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Yan Z, Kavanagh T, Harrabi RDS, Lust ST, Tang C, Beavil R, Müller MM, Beavil A, Ameer-Beg S, da Silva RM, Gentleman E. FRET Sensor-Modified Synthetic Hydrogels for Real-Time Monitoring of Cell-Derived Matrix Metalloproteinase Activity using Fluorescence Lifetime Imaging. ADVANCED FUNCTIONAL MATERIALS 2024; 34:adfm.202309711. [PMID: 38779415 PMCID: PMC7615971 DOI: 10.1002/adfm.202309711] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Figures] [Subscribe] [Scholar Register] [Received: 08/15/2023] [Indexed: 05/25/2024]
Abstract
Matrix remodeling plays central roles in a range of physiological and pathological processes and is driven predominantly by the activity of matrix metalloproteinases (MMPs), which degrade extracellular matrix (ECM) proteins. Our understanding of how MMPs regulate cell and tissue dynamics is often incomplete as in vivo approaches are lacking and many in vitro strategies cannot provide high-resolution, quantitative measures of enzyme activity in situ within tissue-like 3D microenvironments. Here, we incorporate a Förster resonance energy transfer (FRET) sensor of MMP activity into fully synthetic hydrogels that mimic many properties of the native ECM. We then use fluorescence lifetime imaging to provide a real-time, fluorophore concentration-independent quantification of MMP activity, establishing a highly accurate, readily adaptable platform for studying MMP dynamics in situ. MCF7 human breast cancer cells encapsulated within hydrogels highlight the detection of MMP activity both locally, at the sub-micron level, and within the bulk hydrogel. Our versatile platform may find use in a range of biological studies to explore questions in the dynamics of cancer metastasis, development, and tissue repair by providing high-resolution, quantitative and in situ readouts of local MMP activity within native tissue-like environments.
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Affiliation(s)
- Ziqian Yan
- Centre for Craniofacial and Regenerative Biology, King’s College London, London, UK
| | - Thomas Kavanagh
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer and Pharmaceutical Sciences, King’s College London, London, UK
| | | | - Suzette T. Lust
- Centre for Craniofacial and Regenerative Biology, King’s College London, London, UK
| | - Chunling Tang
- Centre for Craniofacial and Regenerative Biology, King’s College London, London, UK
| | - Rebecca Beavil
- Randall Centre of Cell and Molecular Biophysics, King’s College London, London, UK
| | | | - Andrew Beavil
- Randall Centre of Cell and Molecular Biophysics, King’s College London, London, UK
| | - Simon Ameer-Beg
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer and Pharmaceutical Sciences, King’s College London, London, UK
| | | | - Eileen Gentleman
- Centre for Craniofacial and Regenerative Biology, King’s College London, London, UK
- Department of Biomedical Sciences, University of Lausanne, Lausanne, Switzerland
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2
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Balukova A, Bokea K, Barber PR, Ameer-Beg SM, MacRobert AJ, Yaghini E. Cellular Imaging and Time-Domain FLIM Studies of Meso-Tetraphenylporphine Disulfonate as a Photosensitising Agent in 2D and 3D Models. Int J Mol Sci 2024; 25:4222. [PMID: 38673807 PMCID: PMC11050357 DOI: 10.3390/ijms25084222] [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] [Received: 03/04/2024] [Revised: 03/28/2024] [Accepted: 03/29/2024] [Indexed: 04/28/2024] Open
Abstract
Fluorescence lifetime imaging (FLIM) and confocal fluorescence studies of a porphyrin-based photosensitiser (meso-tetraphenylporphine disulfonate: TPPS2a) were evaluated in 2D monolayer cultures and 3D compressed collagen constructs of a human ovarian cancer cell line (HEY). TPPS2a is known to be an effective model photosensitiser for both Photodynamic Therapy (PDT) and Photochemical Internalisation (PCI). This microspectrofluorimetric study aimed firstly to investigate the uptake and subcellular localisation of TPPS2a, and evaluate the photo-oxidative mechanism using reactive oxygen species (ROS) and lipid peroxidation probes combined with appropriate ROS scavengers. Light-induced intracellular redistribution of TPPS2a was observed, consistent with rupture of endolysosomes where the porphyrin localises. Using the same range of light doses, time-lapse confocal imaging permitted observation of PDT-induced generation of ROS in both 2D and 3D cancer models using fluorescence-based ROS together with specific ROS inhibitors. In addition, the use of red light excitation of the photosensitiser to minimise auto-oxidation of the probes was investigated. In the second part of the study, the photophysical properties of TPPS2a in cells were studied using a time-domain FLIM system with time-correlated single photon counting detection. Owing to the high sensitivity and spatial resolution of this system, we acquired FLIM images that enabled the fluorescence lifetime determination of the porphyrin within the endolysosomal vesicles. Changes in the lifetime dynamics upon prolonged illumination were revealed as the vesicles degraded within the cells.
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Affiliation(s)
- Andrea Balukova
- Department of Surgical Biotechnology, Division of Surgery and Interventional Science, University College London, London NW3 2QG, UK; (A.B.); (K.B.)
| | - Kalliopi Bokea
- Department of Surgical Biotechnology, Division of Surgery and Interventional Science, University College London, London NW3 2QG, UK; (A.B.); (K.B.)
| | - Paul R. Barber
- Department of Oncology, UCL Cancer Institute, University College London, London WC1E 6DD, UK;
- Comprehensive Cancer Centre, School of Cancer and Pharmaceutical Sciences, King’s College London, London SE1 9RT, UK;
| | - Simon M. Ameer-Beg
- Comprehensive Cancer Centre, School of Cancer and Pharmaceutical Sciences, King’s College London, London SE1 9RT, UK;
| | - Alexander J. MacRobert
- Department of Surgical Biotechnology, Division of Surgery and Interventional Science, University College London, London NW3 2QG, UK; (A.B.); (K.B.)
| | - Elnaz Yaghini
- Department of Surgical Biotechnology, Division of Surgery and Interventional Science, University College London, London NW3 2QG, UK; (A.B.); (K.B.)
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3
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Wilda CB, Burnstock A, Suhling K, Mattioli Della Rocca F, Henderson RK, Nedbal J. Visualising varnish removal for conservation of paintings by fluorescence lifetime imaging (FLIM). HERITAGE SCIENCE 2023; 11:127. [PMID: 37333623 PMCID: PMC10276100 DOI: 10.1186/s40494-023-00957-w] [Citation(s) in RCA: 1] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Figures] [Subscribe] [Scholar Register] [Received: 01/27/2023] [Accepted: 05/17/2023] [Indexed: 06/20/2023]
Abstract
The removal of varnish from the surface is a key step in painting conservation. Varnish removal is traditionally monitored by examining the painting surface under ultraviolet illumination. We show here that by imaging the fluorescence lifetime instead, much better contrast, sensitivity, and specificity can be achieved. For this purpose, we developed a lightweight (4.8 kg) portable instrument for macroscopic fluorescence lifetime imaging (FLIM). It is based on a time-correlated single-photon avalanche diode (SPAD) camera to acquire the FLIM images and a pulsed 440 nm diode laser to excite the varnish fluorescence. A historical model painting was examined to demonstrate the capabilities of the system. We found that the FLIM images provided information on the distribution of the varnish on the painting surface with greater sensitivity, specificity, and contrast compared to the traditional ultraviolet illumination photography. The distribution of the varnish and other painting materials was assessed using FLIM during and after varnish removal with different solvent application methods. Monitoring of the varnish removal process between successive solvent applications by a swab revealed an evolving image contrast as a function of the cleaning progress. FLIM of dammar and mastic resin varnishes identified characteristic changes to their fluorescence lifetimes depending on their ageing conditions. Thus, FLIM has a potential to become a powerful and versatile tool to visualise varnish removal from paintings. Graphical Abstract
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Affiliation(s)
- Christine B. Wilda
- Department of Physics, King’s College London, Strand, London, WC2R 2LS United Kingdom
- The Courtauld, Somerset House, Strand, London, WC1X 0RN United Kingdom
- ConservArt, 6620 E Rogers Cir, Boca Raton, FL 33487 United States
| | - Aviva Burnstock
- The Courtauld, Somerset House, Strand, London, WC1X 0RN United Kingdom
| | - Klaus Suhling
- Department of Physics, King’s College London, Strand, London, WC2R 2LS United Kingdom
| | - Francesco Mattioli Della Rocca
- School of Engineering, University of Edinburgh, King’s Buildings, Edinburgh, EH9 3JL United Kingdom
- Europe Technology Development Centre, Sony Semiconductor Solutions - Sony Europe B.V., Trento, Italy
| | - Robert K. Henderson
- School of Engineering, University of Edinburgh, King’s Buildings, Edinburgh, EH9 3JL United Kingdom
| | - Jakub Nedbal
- Department of Physics, King’s College London, Strand, London, WC2R 2LS United Kingdom
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4
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Barber PR, Mustapha R, Flores-Borja F, Alfano G, Ng K, Weitsman G, Dolcetti L, Suwaidan AA, Wong F, Vicencio JM, Galazi M, Opzoomer JW, Arnold JN, Thavaraj S, Kordasti S, Doyle J, Greenberg J, Dillon MT, Harrington KJ, Forster M, Coolen ACC, Ng T. Predicting progression-free survival after systemic therapy in advanced head and neck cancer: Bayesian regression and model development. eLife 2022; 11:e73288. [PMID: 36562609 PMCID: PMC9815805 DOI: 10.7554/elife.73288] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/24/2021] [Accepted: 12/22/2022] [Indexed: 12/24/2022] Open
Abstract
Background Advanced head and neck squamous cell carcinoma (HNSCC) is associated with a poor prognosis, and biomarkers that predict response to treatment are highly desirable. The primary aim was to predict progression-free survival (PFS) with a multivariate risk prediction model. Methods Experimental covariates were derived from blood samples of 56 HNSCC patients which were prospectively obtained within a Phase 2 clinical trial (NCT02633800) at baseline and after the first treatment cycle of combined platinum-based chemotherapy with cetuximab treatment. Clinical and experimental covariates were selected by Bayesian multivariate regression to form risk scores to predict PFS. Results A 'baseline' and a 'combined' risk prediction model were generated, each of which featuring clinical and experimental covariates. The baseline risk signature has three covariates and was strongly driven by baseline percentage of CD33+CD14+HLADRhigh monocytes. The combined signature has six covariates, also featuring baseline CD33+CD14+HLADRhigh monocytes but is strongly driven by on-treatment relative change of CD8+ central memory T cells percentages. The combined model has a higher predictive power than the baseline model and was successfully validated to predict therapeutic response in an independent cohort of nine patients from an additional Phase 2 trial (NCT03494322) assessing the addition of avelumab to cetuximab treatment in HNSCC. We identified tissue counterparts for the immune cells driving the models, using imaging mass cytometry, that specifically colocalized at the tissue level and correlated with outcome. Conclusions This immune-based combined multimodality signature, obtained through longitudinal peripheral blood monitoring and validated in an independent cohort, presents a novel means of predicting response early on during the treatment course. Funding Daiichi Sankyo Inc, Cancer Research UK, EU IMI2 IMMUCAN, UK Medical Research Council, European Research Council (335326), Merck Serono. Cancer Research Institute, National Institute for Health Research, Guy's and St Thomas' NHS Foundation Trust and The Institute of Cancer Research. Clinical trial number NCT02633800.
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Affiliation(s)
- Paul R Barber
- UCL Cancer Institute, Paul O'Gorman Building, University College LondonLondonUnited Kingdom
- Comprehensive Cancer Centre, School of Cancer & Pharmaceutical Sciences, King’s College LondonLondonUnited Kingdom
| | - Rami Mustapha
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College LondonLondonUnited Kingdom
| | - Fabian Flores-Borja
- Breast Cancer Now Research Unit, School of Cancer & Pharmaceutical Sciences, King’s College LondonLondonUnited Kingdom
| | - Giovanna Alfano
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College LondonLondonUnited Kingdom
| | - Kenrick Ng
- UCL Cancer Institute, Paul O'Gorman Building, University College LondonLondonUnited Kingdom
| | - Gregory Weitsman
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College LondonLondonUnited Kingdom
| | - Luigi Dolcetti
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College LondonLondonUnited Kingdom
| | - Ali Abdulnabi Suwaidan
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College LondonLondonUnited Kingdom
| | - Felix Wong
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College LondonLondonUnited Kingdom
| | - Jose M Vicencio
- UCL Cancer Institute, Paul O'Gorman Building, University College LondonLondonUnited Kingdom
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College LondonLondonUnited Kingdom
| | - Myria Galazi
- UCL Cancer Institute, Paul O'Gorman Building, University College LondonLondonUnited Kingdom
| | - James W Opzoomer
- Tumor Immunology Group, School of Cancer & Pharmaceutical Sciences, King’s College LondonLondonUnited Kingdom
| | - James N Arnold
- Tumor Immunology Group, School of Cancer & Pharmaceutical Sciences, King’s College LondonLondonUnited Kingdom
| | - Selvam Thavaraj
- Centre for Clinical, Oral & Translational Science, King’s College LondonLondonUnited Kingdom
| | - Shahram Kordasti
- Systems Cancer Immunology, School of Cancer & Pharmaceutical Sciences, King’s College LondonLondonUnited Kingdom
| | - Jana Doyle
- Daiichi Sankyo IncorporatedNewarkUnited States
| | | | | | | | - Martin Forster
- UCL Cancer Institute, Paul O'Gorman Building, University College LondonLondonUnited Kingdom
| | - Anthony CC Coolen
- Institute for Mathematical and Molecular Biomedicine, King’s College LondonLondonUnited Kingdom
- Saddle Point Science LtdLondonUnited Kingdom
| | - Tony Ng
- UCL Cancer Institute, Paul O'Gorman Building, University College LondonLondonUnited Kingdom
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College LondonLondonUnited Kingdom
- Breast Cancer Now Research Unit, School of Cancer & Pharmaceutical Sciences, King’s College LondonLondonUnited Kingdom
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5
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Lawson CD, Peel S, Jayo A, Corrigan A, Iyer P, Baxter Dalrymple M, Marsh RJ, Cox S, Van Audenhove I, Gettemans J, Parsons M. Nuclear fascin regulates cancer cell survival. eLife 2022; 11:79283. [PMID: 36039640 PMCID: PMC9427113 DOI: 10.7554/elife.79283] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/06/2022] [Accepted: 08/04/2022] [Indexed: 11/13/2022] Open
Abstract
Fascin is an important regulator of F-actin bundling leading to enhanced filopodia assembly. Fascin is also overexpressed in most solid tumours where it supports invasion through control of F-actin structures at the periphery and nuclear envelope. Recently, fascin has been identified in the nucleus of a broad range of cell types but the contributions of nuclear fascin to cancer cell behaviour remain unknown. Here, we demonstrate that fascin bundles F-actin within the nucleus to support chromatin organisation and efficient DDR. Fascin associates directly with phosphorylated Histone H3 leading to regulated levels of nuclear fascin to support these phenotypes. Forcing nuclear fascin accumulation through the expression of nuclear-targeted fascin-specific nanobodies or inhibition of Histone H3 kinases results in enhanced and sustained nuclear F-actin bundling leading to reduced invasion, viability, and nuclear fascin-specific/driven apoptosis. These findings represent an additional important route through which fascin can support tumourigenesis and provide insight into potential pathways for targeted fascin-dependent cancer cell killing.
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Affiliation(s)
- Campbell D Lawson
- Randall Centre for Cell and Molecular Biophysics, King’s College London, Guy’s Campus
| | - Samantha Peel
- Discovery Sciences, R&D, AstraZeneca (United Kingdom)
| | - Asier Jayo
- Randall Centre for Cell and Molecular Biophysics, King’s College London, Guy’s Campus
| | - Adam Corrigan
- Discovery Sciences, R&D, AstraZeneca (United Kingdom)
| | - Preeti Iyer
- Molecular AI, Discovery Sciences, R&D, AstraZeneca (Sweden)
| | | | - Richard J Marsh
- Randall Centre for Cell and Molecular Biophysics, King’s College London, Guy’s Campus
| | - Susan Cox
- Randall Centre for Cell and Molecular Biophysics, King’s College London, Guy’s Campus
| | - Isabel Van Audenhove
- Department of Biomolecular Medicine, Faculty of Medicine and Health Sciences, Ghent University
| | - Jan Gettemans
- Department of Biomolecular Medicine, Faculty of Medicine and Health Sciences, Ghent University
| | - Maddy Parsons
- Randall Centre for Cell and Molecular Biophysics, King’s College London, Guy’s Campus
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6
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Laine RF, Poudel C, Kaminski CF. A method for the fast and photon-efficient analysis of time-domain fluorescence lifetime image data over large dynamic ranges. J Microsc 2022; 287:138-147. [PMID: 35676768 PMCID: PMC9544871 DOI: 10.1111/jmi.13128] [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: 09/02/2021] [Revised: 05/30/2022] [Accepted: 05/31/2022] [Indexed: 11/29/2022]
Abstract
Fluorescence lifetime imaging (FLIM) allows the quantification of sub‐cellular processes in situ, in living cells. A number of approaches have been developed to extract the lifetime from time‐domain FLIM data, but they are often limited in terms of speed, photon efficiency, precision or the dynamic range of lifetimes they can measure. Here, we focus on one of the best performing methods in the field, the centre‐of‐mass method (CMM), that conveys advantages in terms of speed and photon efficiency over others. In this paper, however, we identify a loss of photon efficiency of CMM for short lifetimes when background noise is present. We subsequently present a new development and generalization of CMM that provides for the rapid and accurate extraction of fluorescence lifetime over a large lifetime dynamic range. We provide software tools to simulate, validate and analyse FLIM data sets and compare the performance of our approach against the standard CMM and the commonly employed least‐square minimization (LSM) methods. Our method features a better photon efficiency than standard CMM and LSM and is robust in the presence of background noise. The algorithm is applicable to any time‐domain FLIM data set.
