1
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Lou J, Deng Q, Zhang X, Bell CC, Das AB, Bediaga NG, Zlatic CO, Johanson TM, Allan RS, Griffin MDW, Paradkar P, Harvey KF, Dawson MA, Hinde E. Heterochromatin protein 1 alpha (HP1α) undergoes a monomer to dimer transition that opens and compacts live cell genome architecture. Nucleic Acids Res 2024:gkae720. [PMID: 39193905 DOI: 10.1093/nar/gkae720] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/10/2023] [Revised: 07/29/2024] [Accepted: 08/07/2024] [Indexed: 08/29/2024] Open
Abstract
Our understanding of heterochromatin nanostructure and its capacity to mediate gene silencing in a living cell has been prevented by the diffraction limit of optical microscopy. Thus, here to overcome this technical hurdle, and directly measure the nucleosome arrangement that underpins this dense chromatin state, we coupled fluorescence lifetime imaging microscopy (FLIM) of Förster resonance energy transfer (FRET) between histones core to the nucleosome, with molecular editing of heterochromatin protein 1 alpha (HP1α). Intriguingly, this super-resolved readout of nanoscale chromatin structure, alongside fluorescence fluctuation spectroscopy (FFS) and FLIM-FRET analysis of HP1α protein-protein interaction, revealed nucleosome arrangement to be differentially regulated by HP1α oligomeric state. Specifically, we found HP1α monomers to impart a previously undescribed global nucleosome spacing throughout genome architecture that is mediated by trimethylation on lysine 9 of histone H3 (H3K9me3) and locally reduced upon HP1α dimerisation. Collectively, these results demonstrate HP1α to impart a dual action on chromatin that increases the dynamic range of nucleosome proximity. We anticipate that this finding will have important implications for our understanding of how live cell heterochromatin structure regulates genome function.
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Affiliation(s)
- Jieqiong Lou
- School of Physics, University of Melbourne, Melbourne, VIC 3010, Australia
| | - Qiji Deng
- Peter MacCallum Cancer Centre, 305 Grattan St, Melbourne, VIC 3000, Australia
| | - Xiaomeng Zhang
- School of Physics, University of Melbourne, Melbourne, VIC 3010, Australia
| | - Charles C Bell
- Peter MacCallum Cancer Centre, 305 Grattan St, Melbourne, VIC 3000, Australia
| | - Andrew B Das
- Peter MacCallum Cancer Centre, 305 Grattan St, Melbourne, VIC 3000, Australia
- Sir Peter MacCallum Department of Oncology, University of Melbourne, Parkville, VIC 3010, Australia
| | - Naiara G Bediaga
- Peter MacCallum Cancer Centre, 305 Grattan St, Melbourne, VIC 3000, Australia
| | - Courtney O Zlatic
- Department of Biochemistry and Pharmacology, Bio21 Molecular Science and Biotechnology Institute, University of Melbourne, Melbourne, VIC 3010, Australia
| | - Timothy M Johanson
- The Walter and Eliza Hall Institute of Medical Research, Parkville, VIC 3052, Australia
- Department of Medical Biology, The University of Melbourne, Parkville, VIC 3010, Australia
| | - Rhys S Allan
- The Walter and Eliza Hall Institute of Medical Research, Parkville, VIC 3052, Australia
- Department of Medical Biology, The University of Melbourne, Parkville, VIC 3010, Australia
| | - Michael D W Griffin
- Department of Biochemistry and Pharmacology, Bio21 Molecular Science and Biotechnology Institute, University of Melbourne, Melbourne, VIC 3010, Australia
| | - PrasadN Paradkar
- CSIRO Health & Biosecurity, Australian Centre for Disease Preparedness, 5 Portarlington Road, Geelong3220, Australia
| | - Kieran F Harvey
- Peter MacCallum Cancer Centre, 305 Grattan St, Melbourne, VIC 3000, Australia
- Sir Peter MacCallum Department of Oncology, University of Melbourne, Parkville, VIC 3010, Australia
- Department of Anatomy and Developmental Biology and Biomedicine Discovery Institute, Monash University, Clayton, VIC 3168, Australia
| | - Mark A Dawson
- Peter MacCallum Cancer Centre, 305 Grattan St, Melbourne, VIC 3000, Australia
- Sir Peter MacCallum Department of Oncology, University of Melbourne, Parkville, VIC 3010, Australia
- Centre for Cancer Research, University of Melbourne, Melbourne, VIC 3010, Australia
| | - Elizabeth Hinde
- School of Physics, University of Melbourne, Melbourne, VIC 3010, Australia
- Department of Biochemistry and Pharmacology, Bio21 Molecular Science and Biotechnology Institute, University of Melbourne, Melbourne, VIC 3010, Australia
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2
