerio) embryos, nematode worms (Caenorhabditis elegans), and leaves of white clover (Trifolium repens) plants were high pressure frozen, freeze-substituted, infiltrated with either Epon, Epon-Araldite, or LR White resins, and polymerized. Total processing time from freezing to blocks ready to section was about 6 h. For epoxy embedding the specimens were freeze-substituted in 1% osmium tetroxide plus 0.1% uranyl acetate in acetone. For embedding in LR White the freeze-substitution medium was 0.2% uranyl acetate in acetone. Rapid infiltration was achieved by centrifugation through increasing concentrations of resin followed by polymerization at 100°C for 1.5-2 h. The preservation of ultrastructure was comparable to standard freeze substitution and resin embedding methods that take days to complete. On-section immunolabeling results for actin and tubulin molecules were positive with very low background labeling. The LR White methods offer a safer, quicker, and less-expensive alternative to Lowicryl embedding of specimens processed for on-section immunolabeling without traditional aldehyde fixatives.
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Affiliation(s)
- Kent L McDonald
- Electron Microscope Laboratory, University of California, Berkeley, CA 94720, USA
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47
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Abstract
In this chapter, we provide details along with considerations and future directions for the use of miniSOG (for mini Singlet Oxygen Generator), a versatile label for correlated light and electron microscopy of genetically tagged proteins in cells, tissues, and organisms. This new visualizable genetic tag improves the ability of biologists to locate specific proteins at nanoscale resolution and to see these tagged proteins in the environment of structural landmarks that we are used to navigating by, such as mitochondrial membranes and compartments. miniSOG provides high-quality ultrastructural preservation and permits three-dimensional protein localization via electron tomography or serial section block-face scanning electron microscopy. miniSOG is now doing for electron microscopy what the family of green fluorescent protein did for fluorescence microscopy.
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Affiliation(s)
- Guy A Perkins
- National Center for Microscopy and Imaging Research, University of California, San Diego, California, USA.
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48
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Smith EA, Cinquin BP, Do M, McDermott G, Le Gros MA, Larabell CA. Correlative cryogenic tomography of cells using light and soft x-rays. Ultramicroscopy 2013; 143:33-40. [PMID: 24355261 DOI: 10.1016/j.ultramic.2013.10.013] [Citation(s) in RCA: 19] [Impact Index Per Article: 1.7] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/15/2013] [Revised: 10/10/2013] [Accepted: 10/18/2013] [Indexed: 10/26/2022]
Abstract
Correlated imaging is the process of imaging a specimen with two complementary modalities, and then combining the two data sets to create a highly informative, composite view. A recent implementation of this concept has been the combination of soft x-ray tomography (SXT) with fluorescence cryogenic microscopy (FCM). SXT-FCM is used to visualize cells that are held in a near-native, cryopreserved. The resultant images are, therefore, highly representative of both the cellular architecture and molecular organization in vivo. SXT quantitatively visualizes the cell and sub-cellular structures; FCM images the spatial distribution of fluorescently labeled molecules. Here, we review the characteristics of SXT-FCM, and briefly discuss how this method compares with existing correlative imaging techniques. We also describe how the incorporation of a cryo-rotation stage into a cryogenic fluorescence microscope allows acquisition of fluorescence cryogenic tomography (FCT) data. FCT is optimally suited for correlation with SXT, since both techniques image the specimen in 3-D, potentially with similar, isotropic spatial resolution.