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Affiliation(s)
- Romain F Laine
- Laser Analytics Group, Department of Chemical Engineering and Biotechnology, University of Cambridge, Philippa Fawcett Drive, Cambridge, UK.,Medical Research Council Laboratory for Molecular Cell Biology (LMCB), University College London, Gower Street, London, WC1E 6BT
| | - Chetan Poudel
- Laser Analytics Group, Department of Chemical Engineering and Biotechnology, University of Cambridge, Philippa Fawcett Drive, Cambridge, UK.,Department of Chemistry, University of Washington, Seattle, WA 98195, USA
| | - Clemens F Kaminski
- Laser Analytics Group, Department of Chemical Engineering and Biotechnology, University of Cambridge, Philippa Fawcett Drive, Cambridge, UK
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7
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Targeting the HER3 pseudokinase domain with small molecule inhibitors. Methods Enzymol 2022; 667:455-505. [PMID: 35525551 DOI: 10.1016/bs.mie.2022.03.044] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/22/2022]
Abstract
HER3 is a potent oncogenic growth factor receptor belonging to the human epidermal growth factor (HER/EGFR) family of receptor tyrosine kinases. In contrast to other EGFR family members, HER3 is a pseudokinase, lacking functional kinase activity. As such, efforts to develop small molecule tyrosine kinase inhibitors against this family member have been limited. In response to HER3-specific growth factors such as neuregulin (NRG, also known as heregulin or HRG), HER3 must couple with catalytically active family members, including its preferred partner HER2. Dimerization of the intracellular HER2:HER3 kinase domains is a critical part of the activation mechanism and HER3 plays a specialized role as an allosteric activator of the active HER2 kinase partner. Intriguingly, many pseudokinases retain functionally important nucleotide binding capacity, despite loss of kinase activity. We demonstrated that occupation of the nucleotide pocket of the pseudokinase HER3 retains functional importance for growth factor signaling through oncogenic HER2:HER3 heterodimers. Mutation of the HER3 nucleotide pocket both disrupts signaling and disrupts HER2:HER3 dimerization. Conversely, ATP competitive drugs which bind to HER3, but not HER2, can stabilize HER2:HER3 dimers, induce signaling and promote cell growth in breast cancer models. This indicates a nucleotide-dependent conformational role for the HER3 kinase domain. Critically, our recent proof-of-concept work demonstrated that HER3-directed small molecule inhibitors can also disrupt HER2:HER3 dimerization and signaling, supporting the prospect that HER3 can be a direct drug target despite its lack of intrinsic activity. In this chapter we will describe methods for identifying and validating small molecule inhibitors against the HER3 pseudokinase.
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8
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Hertweck A, Vila de Mucha M, Barber PR, Dagil R, Porter H, Ramos A, Lord GM, Jenner RG. The TH1 cell lineage-determining transcription factor T-bet suppresses TH2 gene expression by redistributing GATA3 away from TH2 genes. Nucleic Acids Res 2022; 50:4557-4573. [PMID: 35438764 PMCID: PMC9071441 DOI: 10.1093/nar/gkac258] [Citation(s) in RCA: 14] [Impact Index Per Article: 7.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/09/2021] [Revised: 03/28/2022] [Accepted: 04/13/2022] [Indexed: 11/12/2022] Open
Abstract
Lineage-determining transcription factors (LD-TFs) drive the differentiation of progenitor cells into a specific lineage. In CD4+ T cells, T-bet dictates differentiation of the TH1 lineage, whereas GATA3 drives differentiation of the alternative TH2 lineage. However, LD-TFs, including T-bet and GATA3, are frequently co-expressed but how this affects LD-TF function is not known. By expressing T-bet and GATA3 separately or together in mouse T cells, we show that T-bet sequesters GATA3 at its target sites, thereby removing GATA3 from TH2 genes. This redistribution of GATA3 is independent of GATA3 DNA binding activity and is instead mediated by the T-bet DNA binding domain, which interacts with the GATA3 DNA binding domain and changes GATA3's sequence binding preference. This mechanism allows T-bet to drive the TH1 gene expression program in the presence of GATA3. We propose that redistribution of one LD-TF by another may be a common mechanism that could explain how specific cell fate choices can be made even in the presence of other transcription factors driving alternative differentiation pathways.
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Affiliation(s)
- Arnulf Hertweck
- UCL Cancer Institute and Cancer Research UK UCL Centre, University College London (UCL), London, WC1E 6BT, UK
| | - Maria Vila de Mucha
- UCL Cancer Institute and Cancer Research UK UCL Centre, University College London (UCL), London, WC1E 6BT, UK
| | - Paul R Barber
- UCL Cancer Institute and Cancer Research UK UCL Centre, University College London (UCL), London, WC1E 6BT, UK.,Comprehensive Cancer Centre, School of Cancer & Pharmaceutical Sciences, King's College London, London, SE1 1UL, UK
| | - Robert Dagil
- Research Department of Structural and Molecular Biology, University College London, Darwin Building, Gower Street, London, WC1E 6XA, UK
| | - Hayley Porter
- UCL Cancer Institute and Cancer Research UK UCL Centre, University College London (UCL), London, WC1E 6BT, UK
| | - Andres Ramos
- Research Department of Structural and Molecular Biology, University College London, Darwin Building, Gower Street, London, WC1E 6XA, UK
| | - Graham M Lord
- Faculty of Biology, Medicine and Health, University of Manchester, Manchester, M13 9NT, UK
| | - Richard G Jenner
- UCL Cancer Institute and Cancer Research UK UCL Centre, University College London (UCL), London, WC1E 6BT, UK
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9
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Vicencio JM, Evans R, Green R, An Z, Deng J, Treacy C, Mustapha R, Monypenny J, Costoya C, Lawler K, Ng K, De-Souza K, Coban O, Gomez V, Clancy J, Chen SH, Chalk A, Wong F, Gordon P, Savage C, Gomes C, Pan T, Alfano G, Dolcetti L, Chan JNE, Flores-Borja F, Barber PR, Weitsman G, Sosnowska D, Capone E, Iacobelli S, Hochhauser D, Hartley JA, Parsons M, Arnold JN, Ameer-Beg S, Quezada SA, Yarden Y, Sala G, Ng T. Osimertinib and anti-HER3 combination therapy engages immune dependent tumor toxicity via STING activation in trans. Cell Death Dis 2022; 13:274. [PMID: 35347108 PMCID: PMC8960767 DOI: 10.1038/s41419-022-04701-3] [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: 11/11/2021] [Revised: 02/07/2022] [Accepted: 03/01/2022] [Indexed: 11/28/2022]
Abstract
Over the past decade, immunotherapy delivered novel treatments for many cancer types. However, lung cancer still leads cancer mortality, and non-small-cell lung carcinoma patients with mutant EGFR cannot benefit from checkpoint inhibitors due to toxicity, relying only on palliative chemotherapy and the third-generation tyrosine kinase inhibitor (TKI) osimertinib. This new drug extends lifespan by 9-months vs. second-generation TKIs, but unfortunately, cancers relapse due to resistance mechanisms and the lack of antitumor immune responses. Here we explored the combination of osimertinib with anti-HER3 monoclonal antibodies and observed that the immune system contributed to eliminate tumor cells in mice and co-culture experiments using bone marrow-derived macrophages and human PBMCs. Osimertinib led to apoptosis of tumors but simultaneously, it triggered inositol-requiring-enzyme (IRE1α)-dependent HER3 upregulation, increased macrophage infiltration, and activated cGAS in cancer cells to produce cGAMP (detected by a lentivirally transduced STING activity biosensor), transactivating STING in macrophages. We sought to target osimertinib-induced HER3 upregulation with monoclonal antibodies, which engaged Fc receptor-dependent tumor elimination by macrophages, and STING agonists enhanced macrophage-mediated tumor elimination further. Thus, by engaging a tumor non-autonomous mechanism involving cGAS-STING and innate immunity, the combination of osimertinib and anti-HER3 antibodies could improve the limited therapeutic and stratification options for advanced stage lung cancer patients with mutant EGFR.
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Affiliation(s)
- J M Vicencio
- Molecular Oncology Group, Cancer Institute, Paul O'Gorman Building, University College London, London, UK.
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK.
| | - R Evans
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - R Green
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - Z An
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - J Deng
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - C Treacy
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - R Mustapha
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - J Monypenny
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - C Costoya
- Cancer Immunology Unit, Cancer Institute, University College London, London, UK
| | - K Lawler
- Wellcome Trust-MRC Institute of Metabolic Science, Addenbrooke's Hospital, Cambridge, UK
| | - K Ng
- Molecular Oncology Group, Cancer Institute, Paul O'Gorman Building, University College London, London, UK
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - K De-Souza
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - O Coban
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - V Gomez
- Molecular Oncology Group, Cancer Institute, Paul O'Gorman Building, University College London, London, UK
| | - J Clancy
- Molecular Oncology Group, Cancer Institute, Paul O'Gorman Building, University College London, London, UK
| | - S H Chen
- Molecular Oncology Group, Cancer Institute, Paul O'Gorman Building, University College London, London, UK
| | - A Chalk
- Molecular Oncology Group, Cancer Institute, Paul O'Gorman Building, University College London, London, UK
| | - F Wong
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - P Gordon
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - C Savage
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - C Gomes
- Molecular Oncology Group, Cancer Institute, Paul O'Gorman Building, University College London, London, UK
| | - T Pan
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - G Alfano
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - L Dolcetti
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - J N E Chan
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - F Flores-Borja
- Centre for Immunobiology and Regenerative Medicine, Barts & The London School of Medicine and Dentistry, Queen Mary University of London, London, UK
| | - P R Barber
- Molecular Oncology Group, Cancer Institute, Paul O'Gorman Building, University College London, London, UK
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - G Weitsman
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - D Sosnowska
- School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - E Capone
- Department of Innovative Technologies in Medicine & Dentistry, University of Chieti-Pescara, Center for Advanced Studies and Technology (CAST), Chieti, Italy
| | | | - D Hochhauser
- Molecular Oncology Group, Cancer Institute, Paul O'Gorman Building, University College London, London, UK
| | - J A Hartley
- Molecular Oncology Group, Cancer Institute, Paul O'Gorman Building, University College London, London, UK
| | - M Parsons
- Randall Centre for Cell and Molecular Biophysics, King's College London, London, UK
| | - J N Arnold
- School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - S Ameer-Beg
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK
| | - S A Quezada
- Cancer Immunology Unit, Cancer Institute, University College London, London, UK
| | - Y Yarden
- Department of Biological Regulation, The Weizmann Institute of Science, Rehovot, Israel
| | - G Sala
- Department of Innovative Technologies in Medicine & Dentistry, University of Chieti-Pescara, Center for Advanced Studies and Technology (CAST), Chieti, Italy
| | - T Ng
- Molecular Oncology Group, Cancer Institute, Paul O'Gorman Building, University College London, London, UK.
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King's College London, London, UK.
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10
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Héliot L, Leray A. Simple phasor-based deep neural network for fluorescence lifetime imaging microscopy. Sci Rep 2021; 11:23858. [PMID: 34903737 PMCID: PMC8668934 DOI: 10.1038/s41598-021-03060-x] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/28/2021] [Accepted: 11/23/2021] [Indexed: 12/29/2022] Open
Abstract
Fluorescence lifetime imaging microscopy (FLIM) is a powerful technique to probe the molecular environment of fluorophores. The analysis of FLIM images is usually performed with time consuming fitting methods. For accelerating this analysis, sophisticated deep learning architectures based on convolutional neural networks have been developed for restrained lifetime ranges but they require long training time. In this work, we present a simple neural network formed only with fully connected layers able to analyze fluorescence lifetime images. It is based on the reduction of high dimensional fluorescence intensity temporal decays into four parameters which are the phasor coordinates, the mean and amplitude-weighted lifetimes. This network called Phasor-Net has been applied for a time domain FLIM system excited with an 80 MHz laser repetition frequency, with negligible jitter and afterpulsing. Due to the restricted time interval of 12.5 ns, the training range of the lifetimes was limited between 0.2 and 3.0 ns; and the total photon number was lower than 106, as encountered in live cell imaging. From simulated biexponential decays, we demonstrate that Phasor-Net is more precise and less biased than standard fitting methods. We demonstrate also that this simple architecture gives almost comparable performance than those obtained from more sophisticated networks but with a faster training process (15 min instead of 30 min). We finally apply successfully our method to determine biexponential decays parameters for FLIM experiments in living cells expressing EGFP linked to mCherry and fused to a plasma membrane protein.
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Affiliation(s)
- Laurent Héliot
- PhLAM Laboratoire de Physique Des Lasers, Atomes Et Molécules, UMR 8523, CNRS, University of Lille, Lille, France.
| | - Aymeric Leray
- Laboratoire Interdisciplinaire Carnot de Bourgogne, UMR 6303, CNRS, Université de Bourgogne Franche-Comté, Dijon, France.
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11
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Law AL, Jalal S, Pallett T, Mosis F, Guni A, Brayford S, Yolland L, Marcotti S, Levitt JA, Poland SP, Rowe-Sampson M, Jandke A, Köchl R, Pula G, Ameer-Beg SM, Stramer BM, Krause M. Nance-Horan Syndrome-like 1 protein negatively regulates Scar/WAVE-Arp2/3 activity and inhibits lamellipodia stability and cell migration. Nat Commun 2021; 12:5687. [PMID: 34584076 PMCID: PMC8478917 DOI: 10.1038/s41467-021-25916-6] [Citation(s) in RCA: 10] [Impact Index Per Article: 3.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/24/2018] [Accepted: 09/03/2021] [Indexed: 12/02/2022] Open
Abstract
Cell migration is important for development and its aberrant regulation contributes to many diseases. The Scar/WAVE complex is essential for Arp2/3 mediated lamellipodia formation during mesenchymal cell migration and several coinciding signals activate it. However, so far, no direct negative regulators are known. Here we identify Nance-Horan Syndrome-like 1 protein (NHSL1) as a direct binding partner of the Scar/WAVE complex, which co-localise at protruding lamellipodia. This interaction is mediated by the Abi SH3 domain and two binding sites in NHSL1. Furthermore, active Rac binds to NHSL1 at two regions that mediate leading edge targeting of NHSL1. Surprisingly, NHSL1 inhibits cell migration through its interaction with the Scar/WAVE complex. Mechanistically, NHSL1 may reduce cell migration efficiency by impeding Arp2/3 activity, as measured in cells using a Arp2/3 FRET-FLIM biosensor, resulting in reduced F-actin density of lamellipodia, and consequently impairing the stability of lamellipodia protrusions.
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Affiliation(s)
- Ah-Lai Law
- Krause Group, Randall Centre for Cell and Molecular Biophysics, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
- School of Life Sciences, University of Bedfordshire, Luton, LU1 3JU, UK
| | - Shamsinar Jalal
- Krause Group, Randall Centre for Cell and Molecular Biophysics, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
| | - Tommy Pallett
- Krause Group, Randall Centre for Cell and Molecular Biophysics, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
| | - Fuad Mosis
- Krause Group, Randall Centre for Cell and Molecular Biophysics, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
| | - Ahmad Guni
- Krause Group, Randall Centre for Cell and Molecular Biophysics, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
| | - Simon Brayford
- Stramer Group, Randall Centre for Cell and Molecular Biophysics, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
| | - Lawrence Yolland
- Stramer Group, Randall Centre for Cell and Molecular Biophysics, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
| | - Stefania Marcotti
- Stramer Group, Randall Centre for Cell and Molecular Biophysics, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
| | - James A Levitt
- Ameer-Beg Group, Richard Dimbleby Cancer Research Laboratories, Comprehensive Cancer Centre, School of Cancer and Pharmaceutical Sciences, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
| | - Simon P Poland
- Ameer-Beg Group, Richard Dimbleby Cancer Research Laboratories, Comprehensive Cancer Centre, School of Cancer and Pharmaceutical Sciences, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
| | - Maia Rowe-Sampson
- Krause Group, Randall Centre for Cell and Molecular Biophysics, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
- Stramer Group, Randall Centre for Cell and Molecular Biophysics, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
| | - Anett Jandke
- Krause Group, Randall Centre for Cell and Molecular Biophysics, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
- Immunosurveillance Laboratory, The Francis Crick Institute, 1 Midland Road, London, NW1 1AT, UK
| | - Robert Köchl
- School of Immunology and Microbial Sciences, King's College London, Guy's Campus, London, SE1 1UL, UK
| | - Giordano Pula
- Krause Group, Randall Centre for Cell and Molecular Biophysics, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
- Institute for Clinical Chemistry and Laboratory Medicine, University Medical Center Hamburg (UKE), Martinistrasse 52, O26, 20246, Hamburg, Germany
| | - Simon M Ameer-Beg
- Ameer-Beg Group, Richard Dimbleby Cancer Research Laboratories, Comprehensive Cancer Centre, School of Cancer and Pharmaceutical Sciences, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
| | - Brian Marc Stramer
- Stramer Group, Randall Centre for Cell and Molecular Biophysics, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK
| | - Matthias Krause
- Krause Group, Randall Centre for Cell and Molecular Biophysics, King's College London, New Hunt's House, Guy's Campus, London, SE1 1UL, UK.
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12
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Wang P, Hecht F, Ossato G, Tille S, Fraser SE, Junge JA. Complex wavelet filter improves FLIM phasors for photon starved imaging experiments. BIOMEDICAL OPTICS EXPRESS 2021; 12:3463-3473. [PMID: 34221672 PMCID: PMC8221945 DOI: 10.1364/boe.420953] [Citation(s) in RCA: 12] [Impact Index Per Article: 4.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Received: 01/28/2021] [Revised: 03/22/2021] [Accepted: 03/30/2021] [Indexed: 05/11/2023]
Abstract
Fluorescence lifetime imaging microscopy (FLIM) with phasor analysis provides easy visualization and analysis of fluorophores' lifetimes which is valuable for multiple applications including metabolic imaging, STED imaging, FRET imaging and functional imaging. However, FLIM imaging typically suffers from low photon budgets, leading to unfavorable signal to noise ratios which in many cases prevent extraction of information from the data. Traditionally, median filters are applied in phasor analysis to tackle this problem. This unfortunately degrades high spatial frequency FLIM information in the phasor analysis. These high spatial frequency components are typically edges of features and puncta, which applies to membranes, mitochondria, granules and small organelles in a biological sample. To tackle this problem, we propose a filtering strategy with complex wavelet filtering and Anscombe transform for FLIM phasor analysis. This filtering strategy preserves fine structures and reports accurate lifetimes in photon starved FLIM imaging. Moreover, this filter outperforms median filters and makes FLIM imaging with lower laser power and faster imaging possible.