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Liang Z, Solano A, Lou J, Hinde E. Histone FRET reports the spatial heterogeneity in nanoscale chromatin architecture that is imparted by the epigenetic landscape at the level of single foci in an intact cell nucleus. Chromosoma 2024; 133:5-14. [PMID: 38265456 PMCID: PMC10904561 DOI: 10.1007/s00412-024-00815-z] [Citation(s) in RCA: 2] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/21/2023] [Revised: 12/28/2023] [Accepted: 01/03/2024] [Indexed: 01/25/2024]
Abstract
Genome sequencing has identified hundreds of histone post-translational modifications (PTMs) that define an open or compact chromatin nanostructure at the level of nucleosome proximity, and therefore serve as activators or repressors of gene expression. Direct observation of this epigenetic mode of transcriptional regulation in an intact single nucleus, is however, a complex task. This is because despite the development of fluorescent probes that enable observation of specific histone PTMs and chromatin density, the changes in nucleosome proximity regulating gene expression occur on a spatial scale well below the diffraction limit of optical microscopy. In recent work, to address this research gap, we demonstrated that the phasor approach to fluorescence lifetime imaging microscopy (FLIM) of Förster resonance energy transfer (FRET) between fluorescently labelled histones core to the nucleosome, is a readout of chromatin nanostructure that can be multiplexed with immunofluorescence (IF) against specific histone PTMs. Here from application of this methodology to gold standard gene activators (H3K4Me3 and H3K9Ac) versus repressors (e.g., H3K9Me3 and H3K27Me), we find that while on average these histone marks do impart an open versus compact chromatin nanostructure, at the level of single chromatin foci, there is significant spatial heterogeneity. Collectively this study illustrates the importance of studying the epigenetic landscape as a function of space within intact nuclear architecture and opens the door for the study of chromatin foci sub-populations defined by combinations of histone marks, as is seen in the context of bivalent chromatin.
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Affiliation(s)
- Zhen Liang
- Department of Biochemistry and Pharmacology, University of Melbourne, Melbourne, VIC, Australia
- Cancer and RNA Laboratory, St. Vincent's Institute of Medical Research, Melbourne, VIC, Australia
- Department of Medicine, Melbourne Medical School, St Vincent's Hospital, University of Melbourne, Melbourne, VIC, Australia
| | - Ashleigh Solano
- School of Physics, University of Melbourne, Melbourne, VIC, Australia
| | - Jieqiong Lou
- School of Physics, University of Melbourne, Melbourne, VIC, Australia
| | - Elizabeth Hinde
- Department of Biochemistry and Pharmacology, University of Melbourne, Melbourne, VIC, Australia.
- School of Physics, University of Melbourne, Melbourne, VIC, Australia.
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3
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Díaz M, Malacrida L. Advanced Fluorescence Microscopy Methods to Study Dynamics of Fluorescent Proteins In Vivo. Methods Mol Biol 2023; 2564:53-74. [PMID: 36107337 DOI: 10.1007/978-1-0716-2667-2_3] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 06/15/2023]
Abstract
Fluorescent proteins are standard tools for addressing biological questions in a cell biology laboratory. The genetic tagging of protein of interest with fluorescent proteins opens the opportunity to follow them in vivo and to understand their interactions and dynamics. In addition, the latest advances in optical microscopy image acquisition and processing allow us to study many cellular processes in vivo. Techniques such as fluorescence lifetime microscopy and hyperspectral imaging provide valuable tools for understanding fluorescent protein interactions and their photophysics. Finally, fluorescence fluctuation analysis opens the possibility to address questions of molecular diffusion, protein-protein interactions, and oligomerization, among others, yielding quantitative information on the subject of study. This chapter will cover some of the more important advances in cutting-edge technologies and methods that, combined with fluorescent proteins, open new frontiers for biological studies.
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Affiliation(s)
- Marcela Díaz
- Advanced Bioimaging Unit, Institut Pasteur of Montevideo & Universidad de la República, Montevideo, Uruguay
| | - Leonel Malacrida
- Advanced Bioimaging Unit, Institut Pasteur of Montevideo & Universidad de la República, Montevideo, Uruguay.
