1
|
Johnson SC, Annamdevula NS, Leavesley SJ, Francis CM, Rich TC. Hyperspectral imaging and dynamic region of interest tracking approaches to quantify localized cAMP signals. Biochem Soc Trans 2024; 52:191-203. [PMID: 38334148 PMCID: PMC11115359 DOI: 10.1042/bst20230352] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/15/2023] [Revised: 01/10/2024] [Accepted: 01/15/2024] [Indexed: 02/10/2024]
Abstract
Cyclic adenosine monophosphate (cAMP) is a ubiquitous second messenger known to orchestrate a myriad of cellular functions over a wide range of timescales. In the last 20 years, a variety of single-cell sensors have been developed to measure second messenger signals including cAMP, Ca2+, and the balance of kinase and phosphatase activities. These sensors utilize changes in fluorescence emission of an individual fluorophore or Förster resonance energy transfer (FRET) to detect changes in second messenger concentration. cAMP and kinase activity reporter probes have provided powerful tools for the study of localized signals. Studies relying on these and related probes have the potential to further revolutionize our understanding of G protein-coupled receptor signaling systems. Unfortunately, investigators have not been able to take full advantage of the potential of these probes due to the limited signal-to-noise ratio of the probes and the limited ability of standard epifluorescence and confocal microscope systems to simultaneously measure the distributions of multiple signals (e.g. cAMP, Ca2+, and changes in kinase activities) in real time. In this review, we focus on recently implemented strategies to overcome these limitations: hyperspectral imaging and adaptive thresholding approaches to track dynamic regions of interest (ROI). This combination of approaches increases signal-to-noise ratio and contrast, and allows identification of localized signals throughout cells. These in turn lead to the identification and quantification of intracellular signals with higher effective resolution. Hyperspectral imaging and dynamic ROI tracking approaches offer investigators additional tools with which to visualize and quantify multiplexed intracellular signaling systems.
Collapse
Affiliation(s)
- Santina C Johnson
- Department of Pharmacology, Frederick P. Whiddon College of Medicine, University of South Alabama, Mobile, AL, U.S.A
- Center for Lung Biology, Frederick P. Whiddon College of Medicine, University of South Alabama, Mobile, AL, U.S.A
| | - Naga S Annamdevula
- Department of Pharmacology, Frederick P. Whiddon College of Medicine, University of South Alabama, Mobile, AL, U.S.A
- Department of Physiology and Cell Biology, Frederick P. Whiddon College of Medicine, University of South Alabama, Mobile, AL, U.S.A
- Center for Lung Biology, Frederick P. Whiddon College of Medicine, University of South Alabama, Mobile, AL, U.S.A
| | - Silas J Leavesley
- Department of Pharmacology, Frederick P. Whiddon College of Medicine, University of South Alabama, Mobile, AL, U.S.A
- Center for Lung Biology, Frederick P. Whiddon College of Medicine, University of South Alabama, Mobile, AL, U.S.A
- Chemical and Biomolecular Engineering, University of South Alabama, Mobile, AL, U.S.A
| | - C Michael Francis
- Department of Physiology and Cell Biology, Frederick P. Whiddon College of Medicine, University of South Alabama, Mobile, AL, U.S.A
- Center for Lung Biology, Frederick P. Whiddon College of Medicine, University of South Alabama, Mobile, AL, U.S.A
| | - Thomas C Rich
- Department of Pharmacology, Frederick P. Whiddon College of Medicine, University of South Alabama, Mobile, AL, U.S.A
- Center for Lung Biology, Frederick P. Whiddon College of Medicine, University of South Alabama, Mobile, AL, U.S.A
| |
Collapse
|
2
|
Lohse MJ, Bock A, Zaccolo M. G Protein-Coupled Receptor Signaling: New Insights Define Cellular Nanodomains. Annu Rev Pharmacol Toxicol 2024; 64:387-415. [PMID: 37683278 DOI: 10.1146/annurev-pharmtox-040623-115054] [Citation(s) in RCA: 5] [Impact Index Per Article: 5.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 09/10/2023]
Abstract
G protein-coupled receptors are the largest and pharmacologically most important receptor family and are involved in the regulation of most cell functions. Most of them reside exclusively at the cell surface, from where they signal via heterotrimeric G proteins to control the production of second messengers such as cAMP and IP3 as well as the activity of several ion channels. However, they may also internalize upon agonist stimulation or constitutively reside in various intracellular locations. Recent evidence indicates that their function differs depending on their precise cellular localization. This is because the signals they produce, notably cAMP and Ca2+, are mostly bound to cell proteins that significantly reduce their mobility, allowing the generation of steep concentration gradients. As a result, signals generated by the receptors remain confined to nanometer-sized domains. We propose that such nanometer-sized domains represent the basic signaling units in a cell and a new type of target for drug development.