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Affiliation(s)
- P. Wang
- Translational Imaging Center, Dornsife School of Letters, Arts, and Sciences, University of Southern California, 1002 Childs Way, Los Angeles, CA 90089, USA
| | - F. Hecht
- Leica Microsystems CMS GmbH, Am Friedensplatz 3, Mannheim 68165, Germany
| | - G. Ossato
- Leica Microsystems CMS GmbH, Am Friedensplatz 3, Mannheim 68165, Germany
| | - S. Tille
- Leica Microsystems CMS GmbH, Am Friedensplatz 3, Mannheim 68165, Germany
| | - S. E. Fraser
- Translational Imaging Center, Dornsife School of Letters, Arts, and Sciences, University of Southern California, 1002 Childs Way, Los Angeles, CA 90089, USA
| | - J. A. Junge
- Translational Imaging Center, Dornsife School of Letters, Arts, and Sciences, University of Southern California, 1002 Childs Way, Los Angeles, CA 90089, USA
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13
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Gao D, Barber PR, Chacko JV, Kader Sagar MA, Rueden CT, Grislis AR, Hiner MC, Eliceiri KW. FLIMJ: An open-source ImageJ toolkit for fluorescence lifetime image data analysis. PLoS One 2020; 15:e0238327. [PMID: 33378370 PMCID: PMC7773231 DOI: 10.1371/journal.pone.0238327] [Citation(s) in RCA: 14] [Impact Index Per Article: 3.5] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/11/2020] [Accepted: 12/14/2020] [Indexed: 12/11/2022] Open
Abstract
In the field of fluorescence microscopy, there is continued demand for dynamic technologies that can exploit the complete information from every pixel of an image. One imaging technique with proven ability for yielding additional information from fluorescence imaging is Fluorescence Lifetime Imaging Microscopy (FLIM). FLIM allows for the measurement of how long a fluorophore stays in an excited energy state, and this measurement is affected by changes in its chemical microenvironment, such as proximity to other fluorophores, pH, and hydrophobic regions. This ability to provide information about the microenvironment has made FLIM a powerful tool for cellular imaging studies ranging from metabolic measurement to measuring distances between proteins. The increased use of FLIM has necessitated the development of computational tools for integrating FLIM analysis with image and data processing. To address this need, we have created FLIMJ, an ImageJ plugin and toolkit that allows for easy use and development of extensible image analysis workflows with FLIM data. Built on the FLIMLib decay curve fitting library and the ImageJ Ops framework, FLIMJ offers FLIM fitting routines with seamless integration with many other ImageJ components, and the ability to be extended to create complex FLIM analysis workflows. Building on ImageJ Ops also enables FLIMJ's routines to be used with Jupyter notebooks and integrate naturally with science-friendly programming in, e.g., Python and Groovy. We show the extensibility of FLIMJ in two analysis scenarios: lifetime-based image segmentation and image colocalization. We also validate the fitting routines by comparing them against industry FLIM analysis standards.
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Affiliation(s)
- Dasong Gao
- Laboratory for Optical and Computational Instrumentation, Center for Quantitative Cell Imaging, University of Wisconsin, Madison, WI, United States of America
| | - Paul R. Barber
- UCL Cancer Institute, Paul O’Gorman Building, University College London, London, United Kingdom
| | - Jenu V. Chacko
- Laboratory for Optical and Computational Instrumentation, Center for Quantitative Cell Imaging, University of Wisconsin, Madison, WI, United States of America
| | - Md. Abdul Kader Sagar
- Laboratory for Optical and Computational Instrumentation, Center for Quantitative Cell Imaging, University of Wisconsin, Madison, WI, United States of America
- Department of Biomedical Engineering, University of Wisconsin, Madison, WI, United States of America
| | - Curtis T. Rueden
- Laboratory for Optical and Computational Instrumentation, Center for Quantitative Cell Imaging, University of Wisconsin, Madison, WI, United States of America
| | - Aivar R. Grislis
- Laboratory for Optical and Computational Instrumentation, Center for Quantitative Cell Imaging, University of Wisconsin, Madison, WI, United States of America
| | - Mark C. Hiner
- Laboratory for Optical and Computational Instrumentation, Center for Quantitative Cell Imaging, University of Wisconsin, Madison, WI, United States of America
| | - Kevin W. Eliceiri
- Laboratory for Optical and Computational Instrumentation, Center for Quantitative Cell Imaging, University of Wisconsin, Madison, WI, United States of America
- Department of Biomedical Engineering, University of Wisconsin, Madison, WI, United States of America
- Department of Medical Physics, University of Wisconsin, Madison, WI, United States of America
- Morgridge Institute for Research, University of Wisconsin, Madison, WI, United States of America
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14
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Sen R, Zhdanov AV, Bastiaanssen TFS, Hirvonen LM, Svihra P, Fitzgerald P, Cryan JF, Andersson-Engels S, Nomerotski A, Papkovsky DB. Mapping O 2 concentration in ex-vivo tissue samples on a fast PLIM macro-imager. Sci Rep 2020; 10:19006. [PMID: 33149165 PMCID: PMC7642408 DOI: 10.1038/s41598-020-75928-3] [Citation(s) in RCA: 7] [Impact Index Per Article: 1.8] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/01/2020] [Accepted: 10/08/2020] [Indexed: 12/27/2022] Open
Abstract
O2 PLIM microscopy was employed in various studies, however current platforms have limitations in sensitivity, image acquisition speed, accuracy and general usability. We describe a new PLIM imager based on the Timepix3 camera (Tpx3cam) and its application for imaging of O2 concentration in various tissue samples stained with a nanoparticle based probe, NanO2-IR. Upon passive staining of mouse brain, lung or intestinal tissue surface with minute quantities of NanO2-IR or by microinjecting the probe into the lumen of small or large intestine fragments, robust phosphorescence intensity and lifetime signals were produced, which allow mapping of O2 in the tissue within 20 s. Inhibition of tissue respiration or limitation of O2 diffusion to tissue produced the anticipated increases or decreases in O2 levels, respectively. The difference in O2 concentration between the colonic lumen and air-exposed serosal surface was around 140 µM. Furthermore, subcutaneous injection of 5 µg of the probe in intact organs (a paw or tail of sacrificed mice) enabled efficient O2 imaging at tissue depths of up to 0.5 mm. Overall, the PLIM imager holds promise for metabolic imaging studies with various ex vivo models of animal tissue, and also for use in live animals.
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Affiliation(s)
- Rajannya Sen
- School of Biochemistry and Cell Biology, University College Cork, Cork, Ireland
| | - Alexander V Zhdanov
- School of Biochemistry and Cell Biology, University College Cork, Cork, Ireland
| | - Thomaz F S Bastiaanssen
- APC Microbiome Ireland, University College Cork, Cork, Ireland
- Department of Anatomy and Neuroscience, University College Cork, Cork, Ireland
| | - Liisa M Hirvonen
- Centre for Microscopy, Characterisation and Analysis (CMCA), The University of Western Australia, Crawley, WA, 6009, Australia
| | - Peter Svihra
- Department of Physics, Faculty of Nuclear Sciences and Physical Engineering, Czech Technical University, 115 19, Prague, Czech Republic
- Department of Physics and Astronomy, School of Natural Sciences, The University of Manchester, Manchester, M139PL, UK
| | | | - John F Cryan
- APC Microbiome Ireland, University College Cork, Cork, Ireland
- Department of Anatomy and Neuroscience, University College Cork, Cork, Ireland
| | | | - Andrei Nomerotski
- Physics Department, Brookhaven National Laboratory, Upton, NY, 11973, USA
| | - Dmitri B Papkovsky
- School of Biochemistry and Cell Biology, University College Cork, Cork, Ireland.
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15
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Barber PR, Weitsman G, Lawler K, Barrett JE, Rowley M, Rodriguez-Justo M, Fisher D, Gao F, Tullis IDC, Deng J, Brown L, Kaplan R, Hochhauser D, Adams R, Maughan TS, Vojnovic B, Coolen ACC, Ng T. HER2-HER3 Heterodimer Quantification by FRET-FLIM and Patient Subclass Analysis of the COIN Colorectal Trial. J Natl Cancer Inst 2020; 112:944-954. [PMID: 31851321 PMCID: PMC7492762 DOI: 10.1093/jnci/djz231] [Citation(s) in RCA: 9] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/29/2019] [Revised: 11/27/2019] [Accepted: 12/11/2019] [Indexed: 12/20/2022] Open
Abstract
BACKGROUND The phase III MRC COIN trial showed no statistically significant benefit from adding the EGFR-target cetuximab to oxaliplatin-based chemotherapy in first-line treatment of advanced colorectal cancer. This study exploits additional information on HER2-HER3 dimerization to achieve patient stratification and reveal previously hidden subgroups of patients who had differing disease progression and treatment response. METHODS HER2-HER3 dimerization was quantified by fluorescence lifetime imaging microscopy in primary tumor samples from 550 COIN trial patients receiving oxaliplatin and fluoropyrimidine chemotherapy with or without cetuximab. Bayesian latent class analysis and covariate reduction was performed to analyze the effects of HER2-HER3 dimer, RAS mutation, and cetuximab on progression-free survival and overall survival (OS). All statistical tests were two-sided. RESULTS Latent class analysis on a cohort of 398 patients revealed two patient subclasses with differing prognoses (median OS = 1624 days [95% confidence interval [CI] = 1466 to 1816 days] vs 461 days [95% CI = 431 to 504 days]): Class 1 (15.6%) showed a benefit from cetuximab in OS (hazard ratio = 0.43, 95% CI = 0.25 to 0.76, P = .004). Class 2 showed an association of increased HER2-HER3 with better OS (hazard ratio = 0.64, 95% CI = 0.44 to 0.94, P = .02). A class prediction signature was formed and tested on an independent validation cohort (n = 152) validating the prognostic utility of the dimer assay. Similar subclasses were also discovered in full trial dataset (n = 1630) based on 10 baseline clinicopathological and genetic covariates. CONCLUSIONS Our work suggests that the combined use of HER dimer imaging and conventional mutation analyses will be able to identify a small subclass of patients (>10%) who will have better prognosis following chemotherapy. A larger prospective cohort will be required to confirm its utility in predicting the outcome of anti-EGFR treatment.
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Affiliation(s)
- Paul R Barber
- UCL Cancer Institute, Paul O’Gorman Building, University College London, London, UK
| | - Gregory Weitsman
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King’s College London, London, UK
| | - Katherine Lawler
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King’s College London, London, UK
- Institute for Mathematical and Molecular Biomedicine, King’s College London, Guy’s Medical School Campus, London, UK
| | - James E Barrett
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King’s College London, London, UK
| | - Mark Rowley
- Institute for Mathematical and Molecular Biomedicine, King’s College London, Guy’s Medical School Campus, London, UK
- Saddle Point Science Ltd, London, UK
| | | | - David Fisher
- MRC Clinical Trials Unit at UCL, Institute of Clinical Trials & Methodology, London, UK
| | - Fangfei Gao
- UCL Cancer Institute, Paul O’Gorman Building, University College London, London, UK
| | - Iain D C Tullis
- Department of Oncology, Cancer Research UK and Medical Research Council Oxford Institute for Radiation Oncology, University of Oxford, Oxford, UK
| | - Jinhai Deng
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King’s College London, London, UK
| | - Louise Brown
- MRC Clinical Trials Unit at UCL, Institute of Clinical Trials & Methodology, London, UK
| | - Richard Kaplan
- MRC Clinical Trials Unit at UCL, Institute of Clinical Trials & Methodology, London, UK
| | - Daniel Hochhauser
- UCL Cancer Institute, Paul O’Gorman Building, University College London, London, UK
| | | | - Timothy S. Maughan
- Department of Oncology, Cancer Research UK and Medical Research Council Oxford Institute for Radiation Oncology, University of Oxford, Oxford, UK
| | - Borivoj Vojnovic
- Department of Oncology, Cancer Research UK and Medical Research Council Oxford Institute for Radiation Oncology, University of Oxford, Oxford, UK
| | - Anthony C C Coolen
- Institute for Mathematical and Molecular Biomedicine, King’s College London, Guy’s Medical School Campus, London, UK
- Saddle Point Science Ltd, London, UK
| | - Tony Ng
- UCL Cancer Institute, Paul O’Gorman Building, University College London, London, UK
- Richard Dimbleby Laboratory of Cancer Research, School of Cancer & Pharmaceutical Sciences, King’s College London, London, UK
- Breast Cancer Now Research Unit, Department of Research Oncology, Guy’s Hospital King’s College London, London, UK
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16
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Pfisterer K, Levitt J, Lawson CD, Marsh RJ, Heddleston JM, Wait E, Ameer-Beg SM, Cox S, Parsons M. FMNL2 regulates dynamics of fascin in filopodia. J Cell Biol 2020; 219:e201906111. [PMID: 32294157 PMCID: PMC7199847 DOI: 10.1083/jcb.201906111] [Citation(s) in RCA: 21] [Impact Index Per Article: 5.3] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/19/2019] [Revised: 11/30/2019] [Accepted: 02/20/2020] [Indexed: 12/31/2022] Open
Abstract
Filopodia are peripheral F-actin-rich structures that enable cell sensing of the microenvironment. Fascin is an F-actin-bundling protein that plays a key role in stabilizing filopodia to support efficient adhesion and migration. Fascin is also highly up-regulated in human cancers, where it increases invasive cell behavior and correlates with poor patient prognosis. Previous studies have shown that fascin phosphorylation can regulate F-actin bundling, and that this modification can contribute to subcellular fascin localization and function. However, the factors that regulate fascin dynamics within filopodia remain poorly understood. In the current study, we used advanced live-cell imaging techniques and a fascin biosensor to demonstrate that fascin phosphorylation, localization, and binding to F-actin are highly dynamic and dependent on local cytoskeletal architecture in cells in both 2D and 3D environments. Fascin dynamics within filopodia are under the control of formins, and in particular FMNL2, that binds directly to dephosphorylated fascin. Our data provide new insight into control of fascin dynamics at the nanoscale and into the mechanisms governing rapid cytoskeletal adaptation to environmental changes. This filopodia-driven exploration stage may represent an essential regulatory step in the transition from static to migrating cancer cells.
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Affiliation(s)
- Karin Pfisterer
- Randall Centre for Cell and Molecular Biophysics, King's College London, Guy's Campus, London, UK
| | - James Levitt
- Randall Centre for Cell and Molecular Biophysics, King's College London, Guy's Campus, London, UK
- Microscopy Innovation Centre, King's College London, Guy's Campus, London, UK
| | - Campbell D. Lawson
- Randall Centre for Cell and Molecular Biophysics, King's College London, Guy's Campus, London, UK
| | - Richard J. Marsh
- Randall Centre for Cell and Molecular Biophysics, King's College London, Guy's Campus, London, UK
| | - John M. Heddleston
- Advanced Imaging Centre, Howard Hughes Medical Institute, Janelia Research Campus, Ashburn, VA
| | - Eric Wait
- Advanced Imaging Centre, Howard Hughes Medical Institute, Janelia Research Campus, Ashburn, VA
| | - Simon Morris Ameer-Beg
- Randall Centre for Cell and Molecular Biophysics, King's College London, Guy's Campus, London, UK
- School of Cancer and Pharmaceutical Sciences, King's College London, Guy's Campus, London, UK
| | - Susan Cox
- Randall Centre for Cell and Molecular Biophysics, King's College London, Guy's Campus, London, UK
| | - Maddy Parsons
- Randall Centre for Cell and Molecular Biophysics, King's College London, Guy's Campus, London, UK
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17
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Levitt JA, Poland SP, Krstajic N, Pfisterer K, Erdogan A, Barber PR, Parsons M, Henderson RK, Ameer-Beg SM. Quantitative real-time imaging of intracellular FRET biosensor dynamics using rapid multi-beam confocal FLIM. Sci Rep 2020; 10:5146. [PMID: 32198437 PMCID: PMC7083966 DOI: 10.1038/s41598-020-61478-1] [Citation(s) in RCA: 16] [Impact Index Per Article: 4.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/11/2019] [Accepted: 02/14/2020] [Indexed: 01/21/2023] Open
Abstract
Fluorescence lifetime imaging (FLIM) is a quantitative, intensity-independent microscopical method for measurement of diverse biochemical and physical properties in cell biology. It is a highly effective method for measurements of Förster resonance energy transfer (FRET), and for quantification of protein-protein interactions in cells. Time-domain FLIM-FRET measurements of these dynamic interactions are particularly challenging, since the technique requires excellent photon statistics to derive experimental parameters from the complex decay kinetics often observed from fluorophores in living cells. Here we present a new time-domain multi-confocal FLIM instrument with an array of 64 visible beamlets to achieve parallelised excitation and detection with average excitation powers of ~ 1–2 μW per beamlet. We exemplify this instrument with up to 0.5 frames per second time-lapse FLIM measurements of cAMP levels using an Epac-based fluorescent biosensor in live HeLa cells with nanometer spatial and picosecond temporal resolution. We demonstrate the use of time-dependent phasor plots to determine parameterisation for multi-exponential decay fitting to monitor the fractional contribution of the activated conformation of the biosensor. Our parallelised confocal approach avoids having to compromise on speed, noise, accuracy in lifetime measurements and provides powerful means to quantify biochemical dynamics in living cells.
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Affiliation(s)
- James A Levitt
- Microscopy Innovation Centre, Guy's Campus, Kings College, London, SE1 1UL, UK.,Richard Dimbleby Laboratories, School of Cancer and Pharmaceutical Sciences, Guy's Campus, Kings College London, London, SE1 1UL, UK
| | - Simon P Poland
- Richard Dimbleby Laboratories, School of Cancer and Pharmaceutical Sciences, Guy's Campus, Kings College London, London, SE1 1UL, UK
| | - Nikola Krstajic
- Institute for Microelectronics and Nanosystems, School of Engineering, College of Science and Engineering, University of Edinburgh, Edinburgh, EH9 3FB, UK
| | - Karin Pfisterer
- Randall Centre for Cell and Molecular Biophysics, Guy's Campus, Kings College, London, SE1 1UL, UK
| | - Ahmet Erdogan
- Institute for Microelectronics and Nanosystems, School of Engineering, College of Science and Engineering, University of Edinburgh, Edinburgh, EH9 3FB, UK
| | - Paul R Barber
- UCL Cancer Institute, Paul O'Gorman Building, University College London, London, WC1E 6DD, UK
| | - Maddy Parsons
- Randall Centre for Cell and Molecular Biophysics, Guy's Campus, Kings College, London, SE1 1UL, UK
| | - Robert K Henderson
- Institute for Microelectronics and Nanosystems, School of Engineering, College of Science and Engineering, University of Edinburgh, Edinburgh, EH9 3FB, UK
| | - Simon M Ameer-Beg
- Richard Dimbleby Laboratories, School of Cancer and Pharmaceutical Sciences, Guy's Campus, Kings College London, London, SE1 1UL, UK.