- Departamento de Fisiopatología, Hospital de Clínicas, Facultad de Medicina, Universidad de la República, Montevideo, Uruguay.
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4
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Osterlund EJ, Hirmiz N, Pemberton JM, Nougarède A, Liu Q, Leber B, Fang Q, Andrews DW. Efficacy and specificity of inhibitors of BCL-2 family protein interactions assessed by affinity measurements in live cells. SCIENCE ADVANCES 2022; 8:eabm7375. [PMID: 35442739 PMCID: PMC9020777 DOI: 10.1126/sciadv.abm7375] [Citation(s) in RCA: 7] [Impact Index Per Article: 3.5] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Subscribe] [Scholar Register] [Received: 10/12/2021] [Accepted: 03/01/2022] [Indexed: 06/14/2023]
Abstract
Cytoplasmic and membrane-bound BCL-2 family proteins regulate apoptosis, a form of programmed cell death, via dozens of binary protein interactions confounding measurement of the effects of inhibitors in live cells. In cancer, apoptosis is frequently dysregulated, and cell survival depends on antiapoptotic proteins binding to and inhibiting proapoptotic BH3 proteins. The clinical success of BH3 mimetic inhibitors of antiapoptotic proteins has spawned major efforts by the pharmaceutical industry to develop molecules with different specificities and higher affinities. Here, quantitative fast fluorescence lifetime imaging microscopy enabled comparison of BH3 mimetic drugs in trials and preclinical development by measuring drug effects on binding affinities of interacting protein pairs in live cells. Both selectivity and efficacy were assessed for 15 inhibitors of four antiapoptotic proteins for each of six BH3 protein ligands. While many drugs target the designed interaction, most also have unexpected selectivity and poor efficacy in cells.
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Affiliation(s)
- Elizabeth J. Osterlund
- Biological Sciences, Sunnybrook Research Institute, Toronto, Ontario M4N 3M5, Canada
- Department of Biochemistry, University of Toronto, Toronto, Ontario M5S 2J7, Canada
| | - Nehad Hirmiz
- Biological Sciences, Sunnybrook Research Institute, Toronto, Ontario M4N 3M5, Canada
- School of Biomedical Engineering, McMaster University, Hamilton, Ontario L8S 4L7, Canada
| | - James M. Pemberton
- Biological Sciences, Sunnybrook Research Institute, Toronto, Ontario M4N 3M5, Canada
- Department of Medical Biophysics, University of Toronto, Toronto, Ontario M5S 2J7, Canada
| | - Adrien Nougarède
- Biological Sciences, Sunnybrook Research Institute, Toronto, Ontario M4N 3M5, Canada
| | - Qian Liu
- Biological Sciences, Sunnybrook Research Institute, Toronto, Ontario M4N 3M5, Canada
| | - Brian Leber
- Department of Medicine, McMaster University, Hamilton, Ontario L8N 3Z5, Canada
| | - Qiyin Fang
- School of Biomedical Engineering, McMaster University, Hamilton, Ontario L8S 4L7, Canada
- Department of Engineering Physics, McMaster University, 1280 Main Street West, Hamilton, Ontario L8S 4L7, Canada
| | - David W. Andrews
- Biological Sciences, Sunnybrook Research Institute, Toronto, Ontario M4N 3M5, Canada
- Department of Biochemistry, University of Toronto, Toronto, Ontario M5S 2J7, Canada
- Department of Medical Biophysics, University of Toronto, Toronto, Ontario M5S 2J7, Canada
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5
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Lou J, Solano A, Liang Z, Hinde E. Phasor Histone FLIM-FRET Microscopy Maps Nuclear-Wide Nanoscale Chromatin Architecture With Respect to Genetically Induced DNA Double-Strand Breaks. Front Genet 2021; 12:770081. [PMID: 34956323 PMCID: PMC8702996 DOI: 10.3389/fgene.2021.770081] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/03/2021] [Accepted: 11/10/2021] [Indexed: 12/30/2022] Open
Abstract
A DNA double-strand break (DSB) takes place in the context of chromatin, and there is increasing evidence for chromatin structure to play a functional role in DSB signaling and repair. Thus, there is an emerging need for quantitative microscopy methods that can directly measure chromatin network architecture and detect changes in this structural framework upon DSB induction within an intact nucleus. To address this demand, here we present the phasor approach to fluorescence lifetime imaging microscopy (FLIM) of Förster resonance energy transfer (FRET) between fluorescently labeled histones in the DSB inducible via AsiSI cell system (DIvA), which has sufficient spatial resolution to map nuclear-wide chromatin compaction at the level of nucleosome proximity with respect to multiple site-specific DSBs. We also demonstrate that when phasor histone FLIM-FRET is coupled with immunofluorescence, this technology has the unique advantage of enabling exploration of any heterogeneity that exists in chromatin structure at the spatially distinct and genetically induced DSBs.