Collapse
Affiliation(s)
- Martin J Lohse
- ISAR Bioscience Institute, Planegg/Munich, Germany;
- Rudolf Boehm Institute of Pharmacology and Toxicology, Leipzig University, Leipzig, Germany
| | - Andreas Bock
- Rudolf Boehm Institute of Pharmacology and Toxicology, Leipzig University, Leipzig, Germany
| | - Manuela Zaccolo
- Department of Physiology, Anatomy and Genetics and National Institute for Health and Care Research Oxford Biomedical Research Centre, University of Oxford, Oxford, United Kingdom;
| |
Collapse
|
3
|
Parker M, Annamdevula NS, Pleshinger D, Ijaz Z, Jalkh J, Penn R, Deshpande D, Rich TC, Leavesley SJ. Comparing Performance of Spectral Image Analysis Approaches for Detection of Cellular Signals in Time-Lapse Hyperspectral Imaging Fluorescence Excitation-Scanning Microscopy. Bioengineering (Basel) 2023; 10:642. [PMID: 37370573 PMCID: PMC10295298 DOI: 10.3390/bioengineering10060642] [Citation(s) in RCA: 2] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/24/2023] [Revised: 05/13/2023] [Accepted: 05/16/2023] [Indexed: 06/29/2023] Open
Abstract
Hyperspectral imaging (HSI) technology has been applied in a range of fields for target detection and mixture analysis. While HSI was originally developed for remote sensing applications, modern uses include agriculture, historical document authentication, and medicine. HSI has also shown great utility in fluorescence microscopy. However, traditional fluorescence microscopy HSI systems have suffered from limited signal strength due to the need to filter or disperse the emitted light across many spectral bands. We have previously demonstrated that sampling the fluorescence excitation spectrum may provide an alternative approach with improved signal strength. Here, we report on the use of excitation-scanning HSI for dynamic cell signaling studies-in this case, the study of the second messenger Ca2+. Time-lapse excitation-scanning HSI data of Ca2+ signals in human airway smooth muscle cells (HASMCs) were acquired and analyzed using four spectral analysis algorithms: linear unmixing (LU), spectral angle mapper (SAM), constrained energy minimization (CEM), and matched filter (MF), and the performances were compared. Results indicate that LU and MF provided similar linear responses to increasing Ca2+ and could both be effectively used for excitation-scanning HSI. A theoretical sensitivity framework was used to enable the filtering of analyzed images to reject pixels with signals below a minimum detectable limit. The results indicated that subtle kinetic features might be revealed through pixel filtering. Overall, the results suggest that excitation-scanning HSI can be employed for kinetic measurements of cell signals or other dynamic cellular events and that the selection of an appropriate analysis algorithm and pixel filtering may aid in the extraction of quantitative signal traces. These approaches may be especially helpful for cases where the signal of interest is masked by strong cellular autofluorescence or other competing signals.
Collapse
Affiliation(s)
- Marina Parker
- Department of Chemical and Biomolecular Engineering, University of South Alabama, 150 Student Services Dr., Mobile, AL 36688, USA
- Department of Systems Engineering, University of South Alabama, 150 Student Services Dr., Mobile, AL 36688, USA
| | - Naga S. Annamdevula
- Department of Pharmacology, University of South Alabama, 5851 USA Drive N., Mobile, AL 36688, USA; (N.S.A.)
- Center for Lung Biology, University of South Alabama, 5851 USA Drive N., Mobile, AL 36688, USA
| | - Donald Pleshinger
- Department of Pharmacology, University of South Alabama, 5851 USA Drive N., Mobile, AL 36688, USA; (N.S.A.)
- Center for Lung Biology, University of South Alabama, 5851 USA Drive N., Mobile, AL 36688, USA
| | - Zara Ijaz
- College of Medicine, University of South Alabama, 5851 USA Drive N., Mobile, AL 36688, USA
| | - Josephine Jalkh
- College of Medicine, University of South Alabama, 5851 USA Drive N., Mobile, AL 36688, USA
| | - Raymond Penn
- College of Medicine, Thomas Jefferson University, Philadelphia, PA 19107, USA
| | - Deepak Deshpande
- College of Medicine, Thomas Jefferson University, Philadelphia, PA 19107, USA
| | - Thomas C. Rich
- Department of Pharmacology, University of South Alabama, 5851 USA Drive N., Mobile, AL 36688, USA; (N.S.A.)
- Center for Lung Biology, University of South Alabama, 5851 USA Drive N., Mobile, AL 36688, USA
| | - Silas J. Leavesley
- Department of Chemical and Biomolecular Engineering, University of South Alabama, 150 Student Services Dr., Mobile, AL 36688, USA
- Department of Systems Engineering, University of South Alabama, 150 Student Services Dr., Mobile, AL 36688, USA
- Department of Pharmacology, University of South Alabama, 5851 USA Drive N., Mobile, AL 36688, USA; (N.S.A.)