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18
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Sen R, Hirvonen LM, Zhdanov A, Svihra P, Andersson-Engels S, Nomerotski A, Papkovsky D. New luminescence lifetime macro-imager based on a Tpx3Cam optical camera. BIOMEDICAL OPTICS EXPRESS 2020; 11:77-88. [PMID: 32010501 PMCID: PMC6968763 DOI: 10.1364/boe.11.000077] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Received: 08/26/2019] [Revised: 10/16/2019] [Accepted: 10/16/2019] [Indexed: 05/10/2023]
Abstract
The properties of a novel ultra-fast optical imager, Tpx3Cam, were investigated for macroscopic wide-field phosphorescent lifetime imaging (PLIM) applications. The camera is based on a novel optical sensor and Timepix3 readout chip with a time resolution of 1.6 ns, recording of photon arrival time and time over threshold for each pixel, and readout rate of 80 megapixels per second. In this study, we coupled the camera to an image intensifier, a 760 nm emission filter and a 50 mm lens, and with a super-bright 627nm LED providing pulsed excitation of a 18 × 18 mm sample area. The resulting macro-imager with compact and rigid optical alignment of its main components was characterised using planar phosphorescent O2 sensors and a resolution plate mask. Several acquisition and image processing algorithms were evaluated to optimise the system resolution and performance for the wide-field PLIM, followed by imaging a variety of phosphorescent samples. The new PLIM system looks promising, particularly for phosphorescence lifetime-based imaging of O2 in various chemical and biological samples.
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Affiliation(s)
- Rajannya Sen
- School of Biochemistry and Cell Biology, University College Cork, Cork, Ireland
- These authors contributed equally
| | - Liisa M. Hirvonen
- Centre for Microscopy, Characterisation and Analysis (CMCA), The University of Western Australia, Crawley WA 6009, Australia
- These authors contributed equally
| | - Alexander Zhdanov
- School of Biochemistry and Cell Biology, University College Cork, Cork, Ireland
| | - Peter Svihra
- Department of Physics, Faculty of Nuclear Sciences and Physical Engineering, Czech Technical University, Prague 115 19, Czech Republic
- Department of Physics and Astronomy, School of Natural Sciences, The University of Manchester, Manchester M139PL, United Kingdom
| | | | - Andrei Nomerotski
- Physics Department, Brookhaven National Laboratory, Upton, New York 11973, USA
| | - Dmitri Papkovsky
- School of Biochemistry and Cell Biology, University College Cork, Cork, Ireland
- Irish Photonics Integration Centre, Tyndall National Institute, Cork, Ireland
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19
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Akula AK, Zhang X, Viotti JS, Nestvogel D, Rhee JS, Ebrecht R, Reim K, Wouters F, Liepold T, Jahn O, Bogeski I, Dresbach T. The Calmodulin Binding Region of the Synaptic Vesicle Protein Mover Is Required for Homomeric Interaction and Presynaptic Targeting. Front Mol Neurosci 2019; 12:249. [PMID: 31787876 PMCID: PMC6856015 DOI: 10.3389/fnmol.2019.00249] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/13/2019] [Accepted: 09/26/2019] [Indexed: 01/10/2023] Open
Abstract
Neurotransmitter release is mediated by an evolutionarily conserved machinery. The synaptic vesicle (SV) associated protein Mover/TPRGL/SVAP30 does not occur in all species and all synapses. Little is known about its molecular properties and how it may interact with the conserved components of the presynaptic machinery. Here, we show by deletion analysis that regions required for homomeric interaction of Mover are distributed across the entire molecule, including N-terminal, central and C-terminal regions. The same regions are also required for the accumulation of Mover in presynaptic terminals of cultured neurons. Mutating two phosphorylation sites in N-terminal regions did not affect these properties. In contrast, a point mutation in the predicted Calmodulin (CaM) binding sequence of Mover abolished both homomeric interaction and presynaptic targeting. We show that this sequence indeed binds Calmodulin, and that recombinant Mover increases Calmodulin signaling upon heterologous expression. Our data suggest that presynaptic accumulation of Mover requires homomeric interaction mediated by regions distributed across large areas of the protein, and corroborate the hypothesis that Mover functionally interacts with Calmodulin signaling.
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Affiliation(s)
- Asha Kiran Akula
- Institute for Anatomy and Embryology, University Medical Center Göttingen, Göttingen, Germany
| | - Xin Zhang
- Molecular Physiology, Institute of Cardiovascular Physiology, University Medical Center, Göttingen, Germany
| | - Julio S Viotti
- Institute for Anatomy and Embryology, University Medical Center Göttingen, Göttingen, Germany
| | - Dennis Nestvogel
- Department of Molecular Neurobiology, Max Planck Institute of Experimental Medicine, Göttingen, Germany
| | - Jeong-Seop Rhee
- Department of Molecular Neurobiology, Max Planck Institute of Experimental Medicine, Göttingen, Germany
| | - Rene Ebrecht
- Institute for Neuropathology, University Medical Center Göttingen, Göttingen, Germany
| | - Kerstin Reim
- Department of Molecular Neurobiology, Max Planck Institute of Experimental Medicine, Göttingen, Germany
| | - Fred Wouters
- Institute for Neuropathology, University Medical Center Göttingen, Göttingen, Germany
| | - Thomas Liepold
- Proteomics Group, Max Planck Institute of Experimental Medicine, Göttingen, Germany
| | - Olaf Jahn
- Proteomics Group, Max Planck Institute of Experimental Medicine, Göttingen, Germany
| | - Ivan Bogeski
- Molecular Physiology, Institute of Cardiovascular Physiology, University Medical Center, Göttingen, Germany
| | - Thomas Dresbach
- Institute for Anatomy and Embryology, University Medical Center Göttingen, Göttingen, Germany
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20
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Evans R, Flores-Borja F, Nassiri S, Miranda E, Lawler K, Grigoriadis A, Monypenny J, Gillet C, Owen J, Gordon P, Male V, Cheung A, Noor F, Barber P, Marlow R, Francesch-Domenech E, Fruhwirth G, Squadrito M, Vojnovic B, Tutt A, Festy F, De Palma M, Ng T. Integrin-Mediated Macrophage Adhesion Promotes Lymphovascular Dissemination in Breast Cancer. Cell Rep 2019; 27:1967-1978.e4. [PMID: 31091437 PMCID: PMC6527923 DOI: 10.1016/j.celrep.2019.04.076] [Citation(s) in RCA: 38] [Impact Index Per Article: 7.6] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/12/2018] [Revised: 03/14/2019] [Accepted: 04/17/2019] [Indexed: 12/31/2022] Open
Abstract
Lymphatic vasculature is crucial for metastasis in triple-negative breast cancer (TNBC); however, cellular and molecular drivers controlling lymphovascular metastasis are poorly understood. We define a macrophage-dependent signaling cascade that facilitates metastasis through lymphovascular remodeling. TNBC cells instigate mRNA changes in macrophages, resulting in β4 integrin-dependent adhesion to the lymphovasculature. β4 integrin retains macrophages proximal to lymphatic endothelial cells (LECs), where release of TGF-β1 drives LEC contraction via RhoA activation. Macrophages promote gross architectural changes to lymphovasculature by increasing dilation, hyperpermeability, and disorganization. TGF-β1 drives β4 integrin clustering at the macrophage plasma membrane, further promoting macrophage adhesion and demonstrating the dual functionality of TGF-β1 signaling in this context. β4 integrin-expressing macrophages were identified in human breast tumors, and a combination of vascular-remodeling macrophage gene signature and TGF-β signaling scores correlates with metastasis. We postulate that future clinical strategies for patients with TNBC should target crosstalk between β4 integrin and TGF-β1.
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Affiliation(s)
- Rachel Evans
- Richard Dimbleby Department of Cancer Research, Randall Division & Division of Cancer Studies, Kings College London, London, UK.
| | - Fabian Flores-Borja
- Breast Cancer Now Research Unit, King's College London, Guy's Hospital, London, UK
| | - Sina Nassiri
- Swiss Institute for Experimental Cancer Research (ISREC), School of Life Sciences, École Polytechnique Fédérale de Lausanne (EPFL), Lausanne, Switzerland
| | - Elena Miranda
- Pathology Core Facility, University College London Cancer Institute, London, UK
| | - Katherine Lawler
- Richard Dimbleby Department of Cancer Research, Randall Division & Division of Cancer Studies, Kings College London, London, UK; Institute for Mathematical and Molecular Biomedicine, King's College London, London, UK
| | - Anita Grigoriadis
- Breast Cancer Now Research Unit, King's College London, Guy's Hospital, London, UK
| | - James Monypenny
- Richard Dimbleby Department of Cancer Research, Randall Division & Division of Cancer Studies, Kings College London, London, UK
| | - Cheryl Gillet
- King's Health Partners Cancer Biobank, King's College London, London, UK; Research Oncology, Division of Cancer Studies, Guy's Hospital, King's College London, London, UK
| | - Julie Owen
- King's Health Partners Cancer Biobank, King's College London, London, UK; Research Oncology, Division of Cancer Studies, Guy's Hospital, King's College London, London, UK
| | - Peter Gordon
- Breast Cancer Now Research Unit, King's College London, Guy's Hospital, London, UK
| | - Victoria Male
- Breast Cancer Now Research Unit, King's College London, Guy's Hospital, London, UK
| | - Anthony Cheung
- Breast Cancer Now Research Unit, King's College London, Guy's Hospital, London, UK
| | - Farzana Noor
- Breast Cancer Now Research Unit, King's College London, Guy's Hospital, London, UK
| | - Paul Barber
- Richard Dimbleby Department of Cancer Research, Randall Division & Division of Cancer Studies, Kings College London, London, UK; UCL Cancer Institute, University College London, London, UK
| | - Rebecca Marlow
- Breast Cancer Now Research Unit, King's College London, Guy's Hospital, London, UK
| | | | - Gilbert Fruhwirth
- Division of Imaging Sciences and Biomedical Engineering, King's College London, London, UK
| | - Mario Squadrito
- Swiss Institute for Experimental Cancer Research (ISREC), School of Life Sciences, École Polytechnique Fédérale de Lausanne (EPFL), Lausanne, Switzerland
| | - Borivoj Vojnovic
- Department of Oncology, Cancer Research UK and Medical Research Council, Oxford Institute for Radiation Oncology, University of Oxford, UK
| | - Andrew Tutt
- Breast Cancer Now Research Unit, King's College London, Guy's Hospital, London, UK
| | - Frederic Festy
- Tissue Engineering and Biophotonics, King's College London, London, UK
| | - Michele De Palma
- Swiss Institute for Experimental Cancer Research (ISREC), School of Life Sciences, École Polytechnique Fédérale de Lausanne (EPFL), Lausanne, Switzerland
| | - Tony Ng
- Richard Dimbleby Department of Cancer Research, Randall Division & Division of Cancer Studies, Kings College London, London, UK; Breast Cancer Now Research Unit, King's College London, Guy's Hospital, London, UK; UCL Cancer Institute, University College London, London, UK.
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21
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Aluko J, Perrin C, Devauges V, Nedbal J, Poland S, Matthews D, Whittaker J, Ameer-Beg S. Semi-autonomous real-time programmable fluorescence lifetime segmentation with a digital micromirror device. OPTICS EXPRESS 2018; 26:31055-31074. [PMID: 30650697 DOI: 10.1364/oe.26.031055] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 07/11/2018] [Accepted: 08/22/2018] [Indexed: 06/09/2023]
Abstract
Time-correlated single-photon counting (TCSPC) is the gold standard for performing lifetime spectroscopy in biological assays. Traditional fluorescence lifetime imaging (FLIM) using laser scanning microscopes are inherently slow due to point scanning all pixels in the field-of-view. Wide-field implementations of TCSPC spectroscopy using microchannel plates benefit from particularly fast acquisition times at the expense of temporal resolution, and are fundamentally limited by photon counting rates. Here, we introduce programmable lifetime imaging (PLI), combining the advantages of wide-field imaging using total internal reflection excitation with state-of-the-art TCSPC detector technology for accurate lifetime determination in an object-oriented manner using a digital micromirror device (DMD). The fluorescent emission is projected onto the DMD to facilitate the sequential segmentation of fluorescence from individual objects in the field-of-view, allowing for both image acquisition and fluorescence lifetime determination of the assay. The sensitivity of PLI is demonstrated by manually segmenting fluorescence from fixed cell assays. We also demonstrate an automated implementation of PLI, using a camera as a feedback mechanism to segment fluorescence produced by emitting objects of interest in the imaging field-of-view, highlighting the advantages of measurement only in areas where valuable information exists. As a result, PLI is able to reduce acquisition time of fluorescence lifetime data by at least an order of magnitude compared to laser scanning implementations.
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22
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Claus J, Patel G, Autore F, Colomba A, Weitsman G, Soliman TN, Roberts S, Zanetti-Domingues LC, Hirsch M, Collu F, George R, Ortiz-Zapater E, Barber PR, Vojnovic B, Yarden Y, Martin-Fernandez ML, Cameron A, Fraternali F, Ng T, Parker PJ. Inhibitor-induced HER2-HER3 heterodimerisation promotes proliferation through a novel dimer interface. eLife 2018; 7:e32271. [PMID: 29712619 PMCID: PMC5929906 DOI: 10.7554/elife.32271] [Citation(s) in RCA: 48] [Impact Index Per Article: 8.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/25/2017] [Accepted: 03/21/2018] [Indexed: 12/13/2022] Open
Abstract
While targeted therapy against HER2 is an effective first-line treatment in HER2+ breast cancer, acquired resistance remains a clinical challenge. The pseudokinase HER3, heterodimerisation partner of HER2, is widely implicated in the resistance to HER2-mediated therapy. Here, we show that lapatinib, an ATP-competitive inhibitor of HER2, is able to induce proliferation cooperatively with the HER3 ligand neuregulin. This counterintuitive synergy between inhibitor and growth factor depends on their ability to promote atypical HER2-HER3 heterodimerisation. By stabilising a particular HER2 conformer, lapatinib drives HER2-HER3 kinase domain heterocomplex formation. This dimer exists in a head-to-head orientation distinct from the canonical asymmetric active dimer. The associated clustering observed for these dimers predisposes to neuregulin responses, affording a proliferative outcome. Our findings provide mechanistic insights into the liabilities involved in targeting kinases with ATP-competitive inhibitors and highlight the complex role of protein conformation in acquired resistance.
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Affiliation(s)
- Jeroen Claus
- Protein Phosphorylation LaboratoryThe Francis Crick InstituteLondonUnited Kingdom
| | - Gargi Patel
- Richard Dimbleby Department of Cancer Research, Randall Division and Division of Cancer StudiesKings College LondonLondonUnited Kingdom
- Sussex Cancer CentreBrighton and Sussex University HospitalsBrightonUnited States
| | - Flavia Autore
- Randall Division of Cell & Molecular BiophysicsKings College LondonLondonUnited Kingdom
| | - Audrey Colomba
- Protein Phosphorylation LaboratoryThe Francis Crick InstituteLondonUnited Kingdom
| | - Gregory Weitsman
- Richard Dimbleby Department of Cancer Research, Randall Division and Division of Cancer StudiesKings College LondonLondonUnited Kingdom
| | - Tanya N Soliman
- Protein Phosphorylation LaboratoryThe Francis Crick InstituteLondonUnited Kingdom
| | - Selene Roberts
- Central Laser Facility, Research Complex at Harwell, Science and Technology Facilities CouncilRutherford Appleton LaboratoryDidcotUnited Kingdom
| | - Laura C Zanetti-Domingues
- Central Laser Facility, Research Complex at Harwell, Science and Technology Facilities CouncilRutherford Appleton LaboratoryDidcotUnited Kingdom
| | - Michael Hirsch
- Central Laser Facility, Research Complex at Harwell, Science and Technology Facilities CouncilRutherford Appleton LaboratoryDidcotUnited Kingdom
| | - Francesca Collu
- Randall Division of Cell & Molecular BiophysicsKings College LondonLondonUnited Kingdom
| | - Roger George
- The Structural Biology Science Technology PlatformThe Francis Crick InstituteLondonUnited Kingdom
| | - Elena Ortiz-Zapater
- Department of Asthma, Allergy and Respiratory ScienceKing’s College London, Guy’s HospitalLondonUnited Kingdom
| | - Paul R Barber
- Randall Division of Cell & Molecular BiophysicsKings College LondonLondonUnited Kingdom
- UCL Cancer InstituteUniversity College LondonLondonUnited Kingdom
| | - Boris Vojnovic
- Randall Division of Cell & Molecular BiophysicsKings College LondonLondonUnited Kingdom
- Department of OncologyCancer Research UK and Medical Research Council Oxford Institute for Radiation OncologyOxfordUnited Kingdom
| | - Yosef Yarden
- Department of Biological RegulationWeizmann Institute of ScienceRehovotIsrael
| | - Marisa L Martin-Fernandez
- Central Laser Facility, Research Complex at Harwell, Science and Technology Facilities CouncilRutherford Appleton LaboratoryDidcotUnited Kingdom
| | - Angus Cameron
- Protein Phosphorylation LaboratoryThe Francis Crick InstituteLondonUnited Kingdom
- Barts Cancer InstituteQueen Mary University of LondonLondonUnited Kingdom
| | - Franca Fraternali
- Randall Division of Cell & Molecular BiophysicsKings College LondonLondonUnited Kingdom
| | - Tony Ng
- Richard Dimbleby Department of Cancer Research, Randall Division and Division of Cancer StudiesKings College LondonLondonUnited Kingdom
- UCL Cancer InstituteUniversity College LondonLondonUnited Kingdom
- Breast Cancer Now Research Unit, Department of Research OncologyGuy’s Hospital King’s College London School of MedicineLondonUnited Kingdom
| | - Peter J Parker
- Protein Phosphorylation LaboratoryThe Francis Crick InstituteLondonUnited Kingdom
- School of Cancer and Pharmaceutical SciencesKing’s College London, Guy’s CampusLondonUnited Kingdom
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23
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Le Marois A, Suhling K. Quantitative Live Cell FLIM Imaging in Three Dimensions. ADVANCES IN EXPERIMENTAL MEDICINE AND BIOLOGY 2018; 1035:31-48. [PMID: 29080129 DOI: 10.1007/978-3-319-67358-5_3] [Citation(s) in RCA: 10] [Impact Index Per Article: 1.7] [Reference Citation Analysis] [Abstract] [Key Words] [Subscribe] [Scholar Register] [Indexed: 01/16/2023]
Abstract
In this chapter, the concept of fluorescence lifetime and its utility in quantitative live cell imaging will be introduced, along with methods to record and analyze FLIM data. Relevant applications in 3D tissue and live cell imaging, including multiplexed FLIM detection, will also be detailed.