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Affiliation(s)
- Jieqiong Lou
- School of Physics, University of Melbourne, Melbourne, VIC, Australia.,Department of Biochemistry and Pharmacology, University of Melbourne, Melbourne, VIC, Australia
| | - Ashleigh Solano
- School of Physics, University of Melbourne, Melbourne, VIC, Australia.,Department of Biochemistry and Pharmacology, University of Melbourne, Melbourne, VIC, Australia
| | - Zhen Liang
- Cancer and RNA Laboratory, St. Vincent's Institute of Medical Research, Fitzroy, VIC, Australia.,Department of Medicine, Melbourne Medical School, St Vincent's Hospital, University of Melbourne, Fitzroy, VIC, Australia
| | - Elizabeth Hinde
- School of Physics, University of Melbourne, Melbourne, VIC, Australia.,Department of Biochemistry and Pharmacology, University of Melbourne, Melbourne, VIC, Australia
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6
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Datta R, Gillette A, Stefely M, Skala MC. Recent innovations in fluorescence lifetime imaging microscopy for biology and medicine. JOURNAL OF BIOMEDICAL OPTICS 2021; 26:JBO-210093-PER. [PMID: 34247457 PMCID: PMC8271181 DOI: 10.1117/1.jbo.26.7.070603] [Citation(s) in RCA: 18] [Impact Index Per Article: 6.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 03/30/2021] [Accepted: 06/11/2021] [Indexed: 05/05/2023]
Abstract
SIGNIFICANCE Fluorescence lifetime imaging microscopy (FLIM) measures the decay rate of fluorophores, thus providing insights into molecular interactions. FLIM is a powerful molecular imaging technique that is widely used in biology and medicine. AIM This perspective highlights some of the major advances in FLIM instrumentation, analysis, and biological and clinical applications that we have found impactful over the last year. APPROACH Innovations in FLIM instrumentation resulted in faster acquisition speeds, rapid imaging over large fields of view, and integration with complementary modalities such as single-molecule microscopy or light-sheet microscopy. There were significant developments in FLIM analysis with machine learning approaches to enhance processing speeds, fit-free techniques to analyze images without a priori knowledge, and open-source analysis resources. The advantages and limitations of these recent instrumentation and analysis techniques are summarized. Finally, applications of FLIM in the last year include label-free imaging in biology, ophthalmology, and intraoperative imaging, FLIM of new fluorescent probes, and lifetime-based Förster resonance energy transfer measurements. CONCLUSIONS A large number of high-quality publications over the last year signifies the growing interest in FLIM and ensures continued technological improvements and expanding applications in biomedical research.
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Affiliation(s)
- Rupsa Datta
- Morgridge Institute for Research, Madison, Wisconsin, United States
| | - Amani Gillette
- Morgridge Institute for Research, Madison, Wisconsin, United States
- University of Wisconsin, Department of Biomedical Engineering, Madison, Wisconsin, United States
| | - Matthew Stefely
- Morgridge Institute for Research, Madison, Wisconsin, United States
| | - Melissa C. Skala
- Morgridge Institute for Research, Madison, Wisconsin, United States
- University of Wisconsin, Department of Biomedical Engineering, Madison, Wisconsin, United States
- Address all correspondence to Melissa C. Skala,
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7
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Vallmitjana A, Torrado B, Gratton E. Phasor-based image segmentation: machine learning clustering techniques. BIOMEDICAL OPTICS EXPRESS 2021; 12:3410-3422. [PMID: 34221668 PMCID: PMC8221971 DOI: 10.1364/boe.422766] [Citation(s) in RCA: 13] [Impact Index Per Article: 4.3] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 02/24/2021] [Revised: 04/20/2021] [Accepted: 04/22/2021] [Indexed: 05/30/2023]
Abstract
The phasor approach is a well-established method for data visualization and image analysis in spectral and lifetime fluorescence microscopy. Nevertheless, it is typically applied in a user-dependent manner by manually selecting regions of interest on the phasor space to find distinct regions in the fluorescence images. In this paper we present our work on using machine learning clustering techniques to establish an unsupervised and automatic method that can be used for identifying populations of fluorescent species in spectral and lifetime imaging. We demonstrate our method using both synthetic data, created by sampling photon arrival times and plotting the distributions on the phasor plot, and real live cells samples, by staining cellular organelles with a selection of commercial probes.