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Affiliation(s)
- Elizabeth A Smith
- Department of Anatomy, School of Medicine, University of California San Francisco, San Francisco, CA, United States; National Center for X-ray Tomography, Advanced Light Source, Berkeley, CA, United States
| | - Bertrand P Cinquin
- Department of Anatomy, School of Medicine, University of California San Francisco, San Francisco, CA, United States; National Center for X-ray Tomography, Advanced Light Source, Berkeley, CA, United States
| | - Myan Do
- Department of Anatomy, School of Medicine, University of California San Francisco, San Francisco, CA, United States; National Center for X-ray Tomography, Advanced Light Source, Berkeley, CA, United States
| | - Gerry McDermott
- Department of Anatomy, School of Medicine, University of California San Francisco, San Francisco, CA, United States; National Center for X-ray Tomography, Advanced Light Source, Berkeley, CA, United States
| | - Mark A Le Gros
- Department of Anatomy, School of Medicine, University of California San Francisco, San Francisco, CA, United States; Physical BioSciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA, United States; National Center for X-ray Tomography, Advanced Light Source, Berkeley, CA, United States.
| | - Carolyn A Larabell
- Department of Anatomy, School of Medicine, University of California San Francisco, San Francisco, CA, United States; Physical BioSciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA, United States; National Center for X-ray Tomography, Advanced Light Source, Berkeley, CA, United States.
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49
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Das T, Hoshijima M. Adding a new dimension to cardiac nano-architecture using electron microscopy: coupling membrane excitation to calcium signaling. J Mol Cell Cardiol 2012. [PMID: 23201225 DOI: 10.1016/j.yjmcc.2012.11.009] [Citation(s) in RCA: 13] [Impact Index Per Article: 1.1] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Submit a Manuscript] [Subscribe] [Scholar Register] [Indexed: 10/27/2022]
Abstract
Advances in microscopic imaging technologies and associated computational methods now allow descriptions of cellular anatomy to go beyond 2-dimensions, revealing new micro-domain dynamics at unprecedented resolutions. In cardiomyocytes, electron microscopy (EM) first described junctional membrane complexes between the sarcolemma and sarcoplasmic reticulum over a half-century ago. Since then, 3-dimensional EM technologies such as electron tomography have become successful in determining the realistic nano-geometry of membrane junctions (dyads and peripheral junctions) and associated structures such as transverse tubules (T-tubules, aka. T-system). Concomitantly, super-resolution light microscopy has gone beyond the diffraction-limit to determine the distribution of molecules, such as ryanodine receptors, with 10(-8) meter (10nm) order accuracy. This review provides the current structural perspective and functional interpretation of membrane junction complexes, which are the central machinery controlling cardiac excitation-contraction coupling via calcium signaling.
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Affiliation(s)
- Tapaswini Das
- The Center for Research in Biological Systems, University of California San Diego, La Jolla, CA 92093, USA
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50
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Martell JD, Deerinck TJ, Sancak Y, Poulos TL, Mootha VK, Sosinsky GE, Ellisman MH, Ting AY. Engineered ascorbate peroxidase as a genetically encoded reporter for electron microscopy. Nat Biotechnol 2012; 30:1143-8. [PMID: 23086203 PMCID: PMC3699407 DOI: 10.1038/nbt.2375] [Citation(s) in RCA: 491] [Impact Index Per Article: 40.9] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/29/2012] [Accepted: 08/29/2012] [Indexed: 11/09/2022]
Abstract
Electron microscopy (EM) is the standard method for imaging cellular structures with nanometer resolution, but existing genetic tags are inactive in most cellular compartments or require light and can be difficult to use. Here we report the development of 'APEX', a genetically encodable EM tag that is active in all cellular compartments and does not require light. APEX is a monomeric 28-kDa peroxidase that withstands strong EM fixation to give excellent ultrastructural preservation. We demonstrate the utility of APEX for high-resolution EM imaging of a variety of mammalian organelles and specific proteins using a simple and robust labeling procedure. We also fused APEX to the N or C terminus of the mitochondrial calcium uniporter (MCU), a recently identified channel whose topology is disputed. These fusions give EM contrast exclusively in the mitochondrial matrix, suggesting that both the N and C termini of MCU face the matrix. Because APEX staining is not dependent on light activation, APEX should make EM imaging of any cellular protein straightforward, regardless of the size or thickness of the specimen.
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Affiliation(s)
- Jeffrey D Martell
- Department of Chemistry, Massachusetts Institute of Technology, Cambridge, Massachusetts, USA
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