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Affiliation(s)
- Alix Le Marois
- Department of Physics, King's College London, Strand, London, WC2R 2LS, UK
| | - Klaus Suhling
- Department of Physics, King's College London, Strand, London, WC2R 2LS, UK.
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24
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Fili N, Hari-Gupta Y, Dos Santos Á, Cook A, Poland S, Ameer-Beg SM, Parsons M, Toseland CP. NDP52 activates nuclear myosin VI to enhance RNA polymerase II transcription. Nat Commun 2017; 8:1871. [PMID: 29187741 PMCID: PMC5707354 DOI: 10.1038/s41467-017-02050-w] [Citation(s) in RCA: 39] [Impact Index Per Article: 5.6] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/10/2017] [Accepted: 10/30/2017] [Indexed: 11/09/2022] Open
Abstract
Myosin VI (MVI) has been found to be overexpressed in ovarian, breast and prostate cancers. Moreover, it has been shown to play a role in regulating cell proliferation and migration, and to interact with RNA Polymerase II (RNAPII). Here, we find that backfolding of MVI regulates its ability to bind DNA and that a putative transcription co-activator NDP52 relieves the auto-inhibition of MVI to enable DNA binding. Additionally, we show that the MVI-NDP52 complex binds RNAPII, which is critical for transcription, and that depletion of NDP52 or MVI reduces steady-state mRNA levels. Lastly, we demonstrate that MVI directly interacts with nuclear receptors to drive expression of target genes, thereby suggesting a link to cell proliferation and migration. Overall, we suggest MVI may function as an auxiliary motor to drive transcription.
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Affiliation(s)
- Natalia Fili
- School of Biosciences, University of Kent, Canterbury, CT2 7NJ, UK
| | - Yukti Hari-Gupta
- School of Biosciences, University of Kent, Canterbury, CT2 7NJ, UK
| | - Ália Dos Santos
- School of Biosciences, University of Kent, Canterbury, CT2 7NJ, UK
| | - Alexander Cook
- School of Biosciences, University of Kent, Canterbury, CT2 7NJ, UK
| | - Simon Poland
- Randall Division of Cell and Molecular Biophysics, King's College London, Guys Campus, London, SE1 1UL, UK
| | - Simon M Ameer-Beg
- Randall Division of Cell and Molecular Biophysics, King's College London, Guys Campus, London, SE1 1UL, UK
| | - Maddy Parsons
- Randall Division of Cell and Molecular Biophysics, King's College London, Guys Campus, London, SE1 1UL, UK
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25
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Weitsman G, Mitchell NJ, Evans R, Cheung A, Kalber TL, Bofinger R, Fruhwirth GO, Keppler M, Wright ZVF, Barber PR, Gordon P, de Koning T, Wulaningsih W, Sander K, Vojnovic B, Ameer-Beg S, Lythgoe M, Arnold JN, Årstad E, Festy F, Hailes HC, Tabor AB, Ng T. Detecting intratumoral heterogeneity of EGFR activity by liposome-based in vivo transfection of a fluorescent biosensor. Oncogene 2017; 36:3618-3628. [PMID: 28166195 PMCID: PMC5421598 DOI: 10.1038/onc.2016.522] [Citation(s) in RCA: 11] [Impact Index Per Article: 1.6] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/13/2016] [Revised: 11/12/2016] [Accepted: 12/21/2016] [Indexed: 12/20/2022]
Abstract
Despite decades of research in the epidermal growth factor receptor (EGFR) signalling field, and many targeted anti-cancer drugs that have been tested clinically, the success rate for these agents in the clinic is low, particularly in terms of the improvement of overall survival. Intratumoral heterogeneity is proposed as a major mechanism underlying treatment failure of these molecule-targeted agents. Here we highlight the application of fluorescence lifetime microscopy (FLIM)-based biosensing to demonstrate intratumoral heterogeneity of EGFR activity. For sensing EGFR activity in cells, we used a genetically encoded CrkII-based biosensor which undergoes conformational changes upon tyrosine-221 phosphorylation by EGFR. We transfected this biosensor into EGFR-positive tumour cells using targeted lipopolyplexes bearing EGFR-binding peptides at their surfaces. In a murine model of basal-like breast cancer, we demonstrated a significant degree of intratumoral heterogeneity in EGFR activity, as well as the pharmacodynamic effect of a radionuclide-labeled EGFR inhibitor in situ. Furthermore, a significant correlation between high EGFR activity in tumour cells and macrophage-tumour cell proximity was found to in part account for the intratumoral heterogeneity in EGFR activity observed. The same effect of macrophage infiltrate on EGFR activation was also seen in a colorectal cancer xenograft. In contrast, a non-small cell lung cancer xenograft expressing a constitutively active EGFR conformational mutant exhibited macrophage proximity-independent EGFR activity. Our study validates the use of this methodology to monitor therapeutic response in terms of EGFR activity. In addition, we found iNOS gene induction in macrophages that are cultured in tumour cell-conditioned media as well as an iNOS activity-dependent increase in EGFR activity in tumour cells. These findings point towards an immune microenvironment-mediated regulation that gives rise to the observed intratumoral heterogeneity of EGFR signalling activity in tumour cells in vivo.
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Affiliation(s)
- G Weitsman
- Richard Dimbleby Department of Cancer Research, Randall Division & Division of Cancer Studies, Kings College London, Guy’s Medical School Campus, London, UK
| | - N J Mitchell
- Department of Chemistry, University College London, London, UK
| | - R Evans
- Richard Dimbleby Department of Cancer Research, Randall Division & Division of Cancer Studies, Kings College London, Guy’s Medical School Campus, London, UK
| | - A Cheung
- Richard Dimbleby Department of Cancer Research, Randall Division & Division of Cancer Studies, Kings College London, Guy’s Medical School Campus, London, UK
- Breast Cancer Now Research Unit, King’s College London, London, UK
| | - T L Kalber
- UCL Centre for Advanced Biomedical Imaging, Division of Medicine, University College London, London, UK
| | - R Bofinger
- Department of Chemistry, University College London, London, UK
| | - G O Fruhwirth
- Richard Dimbleby Department of Cancer Research, Randall Division & Division of Cancer Studies, Kings College London, Guy’s Medical School Campus, London, UK
| | - M Keppler
- Richard Dimbleby Department of Cancer Research, Randall Division & Division of Cancer Studies, Kings College London, Guy’s Medical School Campus, London, UK
| | - Z V F Wright
- Department of Chemistry, University College London, London, UK
| | - P R Barber
- Gray Laboratories, Department of Oncology, Cancer Research UK and Medical Research Council Oxford Institute for Radiation Oncology, Oxford, UK
| | - P Gordon
- Breast Cancer Now Research Unit, King’s College London, London, UK
| | - T de Koning
- Division of Cancer Studies, Kings College London, Guy’s Medical School Campus, London, UK
| | - W Wulaningsih
- Cancer Epidemiology Group, Division of Cancer Studies, King’s College London, London, UK
| | - K Sander
- Institute of Nuclear Medicine, University College London, London, UK
| | - B Vojnovic
- Gray Laboratories, Department of Oncology, Cancer Research UK and Medical Research Council Oxford Institute for Radiation Oncology, Oxford, UK
| | - S Ameer-Beg
- Richard Dimbleby Department of Cancer Research, Randall Division & Division of Cancer Studies, Kings College London, Guy’s Medical School Campus, London, UK
| | - M Lythgoe
- UCL Centre for Advanced Biomedical Imaging, Division of Medicine, University College London, London, UK
| | - J N Arnold
- Division of Cancer Studies, Kings College London, Guy’s Medical School Campus, London, UK
| | - E Årstad
- Institute of Nuclear Medicine, University College London, London, UK
| | - F Festy
- King’s College London Dental Institute, Tissue Engineering and Biophotonics, Guy’s Hospital Campus, London, UK
| | - H C Hailes
- Department of Chemistry, University College London, London, UK
| | - A B Tabor
- Department of Chemistry, University College London, London, UK
| | - T Ng
- Richard Dimbleby Department of Cancer Research, Randall Division & Division of Cancer Studies, Kings College London, Guy’s Medical School Campus, London, UK
- Breast Cancer Now Research Unit, King’s College London, London, UK
- UCL Cancer Institute, Paul O’Gorman Building, University College London, London, UK
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26
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Verboogen DRJ, González Mancha N, Ter Beest M, van den Bogaart G. Fluorescence Lifetime Imaging Microscopy reveals rerouting of SNARE trafficking driving dendritic cell activation. eLife 2017; 6. [PMID: 28524818 PMCID: PMC5473687 DOI: 10.7554/elife.23525] [Citation(s) in RCA: 17] [Impact Index Per Article: 2.4] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/22/2016] [Accepted: 05/18/2017] [Indexed: 11/13/2022] Open
Abstract
SNARE proteins play a crucial role in intracellular trafficking by catalyzing membrane fusion, but assigning SNAREs to specific intracellular transport routes is challenging with current techniques. We developed a novel Förster resonance energy transfer-fluorescence lifetime imaging microscopy (FRET-FLIM)-based technique allowing visualization of real-time local interactions of fluorescently tagged SNARE proteins in live cells. We used FRET-FLIM to delineate the trafficking steps underlying the release of the inflammatory cytokine interleukin-6 (IL-6) from human blood-derived dendritic cells. We found that activation of dendritic cells by bacterial lipopolysaccharide leads to increased FRET of fluorescently labeled syntaxin 4 with VAMP3 specifically at the plasma membrane, indicating increased SNARE complex formation, whereas FRET with other tested SNAREs was unaltered. Our results revealed that SNARE complexing is a key regulatory step for cytokine production by immune cells and prove the applicability of FRET-FLIM for visualizing SNARE complexes in live cells with subcellular spatial resolution.
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Affiliation(s)
- Daniëlle Rianne José Verboogen
- Department of Tumor Immunology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, Netherlands
| | - Natalia González Mancha
- Department of Tumor Immunology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, Netherlands
| | - Martin Ter Beest
- Department of Tumor Immunology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, Netherlands
| | - Geert van den Bogaart
- Department of Tumor Immunology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, Netherlands
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27
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Conway JRW, Warren SC, Timpson P. Context-dependent intravital imaging of therapeutic response using intramolecular FRET biosensors. Methods 2017; 128:78-94. [PMID: 28435000 DOI: 10.1016/j.ymeth.2017.04.014] [Citation(s) in RCA: 33] [Impact Index Per Article: 4.7] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/31/2017] [Revised: 03/13/2017] [Accepted: 04/08/2017] [Indexed: 12/18/2022] Open
Abstract
Intravital microscopy represents a more physiologically relevant method for assessing therapeutic response. However, the movement into an in vivo setting brings with it several additional considerations, the primary being the context in which drug activity is assessed. Microenvironmental factors, such as hypoxia, pH, fibrosis, immune infiltration and stromal interactions have all been shown to have pronounced effects on drug activity in a more complex setting, which is often lost in simpler two- or three-dimensional assays. Here we present a practical guide for the application of intravital microscopy, looking at the available fluorescent reporters and their respective expression systems and analysis considerations. Moving in vivo, we also discuss the microscopy set up and methods available for overlaying microenvironmental context to the experimental readouts. This enables a smooth transition into applying higher fidelity intravital imaging to improve the drug discovery process.
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Affiliation(s)
- James R W Conway
- Garvan Institute of Medical Research and The Kinghorn Cancer Centre, Cancer Division, Sydney, NSW 2010, Australia; St Vincent's Clinical School, Faculty of Medicine, University of NSW, Sydney, NSW 2010, Australia
| | - Sean C Warren
- Garvan Institute of Medical Research and The Kinghorn Cancer Centre, Cancer Division, Sydney, NSW 2010, Australia; St Vincent's Clinical School, Faculty of Medicine, University of NSW, Sydney, NSW 2010, Australia
| | - Paul Timpson
- Garvan Institute of Medical Research and The Kinghorn Cancer Centre, Cancer Division, Sydney, NSW 2010, Australia; St Vincent's Clinical School, Faculty of Medicine, University of NSW, Sydney, NSW 2010, Australia.
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Hirvonen LM, Fisher-Levine M, Suhling K, Nomerotski A. Photon counting phosphorescence lifetime imaging with TimepixCam. THE REVIEW OF SCIENTIFIC INSTRUMENTS 2017; 88:013104. [PMID: 28147700 DOI: 10.1063/1.4973717] [Citation(s) in RCA: 7] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 05/25/2023]
Abstract
TimepixCam is a novel fast optical imager based on an optimized silicon pixel sensor with a thin entrance window and read out by a Timepix Application Specific Integrated Circuit. The 256 × 256 pixel sensor has a time resolution of 15 ns at a sustained frame rate of 10 Hz. We used this sensor in combination with an image intensifier for wide-field time-correlated single photon counting imaging. We have characterised the photon detection capabilities of this detector system and employed it on a wide-field epifluorescence microscope to map phosphorescence decays of various iridium complexes with lifetimes of about 1 μs in 200 μm diameter polystyrene beads.
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Affiliation(s)
- Liisa M Hirvonen
- Department of Physics, King's College London, Strand, London WC2R 2LS, United Kingdom
| | | | - Klaus Suhling
- Department of Physics, King's College London, Strand, London WC2R 2LS, United Kingdom
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29
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Adrien V, Rayan G, Reffay M, Porcar L, Maldonado A, Ducruix A, Urbach W, Taulier N. Characterization of a Biomimetic Mesophase Composed of Nonionic Surfactants and an Aqueous Solvent. LANGMUIR : THE ACS JOURNAL OF SURFACES AND COLLOIDS 2016; 32:10268-10275. [PMID: 27618561 DOI: 10.1021/acs.langmuir.6b02744] [Citation(s) in RCA: 4] [Impact Index Per Article: 0.5] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/06/2023]
Abstract
We have investigated the physical and biomimetic properties of a sponge (L3) phase composed of pentaethylene glycol monododecyl ether (C12E5), a nonionic surfactant, an aqueous solvent, and a cosurfactant. The following cosurfactants, commonly used for solubilizing membrane proteins, were incorporated: n-octyl-β-d-glucopyranoside (β-OG), n-dodecyl-β-d-maltopyranoside (DDM), 4-cyclohexyl-1-butyl-β-d-maltoside (CYMAL-4), and 5-cyclohexyl-1-pentyl-β-d-maltoside (CYMAL-5). Partial phase diagrams of these systems were created. The L3 phase was characterized using crossed polarizers, diffusion of a fluorescent probe by fluorescence recovery after pattern photobleaching (FRAPP), and freeze fracture electron microscopy (FFEM). By varying the hydration of the phase, we were able to tune the distance between adjacent bilayers. The characteristic distance (db) of the phase was obtained from small angle scattering (SAXS/SANS) as well as from FFEM, which yielded complementary db values. These db values were neither affected by the nature of the cosurfactant nor by the addition of membrane proteins. These findings illustrate that a biomimetic surfactant sponge phase can be created in the presence of several common membrane protein-solubilizing detergents, thus making it a versatile medium for membrane protein studies.
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Affiliation(s)
- V Adrien
- Laboratoire de Physique Statistique, Ecole Normale Supérieure, PSL Research University; Université Paris Diderot Sorbonne Paris Cité; Sorbonne Universités UPMC Univ Paris 06, CNRS, 24 rue Lhomond, 75005 Paris, France
- Univ Paris Descartes, Sorbonne Paris Cité. Laboratoire de Cristallographie et RMN Biologiques, CNRS UMR 8015, Paris, France
| | - G Rayan
- Laboratoire de Physique Statistique, Ecole Normale Supérieure, PSL Research University; Université Paris Diderot Sorbonne Paris Cité; Sorbonne Universités UPMC Univ Paris 06, CNRS, 24 rue Lhomond, 75005 Paris, France
| | - M Reffay
- Laboratoire de Physique Statistique, Ecole Normale Supérieure, PSL Research University; Université Paris Diderot Sorbonne Paris Cité; Sorbonne Universités UPMC Univ Paris 06, CNRS, 24 rue Lhomond, 75005 Paris, France
| | - L Porcar
- Institut Laue-Langevin, 71 avenue des Martyrs, CS 20156, 38042 Grenoble Cedex 9, France
| | - A Maldonado
- Departamento de Física, Universidad de Sonora , Apdo Postal 1626, 83000 Hermosillo, Sonora Mexico
| | - A Ducruix
- Univ Paris Descartes, Sorbonne Paris Cité. Laboratoire de Cristallographie et RMN Biologiques, CNRS UMR 8015, Paris, France
| | - W Urbach
- Laboratoire de Physique Statistique, Ecole Normale Supérieure, PSL Research University; Université Paris Diderot Sorbonne Paris Cité; Sorbonne Universités UPMC Univ Paris 06, CNRS, 24 rue Lhomond, 75005 Paris, France
- Sorbonne Universités, UPMC Univ Paris 06, CNRS, INSERM, Laboratoire d'Imagerie Biomédicale, F-75006, Paris, France
| | - N Taulier
- Sorbonne Universités, UPMC Univ Paris 06, CNRS, INSERM, Laboratoire d'Imagerie Biomédicale, F-75006, Paris, France
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30
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Rowley MI, Coolen ACC, Vojnovic B, Barber PR. Robust Bayesian Fluorescence Lifetime Estimation, Decay Model Selection and Instrument Response Determination for Low-Intensity FLIM Imaging. PLoS One 2016; 11:e0158404. [PMID: 27355322 PMCID: PMC4927071 DOI: 10.1371/journal.pone.0158404] [Citation(s) in RCA: 25] [Impact Index Per Article: 3.1] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/18/2016] [Accepted: 06/15/2016] [Indexed: 11/26/2022] Open
Abstract
We present novel Bayesian methods for the analysis of exponential decay data that exploit the evidence carried by every detected decay event and enables robust extension to advanced processing. Our algorithms are presented in the context of fluorescence lifetime imaging microscopy (FLIM) and particular attention has been paid to model the time-domain system (based on time-correlated single photon counting) with unprecedented accuracy. We present estimates of decay parameters for mono- and bi-exponential systems, offering up to a factor of two improvement in accuracy compared to previous popular techniques. Results of the analysis of synthetic and experimental data are presented, and areas where the superior precision of our techniques can be exploited in Förster Resonance Energy Transfer (FRET) experiments are described. Furthermore, we demonstrate two advanced processing methods: decay model selection to choose between differing models such as mono- and bi-exponential, and the simultaneous estimation of instrument and decay parameters.