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Affiliation(s)
- Alex Vallmitjana
- Laboratory for Fluorescence Dynamics, Biomedical Engineering, University of California, Irvine, CA 92697, USA
| | - Belén Torrado
- Laboratory for Fluorescence Dynamics, Biomedical Engineering, University of California, Irvine, CA 92697, USA
| | - Enrico Gratton
- Laboratory for Fluorescence Dynamics, Biomedical Engineering, University of California, Irvine, CA 92697, USA
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8
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Priest DG, Bernardini A, Lou J, Mantovani R, Hinde E. Live cell dynamics of the NF-Y transcription factor. Sci Rep 2021; 11:10992. [PMID: 34040015 PMCID: PMC8155045 DOI: 10.1038/s41598-021-90081-1] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/16/2020] [Accepted: 04/29/2021] [Indexed: 11/24/2022] Open
Abstract
Transcription factors (TFs) are core players in the control of gene expression, evolutionarily selected to recognise a subset of specific DNA sequences and nucleate the recruitment of the transcriptional machinery. How TFs assemble and move in the nucleus to locate and bind their DNA targets and cause a transcriptional response, remains mostly unclear. NF-Y is a highly conserved, heterotrimeric TF with important roles in both housekeeping and lineage-specific gene expression, functioning as a promoter organiser. Despite a large number of biochemical, structural and genomic studies of NF-Y, there is a lack of experiments in single living cells; therefore, basic assumptions of NF-Y biology remain unproven in vivo. Here we employ a series of dynamic fluorescence microscopy methods (FLIM-FRET, NB, RICS and FRAP) to study NF-Y dynamics and complex formation in live cells. Specifically, we provide quantitative measurement of NF-Y subunit association and diffusion kinetics in the nucleus that collectively suggest NF-Y to move and bind chromatin as a trimeric complex in vivo.
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Affiliation(s)
- David G Priest
- School of Physics, University of Melbourne, Melbourne, VIC, Australia
- Department of Biochemistry and Molecular Biology, Bio21 Institute, University of Melbourne, Melbourne, VIC, Australia
| | - Andrea Bernardini
- Dipartimento di Bioscienze, Università degli Studi di Milano, Via Celoria 26, 20133, Milan, Italy
| | - Jieqiong Lou
- School of Physics, University of Melbourne, Melbourne, VIC, Australia
| | - Roberto Mantovani
- Dipartimento di Bioscienze, Università degli Studi di Milano, Via Celoria 26, 20133, Milan, Italy.
| | - Elizabeth Hinde
- School of Physics, University of Melbourne, Melbourne, VIC, Australia.
- Department of Biochemistry and Molecular Biology, Bio21 Institute, University of Melbourne, Melbourne, VIC, Australia.
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9
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Fluorescence Fluctuation Spectroscopy enables quantification of potassium channel subunit dynamics and stoichiometry. Sci Rep 2021; 11:10719. [PMID: 34021177 PMCID: PMC8140153 DOI: 10.1038/s41598-021-90002-2] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/04/2020] [Accepted: 04/15/2021] [Indexed: 11/08/2022] Open
Abstract
Voltage-gated potassium (Kv) channels are a family of membrane proteins that facilitate K+ ion diffusion across the plasma membrane, regulating both resting and action potentials. Kv channels comprise four pore-forming α subunits, each with a voltage sensing domain, and they are regulated by interaction with β subunits such as those belonging to the KCNE family. Here we conducted a comprehensive biophysical characterization of stoichiometry and protein diffusion across the plasma membrane of the epithelial KCNQ1-KCNE2 complex, combining total internal reflection fluorescence (TIRF) microscopy and a series of complementary Fluorescence Fluctuation Spectroscopy (FFS) techniques. Using this approach, we found that KCNQ1-KCNE2 has a predominant 4:4 stoichiometry, while non-bound KCNE2 subunits are mostly present as dimers in the plasma membrane. At the same time, we identified unique spatio-temporal diffusion modalities and nano-environment organization for each channel subunit. These findings improve our understanding of KCNQ1-KCNE2 channel function and suggest strategies for elucidating the subunit stoichiometry and forces directing localization and diffusion of ion channel complexes in general.