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Affiliation(s)
- Mark I. Rowley
- Institute for Mathematical and Molecular Biomedicine, King’s College London, London, United Kingdom
| | - Anthonius C. C. Coolen
- Institute for Mathematical and Molecular Biomedicine, King’s College London, London, United Kingdom
| | - Borivoj Vojnovic
- Cancer Research UK and Medical Research Council Oxford Institute for Radiation Oncology, Department of Oncology, University of Oxford, Oxford, United Kingdom
| | - Paul R. Barber
- Institute for Mathematical and Molecular Biomedicine, King’s College London, London, United Kingdom
- Cancer Research UK and Medical Research Council Oxford Institute for Radiation Oncology, Department of Oncology, University of Oxford, Oxford, United Kingdom
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31
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Loison P, Gervais P, Perrier-Cornet JM, Kuimova MK. Effect of ethanol perturbation on viscosity and permeability of an inner membrane in Bacillus subtilis spores. BIOCHIMICA ET BIOPHYSICA ACTA-BIOMEMBRANES 2016; 1858:2060-2069. [PMID: 27267704 DOI: 10.1016/j.bbamem.2016.06.003] [Citation(s) in RCA: 15] [Impact Index Per Article: 1.9] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Subscribe] [Scholar Register] [Received: 09/21/2015] [Revised: 05/30/2016] [Accepted: 06/02/2016] [Indexed: 02/02/2023]
Abstract
In this work, we investigated how a combination of ethanol and high temperature (70°C), affect the properties of the inner membrane of Bacillus subtilis spores. We observed membrane permeabilization for ethanol concentrations ≥50%, as indicated by the staining of the spores' DNA by the cell impermeable dye Propidium Iodide. The loss of membrane integrity was also confirmed by a decrease in the peak corresponding to dipicolinic acid using infrared spectroscopy. Finally, the spore refractivity (as measured by phase contrast microscopy) was decreased after the ethanol-heat treatment, suggesting a partial rehydration of the protoplast. Previously we have used fluorescent lifetime imaging microscopy (FLIM) combined with the fluorescent molecular rotor Bodipy-C12 to study the microscopic viscosity in the inner membrane of B. subtilis spores, and showed that at normal conditions it is characterized by a very high viscosity. Here we demonstrate that the ethanol/high temperature treatment led to a decrease of the viscosity of the inner membrane, from 1000cP to 860cP for wild type spores at 50% of ethanol. Altogether, our present work confirms the deleterious effect of ethanol on the structure of B. subtilis spores, as well as demonstrates the ability of FLIM - Bodipy-C12 to measure changes in the microviscosity of the spores upon perturbation.
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Affiliation(s)
- Pauline Loison
- Univ. Bourgogne Franche-Comté, AgroSup Dijon, PAM UMR A 02.102, PMB, F-21000 Dijon, France
| | - Patrick Gervais
- Univ. Bourgogne Franche-Comté, AgroSup Dijon, PAM UMR A 02.102, PMB, F-21000 Dijon, France
| | - Jean-Marie Perrier-Cornet
- Univ. Bourgogne Franche-Comté, AgroSup Dijon, PAM UMR A 02.102, PMB, F-21000 Dijon, France; Dimacell Imaging Resource Center, Univ. Bourgogne Franche-Comté, AgroSup Dijon, F-21000 Dijon, France.
| | - Marina K Kuimova
- Department of Chemistry, Imperial College London, Exhibition Road, SW7 2AZ, UK.
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32
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Poland SP, Erdogan AT, Krstajić N, Levitt J, Devauges V, Walker RJ, Li DDU, Ameer-Beg SM, Henderson RK. New high-speed centre of mass method incorporating background subtraction for accurate determination of fluorescence lifetime. OPTICS EXPRESS 2016; 24:6899-915. [PMID: 27136986 DOI: 10.1364/oe.24.006899] [Citation(s) in RCA: 17] [Impact Index Per Article: 2.1] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 05/19/2023]
Abstract
We demonstrate an implementation of a centre-of-mass method (CMM) incorporating background subtraction for use in multifocal fluorescence lifetime imaging microscopy to accurately determine fluorescence lifetime in live cell imaging using the Megaframe camera. The inclusion of background subtraction solves one of the major issues associated with centre-of-mass approaches, namely the sensitivity of the algorithm to background signal. The algorithm, which is predominantly implemented in hardware, provides real-time lifetime output and allows the user to effectively condense large amounts of photon data. Instead of requiring the transfer of thousands of photon arrival times, the lifetime is simply represented by one value which allows the system to collect data up to limit of pulse pile-up without any limitations on data transfer rates. In order to evaluate the performance of this new CMM algorithm with existing techniques (i.e. rapid lifetime determination and Levenburg-Marquardt), we imaged live MCF-7 human breast carcinoma cells transiently transfected with FRET standards. We show that, it offers significant advantages in terms of lifetime accuracy and insensitivity to variability in dark count rate (DCR) between Megaframe camera pixels. Unlike other algorithms no prior knowledge of the expected lifetime is required to perform lifetime determination. The ability of this technique to provide real-time lifetime readout makes it extremely useful for a number of applications.
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33
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Spectral variation of fluorescence lifetime near single metal nanoparticles. Sci Rep 2016; 6:21349. [PMID: 26876780 PMCID: PMC4753420 DOI: 10.1038/srep21349] [Citation(s) in RCA: 25] [Impact Index Per Article: 3.1] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/30/2015] [Accepted: 01/05/2016] [Indexed: 11/17/2022] Open
Abstract
We explore the spectral dependence of fluorescence enhancement and the associated lifetime modification of fluorescent molecules coupled to single metal nanoparticles. Fluorescence lifetime imaging microscopy and single-particle dark-field spectroscopy are combined to correlate the dependence of fluorescence lifetime reduction on the spectral overlap between the fluorescence emission and the localised surface plasmon (LSP) spectra of individual gold nanoparticles. A maximum lifetime reduction is observed when the fluorescence and LSP resonances coincide, with good agreement provided by numerical simulations. The explicit comparison between experiment and simulation, that we obtain, offers an insight into the spectral engineering of LSP mediated fluorescence and may lead to optimized application in sensing and biomedicine.
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34
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Fitzgerald C, Hosny NA, Tong H, Seville PC, Gallimore PJ, Davidson NM, Athanasiadis A, Botchway SW, Ward AD, Kalberer M, Kuimova MK, Pope FD. Fluorescence lifetime imaging of optically levitated aerosol: a technique to quantitatively map the viscosity of suspended aerosol particles. Phys Chem Chem Phys 2016; 18:21710-9. [DOI: 10.1039/c6cp03674k] [Citation(s) in RCA: 21] [Impact Index Per Article: 2.6] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/17/2022]
Abstract
A technique to measure the viscosity of stably levitated single micron-sized aerosol particles.
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Affiliation(s)
- C. Fitzgerald
- Department of Chemistry
- University of Cambridge
- Cambridge
- UK
| | - N. A. Hosny
- Department of Chemistry
- Imperial College London
- London
- UK
| | - H. Tong
- Department of Chemistry
- University of Cambridge
- Cambridge
- UK
| | - P. C. Seville
- School of Pharmacy and Biomedical Sciences
- University of Central Lancashire
- Preston
- UK
| | | | - N. M. Davidson
- School of Geography
- Earth and Environmental Sciences
- University of Birmingham
- Birmingham
- UK
| | | | - S. W. Botchway
- The Science and Technology Facilities Council
- Rutherford Appleton Laboratory
- Research Complex at Harwell
- Oxfordshire
- UK
| | - A. D. Ward
- The Science and Technology Facilities Council
- Rutherford Appleton Laboratory
- Research Complex at Harwell
- Oxfordshire
- UK
| | - M. Kalberer
- Department of Chemistry
- University of Cambridge
- Cambridge
- UK
| | - M. K. Kuimova
- Department of Chemistry
- Imperial College London
- London
- UK
| | - F. D. Pope
- School of Geography
- Earth and Environmental Sciences
- University of Birmingham
- Birmingham
- UK
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35
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Hosny NA, Fitzgerald C, Vyšniauskas A, Athanasiadis A, Berkemeier T, Uygur N, Pöschl U, Shiraiwa M, Kalberer M, Pope FD, Kuimova MK. Direct imaging of changes in aerosol particle viscosity upon hydration and chemical aging. Chem Sci 2015; 7:1357-1367. [PMID: 29910892 PMCID: PMC5975791 DOI: 10.1039/c5sc02959g] [Citation(s) in RCA: 50] [Impact Index Per Article: 5.6] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/10/2015] [Accepted: 11/07/2015] [Indexed: 12/22/2022] Open
Abstract
We report quantitative, real-time, online observations of microscopic viscosity changes in aerosol particles of atmospherically relevant composition, using fluorescence lifetime imaging (FLIM) of viscosity.
Organic aerosol particles (OA) play major roles in atmospheric chemistry, climate, and public health. Aerosol particle viscosity is highly important since it can determine the ability of chemical species such as oxidants, organics or water to diffuse into the particle bulk. Recent measurements indicate that OA may be present in highly viscous states, however, diffusion rates of small molecules such as water are not limited by these high viscosities. Direct observational evidence of kinetic barriers caused by high viscosity and low diffusivity in aerosol particles were not available until recently; and techniques that are able to dynamically quantify and track viscosity changes during atmospherically relevant processes are still unavailable for atmospheric aerosols. Here we report quantitative, real-time, online observations of microscopic viscosity changes in aerosol particles of atmospherically relevant composition, using fluorescence lifetime imaging (FLIM) of viscosity. We show that microviscosity in ozonated oleic acid droplets and secondary organic aerosol (SOA) particles formed by ozonolysis of myrcene increases substantially with decreasing humidity and atmospheric oxidative aging processes. Furthermore, we found unexpected heterogeneities of microviscosity inside individual aerosol particles. The results of this study enhance our understanding of organic aerosol processes on microscopic scales and may have important implications for the modeling of atmospheric aerosol growth, composition and interactions with trace gases and clouds.
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Affiliation(s)
- N A Hosny
- Department of Chemistry , Imperial College London , London , SW7 2AZ , UK .
| | - C Fitzgerald
- Department of Chemistry , University of Cambridge , Cambridge , CB2 1EW , UK .
| | - A Vyšniauskas
- Department of Chemistry , Imperial College London , London , SW7 2AZ , UK .
| | - A Athanasiadis
- Department of Chemistry , Imperial College London , London , SW7 2AZ , UK .
| | - T Berkemeier
- Multiphase Chemistry Department , Max Planck Institute for Chemistry , Hahn-Meitner Weg 1 , 55128 , Mainz , Germany
| | - N Uygur
- Multiphase Chemistry Department , Max Planck Institute for Chemistry , Hahn-Meitner Weg 1 , 55128 , Mainz , Germany
| | - U Pöschl
- Multiphase Chemistry Department , Max Planck Institute for Chemistry , Hahn-Meitner Weg 1 , 55128 , Mainz , Germany
| | - M Shiraiwa
- Multiphase Chemistry Department , Max Planck Institute for Chemistry , Hahn-Meitner Weg 1 , 55128 , Mainz , Germany
| | - M Kalberer
- Department of Chemistry , University of Cambridge , Cambridge , CB2 1EW , UK .
| | - F D Pope
- School of Geography , Earth and Environmental Science , University of Birmingham , Edgbaston , B15 2TT , UK .
| | - M K Kuimova
- Department of Chemistry , Imperial College London , London , SW7 2AZ , UK .
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36
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Levitt JA, Morton PE, Fruhwirth GO, Santis G, Chung PH, Parsons M, Suhling K. Simultaneous FRAP, FLIM and FAIM for measurements of protein mobility and interaction in living cells. BIOMEDICAL OPTICS EXPRESS 2015; 6:3842-54. [PMID: 26504635 PMCID: PMC4605044 DOI: 10.1364/boe.6.003842] [Citation(s) in RCA: 11] [Impact Index Per Article: 1.2] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 06/29/2015] [Revised: 08/16/2015] [Accepted: 08/18/2015] [Indexed: 05/23/2023]
Abstract
We present a novel integrated multimodal fluorescence microscopy technique for simultaneous fluorescence recovery after photobleaching (FRAP), fluorescence lifetime imaging (FLIM) and fluorescence anisotropy imaging (FAIM). This approach captures a series of polarization-resolved fluorescence lifetime images during a FRAP recovery, maximizing the information available from a limited photon budget. We have applied this method to analyse the behaviour of GFP-labelled coxsackievirus and adenovirus receptor (CAR) in living human epithelial cells. Our data reveal that CAR exists in oligomeric states throughout the cell, and that these complexes occur in conjunction with high immobile fractions of the receptor at cell-cell junctions. These findings shed light on previously unknown molecular associations between CAR receptors in intact cells and demonstrate the power of combined FRAP, FLIM and FAIM microscopy as a robust method to analyse complex multi-component dynamics in living cells.
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Affiliation(s)
- James A. Levitt
- Department of Physics, King’s College London, Strand, London WC2R 2LS, UK
| | - Penny E. Morton
- Division of Asthma, Allergy, and Lung Biology, Guys Campus, King’s College London, London, UK
- Randall Division of Cell and Molecular Biophysics, Guys Campus, King’s College London, London, SE1 1UL, UK
| | - Gilbert O. Fruhwirth
- Department of Imaging Chemsitry and Biology, Division of Imaging Sciences and Biomedical Engineering, St. Thomas Hospital, King's College London, SE1 7EH, UK
| | - George Santis
- Division of Asthma, Allergy, and Lung Biology, Guys Campus, King’s College London, London, UK
| | - Pei-Hua Chung
- Department of Physics, King’s College London, Strand, London WC2R 2LS, UK
| | - Maddy Parsons
- Randall Division of Cell and Molecular Biophysics, Guys Campus, King’s College London, London, SE1 1UL, UK
| | - Klaus Suhling
- Department of Physics, King’s College London, Strand, London WC2R 2LS, UK
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37
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The interactions between a small molecule and G-quadruplexes are visualized by fluorescence lifetime imaging microscopy. Nat Commun 2015; 6:8178. [PMID: 26350962 PMCID: PMC4579598 DOI: 10.1038/ncomms9178] [Citation(s) in RCA: 172] [Impact Index Per Article: 19.1] [Reference Citation Analysis] [Abstract] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/20/2015] [Accepted: 07/25/2015] [Indexed: 12/12/2022] Open
Abstract
Guanine-rich oligonucleotides can fold into quadruple-stranded helical structures known as G-quadruplexes. Mounting experimental evidence has gathered suggesting that these non-canonical nucleic acid structures form in vivo and play essential biological roles. However, to date, there are no small-molecule optical probes to image G-quadruplexes in live cells. Herein, we report the design and development of a small fluorescent molecule, which can be used as an optical probe for G-quadruplexes. We demonstrate that the fluorescence lifetime of this new probe changes considerably upon interaction with different nucleic acid topologies. Specifically, longer fluorescence lifetimes are observed in vitro for G-quadruplexes than for double- and single-stranded nucleic acids. Cellular studies confirm that this molecule is cell permeable, has low cytotoxicity and localizes primarily in the cell nucleus. Furthermore, using fluorescence lifetime imaging microscopy, live-cell imaging suggests that the probe can be used to study the interaction of small molecules with G-quadruplexes in vivo. Guanine-rich oligonucleotides can fold into secondary structures known as G-quadruplexes that are proposed to have various biological roles. Here, Shivalingam et al. develop a cell-permeable, low-toxicity probe that can be used to probe interactions between G-quadruplexes and small molecules in vivo.
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38
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Levitt JA, Chung PH, Suhling K. Spectrally resolved fluorescence lifetime imaging of Nile red for measurements of intracellular polarity. JOURNAL OF BIOMEDICAL OPTICS 2015; 20:096002. [PMID: 26334975 DOI: 10.1117/1.jbo.20.9.096002] [Citation(s) in RCA: 26] [Impact Index Per Article: 2.9] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 01/20/2015] [Accepted: 07/17/2015] [Indexed: 06/05/2023]
Abstract
Spectrally resolved confocal microscopy and fluorescence lifetime imaging have been used to measure the polarity of lipid-rich regions in living HeLa cells stained with Nile red. The emission peak from the solvatochromic dye in lipid droplets is at a shorter wavelength than other, more polar, stained internal membranes, and this is indicative of a low polarity environment. We estimate that the dielectric constant, ϵ , is around 5 in lipid droplets and 25<ϵ<40 in other lipid-rich regions. Our spectrally resolved fluorescence lifetime imaging microscopy (FLIM) data show that intracellular Nile red exhibits complex, multiexponential fluorescence decays due to emission from a short lifetime locally excited state and a longer lifetime intramolecular charge transfer state. We measure an increase in the average fluorescence lifetime of the dye with increasing emission wavelength, as shown using phasor plots of the FLIM data. We also show using these phasor plots that the shortest lifetime decay components arise from lipid droplets. Thus, fluorescence lifetime is a viable contrast parameter for distinguishing lipid droplets from other stained lipid-rich regions. Finally, we discuss the FLIM of Nile red as a method for simultaneously mapping both polarity and relative viscosity based on fluorescence lifetime measurements.
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39
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Thompson AJ, Herling TW, Kubánková M, Vyšniauskas A, Knowles TPJ, Kuimova MK. Molecular Rotors Provide Insights into Microscopic Structural Changes During Protein Aggregation. J Phys Chem B 2015; 119:10170-9. [DOI: 10.1021/acs.jpcb.5b05099] [Citation(s) in RCA: 29] [Impact Index Per Article: 3.2] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/29/2022]
Affiliation(s)
| | - Therese W. Herling
- Department
of Chemistry, University of Cambridge, Lensfield Road, Cambridge CB2 1EW, U.K
| | - Markéta Kubánková
- Chemistry
Department, Imperial College London, Exhibition Road, London SW7 2AZ, U.K
| | - Aurimas Vyšniauskas
- Chemistry
Department, Imperial College London, Exhibition Road, London SW7 2AZ, U.K
| | - Tuomas P. J. Knowles
- Department
of Chemistry, University of Cambridge, Lensfield Road, Cambridge CB2 1EW, U.K
| | - Marina K. Kuimova
- Chemistry
Department, Imperial College London, Exhibition Road, London SW7 2AZ, U.K
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40
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Nuciforo P, Radosevic-Robin N, Ng T, Scaltriti M. Quantification of HER family receptors in breast cancer. Breast Cancer Res 2015; 17:53. [PMID: 25887735 PMCID: PMC4389676 DOI: 10.1186/s13058-015-0561-8] [Citation(s) in RCA: 35] [Impact Index Per Article: 3.9] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/17/2022] Open
Abstract
The clinical success of trastuzumab in breast cancer taught us that appropriate tumor evaluation is mandatory for the correct identification of patients eligible for targeted therapies. Although HER2 protein expression by immunohistochemistry (IHC) and gene amplification by fluorescence in situ hybridization (FISH) assays are routinely used to select patients to receive trastuzumab, both assays only partially predict response to the drug. In the case of epidermal growth factor receptor (EGFR), the link between the presence of the receptor or its amplification and response to anti-EGFR therapies could not be demonstrated. Even less is known for HER3 and HER4, mainly due to lack of robust and validated assays detecting these proteins. It is becoming evident that, besides FISH and IHC, we need better assays to quantify HER receptors and categorize the patients for individualized treatments. Here, we present the current available methodologies to measure HER family receptors and discuss the clinical implications of target quantification.
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Affiliation(s)
- Paolo Nuciforo
- Molecular Oncology Laboratory, Vall d'Hebron Institute of Oncology, Passeig Vall d'Hebron 119-129, Barcelona, 08035, Spain.
- Universitat Autònoma de Barcelona, Barcelona, 08035, Spain.
| | - Nina Radosevic-Robin
- ERTICa Research Group, University of Auvergne EA4677, 63000, Clermont-Ferrand, France.
- Biopathology, Jean Perrin Comprehensive Cancer Center, 58 rue Montalembert, 63011, Clermont-Ferrand, France.
| | - Tony Ng
- Richard Dimbleby Department of Cancer Research, Randall Division of Cell and Molecular Biophysics and Division of Cancer Studies, King's College London, London, SE1 1UL, UK.
- UCL Cancer Institute, Paul O'Gorman Building, University College London, London, WC1E 6DD, UK.
- Breakthrough Breast Cancer Research Unit, Department of Research Oncology, Guy's Hospital King's College London School of Medicine, London, SE1 9RT, UK.
| | - Maurizio Scaltriti
- Human Oncology and Pathogenesis Program (HOPP), Memorial Sloan Kettering Cancer Center, 1275 York Avenue, Box 20, New York, NY, 10065, USA.
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41
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Coban O, Zanetti-Dominguez LC, Matthews DR, Rolfe DJ, Weitsman G, Barber PR, Barbeau J, Devauges V, Kampmeier F, Winn M, Vojnovic B, Parker PJ, Lidke KA, Lidke DS, Ameer-Beg SM, Martin-Fernandez ML, Ng T. Effect of phosphorylation on EGFR dimer stability probed by single-molecule dynamics and FRET/FLIM. Biophys J 2015; 108:1013-26. [PMID: 25762314 PMCID: PMC4375452 DOI: 10.1016/j.bpj.2015.01.005] [Citation(s) in RCA: 34] [Impact Index Per Article: 3.8] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/01/2014] [Revised: 12/06/2014] [Accepted: 01/07/2015] [Indexed: 12/22/2022] Open
Abstract
Deregulation of epidermal growth factor receptor (EGFR) signaling has been correlated with the development of a variety of human carcinomas. EGF-induced receptor dimerization and consequent trans- auto-phosphorylation are among the earliest events in signal transduction. Binding of EGF is thought to induce a conformational change that consequently unfolds an ectodomain loop required for dimerization indirectly. It may also induce important allosteric changes in the cytoplasmic domain. Despite extensive knowledge on the physiological activation of EGFR, the effect of targeted therapies on receptor conformation is not known and this particular aspect of receptor function, which can potentially be influenced by drug treatment, may in part explain the heterogeneous clinical response among cancer patients. Here, we used Förster resonance energy transfer/fluorescence lifetime imaging microscopy (FRET/FLIM) combined with two-color single-molecule tracking to study the effect of ATP-competitive small molecule tyrosine kinase inhibitors (TKIs) and phosphatase-based manipulation of EGFR phosphorylation on live cells. The distribution of dimer on-times was fitted to a monoexponential to extract dimer off-rates (koff). Our data show that pretreatment with gefitinib (active conformation binder) stabilizes the EGFR ligand-bound homodimer. Overexpression of EGFR-specific DEP-1 phosphatase was also found to have a stabilizing effect on the homodimer. No significant difference in the koff of the dimer could be detected when an anti-EGFR antibody (425 Snap single-chain variable fragment) that allows for dimerization of ligand-bound receptors, but not phosphorylation, was used. These results suggest that both the conformation of the extracellular domain and phosphorylation status of the receptor are involved in modulating the stability of the dimer. The relative fractions of these two EGFR subpopulations (interacting versus free) were obtained by a fractional-intensity analysis of ensemble FRET/FLIM images. Our combined imaging approach showed that both the fraction and affinity (surrogate of conformation at a single-molecule level) increased after gefitinib pretreatment or DEP-1 phosphatase overexpression. Using an EGFR mutation (I706Q, V948R) that perturbs the ability of EGFR to dimerize intracellularly, we showed that a modest drug-induced increase in the fraction/stability of the EGFR homodimer may have a significant biological impact on the tumor cell's proliferation potential.
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Affiliation(s)
- Oana Coban
- Richard Dimbleby Department of Cancer Research, King's College London, London, UK; Randall Division of Cellular and Molecular Biophysics, King's College London, London, UK.
| | - Laura C Zanetti-Dominguez
- Science and Technology Facilities Council, Research Complex at Harwell, Rutherford Appleton Laboratory, Didcot, UK
| | - Daniel R Matthews
- Richard Dimbleby Department of Cancer Research, King's College London, London, UK; Randall Division of Cellular and Molecular Biophysics, King's College London, London, UK
| | - Daniel J Rolfe
- Science and Technology Facilities Council, Research Complex at Harwell, Rutherford Appleton Laboratory, Didcot, UK
| | - Gregory Weitsman
- Richard Dimbleby Department of Cancer Research, King's College London, London, UK; Randall Division of Cellular and Molecular Biophysics, King's College London, London, UK
| | - Paul R Barber
- Gray Institute for Radiation Oncology & Biology, Department of Oncology, University of Oxford, Oxford, UK
| | - Jody Barbeau
- Richard Dimbleby Department of Cancer Research, King's College London, London, UK; Randall Division of Cellular and Molecular Biophysics, King's College London, London, UK
| | - Viviane Devauges
- Richard Dimbleby Department of Cancer Research, King's College London, London, UK; Randall Division of Cellular and Molecular Biophysics, King's College London, London, UK
| | - Florian Kampmeier
- Division of Imaging Sciences, King's College London, The Rayne Institute, St. Thomas Hospital, London, UK
| | - Martyn Winn
- Computational Science and Engineering Department, Science and Technology Facilities Council, Rutherford Appleton Laboratory, Didcot, UK
| | - Borivoj Vojnovic
- Randall Division of Cellular and Molecular Biophysics, King's College London, London, UK; Gray Institute for Radiation Oncology & Biology, Department of Oncology, University of Oxford, Oxford, UK
| | - Peter J Parker
- Division of Cancer Studies, King's College London, London, UK; Cancer Research UK, London Research Institute, London, UK
| | - Keith A Lidke
- Department of Physics and Astronomy, University of New Mexico, Albuquerque, New Mexico
| | - Diane S Lidke
- Department of Pathology, University of New Mexico School of Medicine, Albuquerque, New Mexico; Cancer Research and Treatment Center, University of New Mexico School of Medicine, Albuquerque, New Mexico
| | - Simon M Ameer-Beg
- Richard Dimbleby Department of Cancer Research, King's College London, London, UK; Randall Division of Cellular and Molecular Biophysics, King's College London, London, UK; Division of Cancer Studies, King's College London, London, UK
| | - Marisa L Martin-Fernandez
- Science and Technology Facilities Council, Research Complex at Harwell, Rutherford Appleton Laboratory, Didcot, UK
| | - Tony Ng
- Richard Dimbleby Department of Cancer Research, King's College London, London, UK; Randall Division of Cellular and Molecular Biophysics, King's College London, London, UK; Division of Cancer Studies, King's College London, London, UK
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Nedbal J, Visitkul V, Ortiz-Zapater E, Weitsman G, Chana P, Matthews DR, Ng T, Ameer-Beg SM. Time-domain microfluidic fluorescence lifetime flow cytometry for high-throughput Förster resonance energy transfer screening. Cytometry A 2015; 87:104-18. [PMID: 25523156 PMCID: PMC4440390 DOI: 10.1002/cyto.a.22616] [Citation(s) in RCA: 27] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/15/2014] [Revised: 11/12/2014] [Accepted: 12/03/2014] [Indexed: 01/22/2023]
Abstract
Sensing ion or ligand concentrations, physico-chemical conditions, and molecular dimerization or conformation change is possible by assays involving fluorescent lifetime imaging. The inherent low throughput of imaging impedes rigorous statistical data analysis on large cell numbers. We address this limitation by developing a fluorescence lifetime-measuring flow cytometer for fast fluorescence lifetime quantification in living or fixed cell populations. The instrument combines a time-correlated single photon counting epifluorescent microscope with microfluidics cell-handling system. The associated computer software performs burst integrated fluorescence lifetime analysis to assign fluorescence lifetime, intensity, and burst duration to each passing cell. The maximum safe throughput of the instrument reaches 3,000 particles per minute. Living cells expressing spectroscopic rulers of varying peptide lengths were distinguishable by Förster resonant energy transfer measured by donor fluorescence lifetime. An epidermal growth factor (EGF)-stimulation assay demonstrated the technique's capacity to selectively quantify EGF receptor phosphorylation in cells, which was impossible by measuring sensitized emission on a standard flow cytometer. Dual-color fluorescence lifetime detection and cell-specific chemical environment sensing were exemplified using di-4-ANEPPDHQ, a lipophilic environmentally sensitive dye that exhibits changes in its fluorescence lifetime as a function of membrane lipid order. To our knowledge, this instrument opens new applications in flow cytometry which were unavailable due to technological limitations of previously reported fluorescent lifetime flow cytometers. The presented technique is sensitive to lifetimes of most popular fluorophores in the 0.5-5 ns range including fluorescent proteins and is capable of detecting multi-exponential fluorescence lifetime decays. This instrument vastly enhances the throughput of experiments involving fluorescence lifetime measurements, thereby providing statistically significant quantitative data for analysis of large cell populations. © 2014 International Society for Advancement of Cytometry.
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Affiliation(s)
- Jakub Nedbal
- Division of Cancer Studies, King's College LondonUnited Kingdom
- Randall Division of Cell and Molecular Biophysics, King's College LondonUnited Kingdom
| | - Viput Visitkul
- Randall Division of Cell and Molecular Biophysics, King's College LondonUnited Kingdom
| | - Elena Ortiz-Zapater
- Division of Asthma, Allergy & Lung Biology, King's College LondonUnited Kingdom
| | | | - Prabhjoat Chana
- Immune Monitoring Laboratory, NIHR Biomedical Research Centre at Guy's and St Thomas' NHS Foundation Trust and King's College LondonUnited Kingdom
| | - Daniel R Matthews
- Queensland Brain Institute, The University of QueenslandSt Lucia, Australia
| | - Tony Ng
- Division of Cancer Studies, King's College LondonUnited Kingdom
- Randall Division of Cell and Molecular Biophysics, King's College LondonUnited Kingdom
- UCL Cancer Institute, University College LondonUnited Kingdom
| | - Simon M Ameer-Beg
- Division of Cancer Studies, King's College LondonUnited Kingdom
- Randall Division of Cell and Molecular Biophysics, King's College LondonUnited Kingdom
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43
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Poland SP, Krstajić N, Monypenny J, Coelho S, Tyndall D, Walker RJ, Devauges V, Richardson J, Dutton N, Barber P, Li DDU, Suhling K, Ng T, Henderson RK, Ameer-Beg SM. A high speed multifocal multiphoton fluorescence lifetime imaging microscope for live-cell FRET imaging. BIOMEDICAL OPTICS EXPRESS 2015; 6:277-96. [PMID: 25780724 PMCID: PMC4354599 DOI: 10.1364/boe.6.000277] [Citation(s) in RCA: 49] [Impact Index Per Article: 5.4] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 09/12/2014] [Revised: 11/28/2014] [Accepted: 11/30/2014] [Indexed: 05/18/2023]
Abstract
We demonstrate diffraction limited multiphoton imaging in a massively parallel, fully addressable time-resolved multi-beam multiphoton microscope capable of producing fluorescence lifetime images with sub-50ps temporal resolution. This imaging platform offers a significant improvement in acquisition speed over single-beam laser scanning FLIM by a factor of 64 without compromising in either the temporal or spatial resolutions of the system. We demonstrate FLIM acquisition at 500 ms with live cells expressing green fluorescent protein. The applicability of the technique to imaging protein-protein interactions in live cells is exemplified by observation of time-dependent FRET between the epidermal growth factor receptor (EGFR) and the adapter protein Grb2 following stimulation with the receptor ligand. Furthermore, ligand-dependent association of HER2-HER3 receptor tyrosine kinases was observed on a similar timescale and involved the internalisation and accumulation or receptor heterodimers within endosomes. These data demonstrate the broad applicability of this novel FLIM technique to the spatio-temporal dynamics of protein-protein interaction.
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Affiliation(s)
- Simon P. Poland
- Division of Cancer Studies, Guy’s Campus, Kings College, London,
UK
- Randall Division of Cell and Molecular Biophysics, Guy’s Campus, Kings College, London,
UK
| | - Nikola Krstajić
- Institute for Integrated Micro and Nano Systems, School of Engineering, University of Edinburgh, Edinburgh,
UK
| | - James Monypenny
- Division of Cancer Studies, Guy’s Campus, Kings College, London,
UK
- Randall Division of Cell and Molecular Biophysics, Guy’s Campus, Kings College, London,
UK
| | - Simao Coelho
- Division of Cancer Studies, Guy’s Campus, Kings College, London,
UK
- Randall Division of Cell and Molecular Biophysics, Guy’s Campus, Kings College, London,
UK
| | - David Tyndall
- Institute for Integrated Micro and Nano Systems, School of Engineering, University of Edinburgh, Edinburgh,
UK
| | - Richard J. Walker
- Institute for Integrated Micro and Nano Systems, School of Engineering, University of Edinburgh, Edinburgh,
UK
- Photon-Force Ltd., Edinburgh,
UK
| | - Viviane Devauges
- Division of Cancer Studies, Guy’s Campus, Kings College, London,
UK
- Randall Division of Cell and Molecular Biophysics, Guy’s Campus, Kings College, London,
UK
| | - Justin Richardson
- Institute for Integrated Micro and Nano Systems, School of Engineering, University of Edinburgh, Edinburgh,
UK
- Photon-Force Ltd., Edinburgh,
UK
| | - Neale Dutton
- Institute for Integrated Micro and Nano Systems, School of Engineering, University of Edinburgh, Edinburgh,
UK
| | - Paul Barber
- Gray Institute for Radiation Oncology & Biology, University of Oxford, Old Road Campus Research Building, Roosevelt Drive, Oxford OX3 7DQ
UK
| | - David Day-Uei Li
- Strathclyde Institute of Pharmacy and Biomedical Sciences, 161 Cathedral Street, Glasgow, G4 0RE,
UK
| | - Klaus Suhling
- Department of Physics, King's College London, Strand, London,
UK
| | - Tony Ng
- Division of Cancer Studies, Guy’s Campus, Kings College, London,
UK
- Randall Division of Cell and Molecular Biophysics, Guy’s Campus, Kings College, London,
UK
- UCL Cancer Institute, Paul O'Gorman Building, University College London, London WC1E 6DD,
UK
| | - Robert K. Henderson
- Institute for Integrated Micro and Nano Systems, School of Engineering, University of Edinburgh, Edinburgh,
UK
| | - Simon M. Ameer-Beg
- Division of Cancer Studies, Guy’s Campus, Kings College, London,
UK
- Randall Division of Cell and Molecular Biophysics, Guy’s Campus, Kings College, London,
UK
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44
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45
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Devauges V, Matthews DR, Aluko J, Nedbal J, Levitt JA, Poland SP, Coban O, Weitsman G, Monypenny J, Ng T, Ameer-Beg SM. Steady-state acceptor fluorescence anisotropy imaging under evanescent excitation for visualisation of FRET at the plasma membrane. PLoS One 2014; 9:e110695. [PMID: 25360776 PMCID: PMC4215982 DOI: 10.1371/journal.pone.0110695] [Citation(s) in RCA: 9] [Impact Index Per Article: 0.9] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/25/2014] [Accepted: 09/15/2014] [Indexed: 11/22/2022] Open
Abstract
We present a novel imaging system combining total internal reflection fluorescence (TIRF) microscopy with measurement of steady-state acceptor fluorescence anisotropy in order to perform live cell Förster Resonance Energy Transfer (FRET) imaging at the plasma membrane. We compare directly the imaging performance of fluorescence anisotropy resolved TIRF with epifluorescence illumination. The use of high numerical aperture objective for TIRF required correction for induced depolarization factors. This arrangement enabled visualisation of conformational changes of a Raichu-Cdc42 FRET biosensor by measurement of intramolecular FRET between eGFP and mRFP1. Higher activity of the probe was found at the cell plasma membrane compared to intracellularly. Imaging fluorescence anisotropy in TIRF allowed clear differentiation of the Raichu-Cdc42 biosensor from negative control mutants. Finally, inhibition of Cdc42 was imaged dynamically in live cells, where we show temporal changes of the activity of the Raichu-Cdc42 biosensor.