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10
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Tavakoli M, Jazani S, Sgouralis I, Heo W, Ishii K, Tahara T, Pressé S. Direct Photon-by-Photon Analysis of Time-Resolved Pulsed Excitation Data using Bayesian Nonparametrics. CELL REPORTS. PHYSICAL SCIENCE 2020; 1:100234. [PMID: 34414380 PMCID: PMC8373049 DOI: 10.1016/j.xcrp.2020.100234] [Citation(s) in RCA: 9] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 05/18/2023]
Abstract
Lifetimes of chemical species are typically estimated by either fitting time-correlated single-photon counting (TCSPC) histograms or phasor analysis from time-resolved photon arrivals. While both methods yield lifetimes in a computationally efficient manner, their performance is limited by choices made on the number of distinct chemical species contributing photons. However, the number of species is encoded in the photon arrival times collected for each illuminated spot and need not be set by hand a priori. Here, we propose a direct photon-by-photon analysis of data drawn from pulsed excitation experiments to infer, simultaneously and self-consistently, the number of species and their associated lifetimes from a few thousand photons. We do so by leveraging new mathematical tools within the Bayesian nonparametric. We benchmark our method for both simulated and experimental data for 1-4 species.
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Affiliation(s)
- Meysam Tavakoli
- Department of Physics, Indiana University-Purdue University, Indianapolis, IN 46202, USA
| | - Sina Jazani
- Center for Biological Physics, Department of Physics, Arizona State University, Tempe, AZ 85287, USA
| | - Ioannis Sgouralis
- Center for Biological Physics, Department of Physics, Arizona State University, Tempe, AZ 85287, USA
| | - Wooseok Heo
- Molecular Spectroscopy Laboratory, RIKEN, 2-1 Hirosawa, Wako, Saitama 351-0198, Japan
| | - Kunihiko Ishii
- Molecular Spectroscopy Laboratory, RIKEN, 2-1 Hirosawa, Wako, Saitama 351-0198, Japan
- Ultrafast Spectroscopy Research Team, RIKEN Center for Advanced Photonics (RAP), 2-1 Hirosawa, Wako, Saitama 351-0198, Japan
| | - Tahei Tahara
- Molecular Spectroscopy Laboratory, RIKEN, 2-1 Hirosawa, Wako, Saitama 351-0198, Japan
- Ultrafast Spectroscopy Research Team, RIKEN Center for Advanced Photonics (RAP), 2-1 Hirosawa, Wako, Saitama 351-0198, Japan
| | - Steve Pressé
- Center for Biological Physics, Department of Physics, Arizona State University, Tempe, AZ 85287, USA
- School of Molecular Sciences, Arizona State University, Tempe, AZ 85287, USA
- Lead Contact
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Coppola S, Antonacci G, Lanzanò L. High-resolution microscopy and spectroscopy datasets meet Data in Brief. Data Brief 2020; 30:105596. [PMID: 32382608 PMCID: PMC7200866 DOI: 10.1016/j.dib.2020.105596] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Track Full Text] [Download PDF] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/03/2022] Open
Affiliation(s)
- Stefano Coppola
- Division of Gene Regulation, The Netherlands Cancer Institute (NKI), Oncode Institute, Amsterdam, Plesmanlaan 121, 1066 CX Amsterdam, The Netherlands
| | - Giuseppe Antonacci
- Photonics Research Group, INTEC, Ghent University-imec, Ghent 9052, Belgium
- Present address: Dipartimento di Fisica, Politecnico di Milano, Piazza Leonardo da Vinci 32, I-20133 Milano, Italy
| | - Luca Lanzanò
- Dipartimento di Fisica e Astronomia "Ettore Majorana", Università degli Studi di Catania, Via S. Sofia, 64, 95123 Catania, Italy
- Nanoscopy and NIC@IIT, Istituto Italiano di Tecnologia, Via Melen 83, Genoa, Italy
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