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Affiliation(s)
- Viviane Devauges
- Richard Dimbleby Cancer Research Laboratory, Division of Cancer Studies and Randall Division of Cell and Molecular Biophysics, King's College London, London, United Kingdom
| | - Daniel R. Matthews
- Richard Dimbleby Cancer Research Laboratory, Division of Cancer Studies and Randall Division of Cell and Molecular Biophysics, King's College London, London, United Kingdom
| | - Justin Aluko
- Department of Physics, King's College London, London, United Kingdom
| | - Jakub Nedbal
- Richard Dimbleby Cancer Research Laboratory, Division of Cancer Studies and Randall Division of Cell and Molecular Biophysics, King's College London, London, United Kingdom
| | - James A. Levitt
- Richard Dimbleby Cancer Research Laboratory, Division of Cancer Studies and Randall Division of Cell and Molecular Biophysics, King's College London, London, United Kingdom
| | - Simon P. Poland
- Richard Dimbleby Cancer Research Laboratory, Division of Cancer Studies and Randall Division of Cell and Molecular Biophysics, King's College London, London, United Kingdom
| | - Oana Coban
- Richard Dimbleby Cancer Research Laboratory, Division of Cancer Studies and Randall Division of Cell and Molecular Biophysics, King's College London, London, United Kingdom
| | - Gregory Weitsman
- Richard Dimbleby Cancer Research Laboratory, Division of Cancer Studies and Randall Division of Cell and Molecular Biophysics, King's College London, London, United Kingdom
| | - James Monypenny
- Richard Dimbleby Cancer Research Laboratory, Division of Cancer Studies and Randall Division of Cell and Molecular Biophysics, King's College London, London, United Kingdom
| | - Tony Ng
- Richard Dimbleby Cancer Research Laboratory, Division of Cancer Studies and Randall Division of Cell and Molecular Biophysics, King's College London, London, United Kingdom
- UCL Cancer Institute, University College London, London, United Kingdom
| | - Simon M. Ameer-Beg
- Richard Dimbleby Cancer Research Laboratory, Division of Cancer Studies and Randall Division of Cell and Molecular Biophysics, King's College London, London, United Kingdom
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46
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Manickavasagam A, Hirvonen LM, Melita LN, Chong EZ, Cook RJ, Bozec L, Festy F. Multimodal optical characterisation of collagen photodegradation by femtosecond infrared laser ablation. Analyst 2014; 139:6135-43. [PMID: 25318007 DOI: 10.1039/c4an01523a] [Citation(s) in RCA: 14] [Impact Index Per Article: 1.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/21/2022]
Abstract
Collagen is a structural component of the human body, as a connective tissue it can become altered as a result of pathophysiological conditions. Although the collagen degradation mechanism is not fully understood, it plays an important role in ageing, disease progression and applications in therapeutic laser treatments. To fully understand the mechanism of collagen alteration, in our study photo-disruptive effects were induced in collagen I matrix by point-irradiation with a femtosecond Ti-sapphire laser under controlled laser ablation settings. This was followed by multi-modal imaging of the irradiated and surrounding areas to analyse the degradation mechanism. Our multi-modal methodology was based on second harmonic generation (SHG), scanning electron microscope (SEM), autofluorescence (AF) average intensities and the average fluorescence lifetime. This allowed us to quantitatively characterise the degraded area into four distinct zones: (1) depolymerised zone in the laser focal spot as indicated by the loss of SHG signal, (2) enhanced crosslinking zone in the inner boundary of the laser induced cavity as represented by the high fluorescence ring, (3) reduced crosslinking zone formed the outer boundary of the cavity as marked by the increased SHG signal and (4) native collagen. These identified distinct zones were in good agreement with the expected photochemical changes shown using Raman spectroscopy. In addition, imaging using polarisation-resolved SHG (p-SHG) revealed both a high degree of fibre re-orientation and a SHG change in tensor ratios around the irradiation spot. Our multi-modal optical imaging approach can provide a new methodology for defining distinct zones that can be used in a clinical setting to determine suitable thresholds for applying safe laser treatments without affecting the surrounding tissues. Furthermore this technique can be extended to address challenges observed in collagen based tissue engineering and used as a minimally invasive diagnostic tool to characterise diseased and non-diseased collagen rich tissues.
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Affiliation(s)
- A Manickavasagam
- Biomaterial, Biomimetics & Biophotonics Division, King's College London Dental Institute, London, UK.
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47
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Hirvonen LM, Festy F, Suhling K. Wide-field time-correlated single-photon counting (TCSPC) lifetime microscopy with microsecond time resolution. OPTICS LETTERS 2014; 39:5602-5. [PMID: 25360938 DOI: 10.1364/ol.39.005602] [Citation(s) in RCA: 16] [Impact Index Per Article: 1.6] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 05/25/2023]
Abstract
A 1 MHz frame rate complementary metal-oxide semiconductor (CMOS) camera was used in combination with an image intensifier for wide-field time-correlated single-photon counting (TCSPC) imaging. The system combines an ultrafast frame rate with single-photon sensitivity and was employed on a fluorescence microscope to image decays of ruthenium compound Ru(dpp) with lifetimes from around 1 to 5 μs. A submicrowatt excitation power over the whole field of view is sufficient for this approach, and compatibility with live-cell imaging was demonstrated by imaging europium-containing beads with a lifetime of 570 μs in living HeLa cells. A standard two-photon excitation scanning fluorescence lifetime imaging (FLIM) system was used to independently verify the lifetime for the europium beads. This approach brings together advantageous features for time-resolved live-cell imaging such as low excitation intensity, single-photon sensitivity, ultrafast camera frame rates, and short acquisition times.
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48
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Kiuchi T, Ortiz-Zapater E, Monypenny J, Matthews DR, Nguyen LK, Barbeau J, Coban O, Lawler K, Burford B, Rolfe DJ, de Rinaldis E, Dafou D, Simpson MA, Woodman N, Pinder S, Gillett CE, Devauges V, Poland SP, Fruhwirth G, Marra P, Boersma YL, Plückthun A, Gullick WJ, Yarden Y, Santis G, Winn M, Kholodenko BN, Martin-Fernandez ML, Parker P, Tutt A, Ameer-Beg SM, Ng T. The ErbB4 CYT2 variant protects EGFR from ligand-induced degradation to enhance cancer cell motility. Sci Signal 2014; 7:ra78. [PMID: 25140053 DOI: 10.1126/scisignal.2005157] [Citation(s) in RCA: 32] [Impact Index Per Article: 3.2] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/16/2022]
Abstract
The epidermal growth factor receptor (EGFR) is a member of the ErbB family that can promote the migration and proliferation of breast cancer cells. Therapies that target EGFR can promote the dimerization of EGFR with other ErbB receptors, which is associated with the development of drug resistance. Understanding how interactions among ErbB receptors alter EGFR biology could provide avenues for improving cancer therapy. We found that EGFR interacted directly with the CYT1 and CYT2 variants of ErbB4 and the membrane-anchored intracellular domain (mICD). The CYT2 variant, but not the CYT1 variant, protected EGFR from ligand-induced degradation by competing with EGFR for binding to a complex containing the E3 ubiquitin ligase c-Cbl and the adaptor Grb2. Cultured breast cancer cells overexpressing both EGFR and ErbB4 CYT2 mICD exhibited increased migration. With molecular modeling, we identified residues involved in stabilizing the EGFR dimer. Mutation of these residues in the dimer interface destabilized the complex in cells and abrogated growth factor-stimulated cell migration. An exon array analysis of 155 breast tumors revealed that the relative mRNA abundance of the ErbB4 CYT2 variant was increased in ER+ HER2- breast cancer patients, suggesting that our findings could be clinically relevant. We propose a mechanism whereby competition for binding to c-Cbl in an ErbB signaling heterodimer promotes migration in response to a growth factor gradient.
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Affiliation(s)
- Tai Kiuchi
- Richard Dimbleby Department of Cancer Research, Randall Division of Cell and Molecular Biophysics, King's College London, Guy's Medical School Campus, London SE1 1UL, UK. Division of Cancer Studies, King's College London, London SE1 1UL, UK. Breakthrough Breast Cancer Research Unit, Research Oncology, King's College London, Guy's Hospital, London SE1 9RT, UK
| | - Elena Ortiz-Zapater
- Department of Asthma, Allergy and Respiratory Science, King's College London, Guy's Hospital, London SE1 9RT, UK
| | - James Monypenny
- Richard Dimbleby Department of Cancer Research, Randall Division of Cell and Molecular Biophysics, King's College London, Guy's Medical School Campus, London SE1 1UL, UK. Division of Cancer Studies, King's College London, London SE1 1UL, UK. Breakthrough Breast Cancer Research Unit, Research Oncology, King's College London, Guy's Hospital, London SE1 9RT, UK
| | - Daniel R Matthews
- Richard Dimbleby Department of Cancer Research, Randall Division of Cell and Molecular Biophysics, King's College London, Guy's Medical School Campus, London SE1 1UL, UK. Division of Cancer Studies, King's College London, London SE1 1UL, UK
| | - Lan K Nguyen
- Systems Biology Ireland, University College Dublin, Belfield, Dublin 4, Ireland
| | - Jody Barbeau
- Richard Dimbleby Department of Cancer Research, Randall Division of Cell and Molecular Biophysics, King's College London, Guy's Medical School Campus, London SE1 1UL, UK. Division of Cancer Studies, King's College London, London SE1 1UL, UK
| | - Oana Coban
- Richard Dimbleby Department of Cancer Research, Randall Division of Cell and Molecular Biophysics, King's College London, Guy's Medical School Campus, London SE1 1UL, UK. Division of Cancer Studies, King's College London, London SE1 1UL, UK
| | - Katherine Lawler
- Richard Dimbleby Department of Cancer Research, Randall Division of Cell and Molecular Biophysics, King's College London, Guy's Medical School Campus, London SE1 1UL, UK. Division of Cancer Studies, King's College London, London SE1 1UL, UK
| | - Brian Burford
- Breakthrough Breast Cancer Research Unit, Research Oncology, King's College London, Guy's Hospital, London SE1 9RT, UK
| | - Daniel J Rolfe
- Central Laser Facility, Rutherford Appleton Laboratory, Science and Technology Facilities Council, Research Complex at Harwell, Didcot OX11 0QX, UK
| | - Emanuele de Rinaldis
- Breakthrough Breast Cancer Research Unit, Research Oncology, King's College London, Guy's Hospital, London SE1 9RT, UK
| | - Dimitra Dafou
- Genetics and Molecular Medicine, King's College London, Guy's Hospital, London SE1 9RT, UK
| | - Michael A Simpson
- Genetics and Molecular Medicine, King's College London, Guy's Hospital, London SE1 9RT, UK
| | - Natalie Woodman
- Guy's and St Thomas' Breast Tissue and Data Bank, King's College London, Guy's Hospital, London SE1 9RT, UK. Research Oncology, Division of Cancer Studies, King's College London, Guy's Hospital, London SE1 9RT, UK
| | - Sarah Pinder
- Guy's and St Thomas' Breast Tissue and Data Bank, King's College London, Guy's Hospital, London SE1 9RT, UK. Research Oncology, Division of Cancer Studies, King's College London, Guy's Hospital, London SE1 9RT, UK
| | - Cheryl E Gillett
- Guy's and St Thomas' Breast Tissue and Data Bank, King's College London, Guy's Hospital, London SE1 9RT, UK. Research Oncology, Division of Cancer Studies, King's College London, Guy's Hospital, London SE1 9RT, UK
| | - Viviane Devauges
- Richard Dimbleby Department of Cancer Research, Randall Division of Cell and Molecular Biophysics, King's College London, Guy's Medical School Campus, London SE1 1UL, UK. Division of Cancer Studies, King's College London, London SE1 1UL, UK
| | - Simon P Poland
- Richard Dimbleby Department of Cancer Research, Randall Division of Cell and Molecular Biophysics, King's College London, Guy's Medical School Campus, London SE1 1UL, UK. Division of Cancer Studies, King's College London, London SE1 1UL, UK
| | - Gilbert Fruhwirth
- Richard Dimbleby Department of Cancer Research, Randall Division of Cell and Molecular Biophysics, King's College London, Guy's Medical School Campus, London SE1 1UL, UK. Division of Cancer Studies, King's College London, London SE1 1UL, UK
| | - Pierfrancesco Marra
- Breakthrough Breast Cancer Research Unit, Research Oncology, King's College London, Guy's Hospital, London SE1 9RT, UK
| | - Ykelien L Boersma
- Department of Biochemistry, University of Zurich, 190, 8057 Zurich, Switzerland
| | - Andreas Plückthun
- Department of Biochemistry, University of Zurich, 190, 8057 Zurich, Switzerland
| | - William J Gullick
- Department of Biosciences, University of Kent, Canterbury, Kent CT2 7NJ, UK
| | - Yosef Yarden
- Department of Biological Regulation, The Weizmann Institute of Science, Rehovot 76100, Israel
| | - George Santis
- Department of Asthma, Allergy and Respiratory Science, King's College London, Guy's Hospital, London SE1 9RT, UK
| | - Martyn Winn
- Computational Science and Engineering Department, Daresbury Laboratory, Science and Technology Facilities Council, Research Complex at Warrington, Warrington WA4 4AD, UK
| | - Boris N Kholodenko
- Systems Biology Ireland, University College Dublin, Belfield, Dublin 4, Ireland
| | - Marisa L Martin-Fernandez
- Central Laser Facility, Rutherford Appleton Laboratory, Science and Technology Facilities Council, Research Complex at Harwell, Didcot OX11 0QX, UK
| | - Peter Parker
- Division of Cancer Studies, King's College London, London SE1 1UL, UK. Protein Phosphorylation Laboratory, Cancer Research UK, London Research Institute, Lincoln's Inn Fields, London WC2A 3PX, UK
| | - Andrew Tutt
- Breakthrough Breast Cancer Research Unit, Research Oncology, King's College London, Guy's Hospital, London SE1 9RT, UK
| | - Simon M Ameer-Beg
- Richard Dimbleby Department of Cancer Research, Randall Division of Cell and Molecular Biophysics, King's College London, Guy's Medical School Campus, London SE1 1UL, UK. Division of Cancer Studies, King's College London, London SE1 1UL, UK.
| | - Tony Ng
- Richard Dimbleby Department of Cancer Research, Randall Division of Cell and Molecular Biophysics, King's College London, Guy's Medical School Campus, London SE1 1UL, UK. Division of Cancer Studies, King's College London, London SE1 1UL, UK. Breakthrough Breast Cancer Research Unit, Research Oncology, King's College London, Guy's Hospital, London SE1 9RT, UK. UCL Cancer Institute, Paul O'Gorman Building, University College London, London WC1E 6BT, UK.
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49
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Abstract
This chapter describes the procedure for globally analyzing fluorescence lifetime imaging (FLIM) data for the observation and quantification of Förster resonance energy transfer (FRET) in live plant cells. The procedure is illustrated by means of a case study, for which plant protoplasts were transfected with different visible fluorescent proteins and subsequently imaged using two-photon excitation FLIM. Spatially resolved fluorescence lifetime images were obtained by application of global analysis using the program Glotaran, which is open-source and freely available software. Using this procedure it is possible to extract the fraction and distance of interacting species between, or conformational changes within proteins, from complex experimental FRET-FLIM datasets, even at low signal-to-noise ratios. In addition, the software allows excluding inherently present autofluorescence from the plant cells, which improves the accuracy of the FRET analysis. The results from the case study are presented and interpreted in the context of the current scientific understanding of these biological systems.
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50
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Damayanti NP, Craig AP, Irudayaraj J. A hybrid FLIM-elastic net platform for label free profiling of breast cancer. Analyst 2014; 138:7127-34. [PMID: 24106733 DOI: 10.1039/c3an01097j] [Citation(s) in RCA: 12] [Impact Index Per Article: 1.2] [Reference Citation Analysis] [Abstract] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/21/2022]
Abstract
We report a label-free fluorescence lifetime profiling strategy to classify breast cancer cells, MCF10CA1h (malignant), MCF10A (nonmalignant), and MCF10AneoT (premalignant) in different stages of malignancy. Fluorescence Lifetime Imaging Microscopy (FLIM) was used to record the lifetime of autofluorescence of endogenous flavin in MCF10 cells in different stages of malignancy. Predominant differences in lifetimes ascertained by multi-exponential fitting curves can be attributed to the different forms of flavin protein; flavin mononucleotide (FMN), free flavin adenine dinucleotide (FAD), semiquinone, and bound FAD. A lifetime map of the metabolite was derived from the contribution of the lifetime of each metabolite by iterative reconvolution fitting of the Time Correlated Single Photon Counting (TCSPC) decay curves. Lifetime maps were constructed by mapping the average lifetime values pixel by pixel using MATLAB. The FLIM image (150 × 150 pixels) of each cell was extracted, resized and centered into 100 × 100 pixels using the nearest neighbor algorithm. Principal Component Analysis (PCA) in conjunction with Elastic net Analysis (EnA) was then used to classify the different stages of MCF10 cell lines based on average lifetime values. The EnA model provided an excellent classification of the cells at different stages of tumorigenesis yielding 100% accuracy.
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Affiliation(s)
- Nur P Damayanti
- Department of Agricultural and Biological Engineering and Bindley Bioscience Center, 225 S. University Street, West Lafayette, IN, USA
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