- Center for Lung Biology, University of South Alabama, 5851 USA Drive N., Mobile, AL 36688, USA
| |
Collapse
|
4
|
Dunn P, Annamdevula NS, Leavesley SJ, Rich TC, Phan AV. A two-dimensional finite element model of intercellular cAMP signaling through gap junction channels. J Biomech 2023; 152:111588. [PMID: 37094384 PMCID: PMC10173664 DOI: 10.1016/j.jbiomech.2023.111588] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/16/2022] [Revised: 04/02/2023] [Accepted: 04/11/2023] [Indexed: 04/26/2023]
Abstract
While cyclic adenosine monophosphate (cAMP) is typically considered an intracellular signal, it has been shown to spread between adjacent cells through connexin-based gap junction channels, promoting gap junctional intercellular communication (GJIC). Gap junction-mediated signaling is critical for the coordinated function of many tissues, and have been linked with cardiovascular disease, neurogenerative disease, and cancers. In particular, it plays a complex role in tumor suppression or promotion. This work introduces a two-dimensional finite element model that can describe intercellular cAMP signaling in the presence of gap junctions on membrane interfaces. The model was utilized to simulate cAMP transfer through one and two gap junction channels on the interface of a cluster of two pulmonary microvascular endothelial cells. The simulation results were found to generally agree with what has been observed in the literature in terms of GJIC. The research outcomes suggest that the proposed model can be employed to evaluate the permeability properties of a gap junction channel if its cAMP volumetric flow rate can be experimentally measured.
Collapse
Affiliation(s)
- P Dunn
- Department of Mechanical, Aerospace and Biomedical Engineering, University of South Alabama, Mobile, AL 36688, USA
| | - N S Annamdevula
- Center for Lung Biology & Department of Pharmacology University of South Alabama, Mobile, AL 36688, USA
| | - S J Leavesley
- Center for Lung Biology & Department of Pharmacology University of South Alabama, Mobile, AL 36688, USA; Department of Chemical and Biomolecular Engineering, University of South Alabama, Mobile, AL 36688, USA
| | - T C Rich
- Center for Lung Biology & Department of Pharmacology University of South Alabama, Mobile, AL 36688, USA
| | - A-V Phan
- Department of Mechanical, Aerospace and Biomedical Engineering, University of South Alabama, Mobile, AL 36688, USA.
| |
Collapse
|
5
|
Rebenku I, Lloyd CB, Szöllősi J, Vereb G. Pixel-by-pixel autofluorescence corrected FRET in fluorescence microscopy improves accuracy for samples with spatially varied autofluorescence to signal ratio. Sci Rep 2023; 13:2934. [PMID: 36804608 PMCID: PMC9941493 DOI: 10.1038/s41598-023-30098-w] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/23/2022] [Accepted: 02/14/2023] [Indexed: 02/22/2023] Open
Abstract
The actual interaction between signaling species in cellular processes is often more important than their expression levels. Förster resonance energy transfer (FRET) is a popular tool for studying molecular interactions, since it is highly sensitive to proximity in the range of 2-10 nm. Spectral spillover-corrected quantitative (3-cube) FRET is a cost effective and versatile approach, which can be applied in flow cytometry and various modalities of fluorescence microscopy, but may be hampered by varying levels of autofluorescence. Here, we have implemented pixel-by-pixel autofluorescence correction in microscopy FRET measurements, exploiting cell-free calibration standards void of autofluorescence that allow the correct determination of all spectral spillover factors. We also present an ImageJ/Fiji plugin for interactive analysis of single images as well as automatic creation of quantitative FRET efficiency maps from large image sets. For validation, we used bead and cell based FRET models covering a range of signal to autofluorescence ratios and FRET efficiencies and compared the approach with conventional average autofluorescence/background correction. Pixel-by-pixel autofluorescence correction proved to be superior in the accuracy of results, particularly for samples with spatially varying autofluorescence and low fluorescence to autofluorescence ratios, the latter often being the case for physiological expression levels.
Collapse
Affiliation(s)
- István Rebenku
- grid.7122.60000 0001 1088 8582Department of Biophysics and Cell Biology, Faculty of Medicine, University of Debrecen, Egyetem tér 1, Debrecen, 4032 Hungary ,grid.7122.60000 0001 1088 8582ELKH-DE Cell Biology and Signaling Research Group, Faculty of Medicine, University of Debrecen, Egyetem tér 1, Debrecen, 4032 Hungary
| | - Cameron B. Lloyd
- grid.7122.60000 0001 1088 8582Department of Biophysics and Cell Biology, Faculty of Medicine, University of Debrecen, Egyetem tér 1, Debrecen, 4032 Hungary
| | - János Szöllősi
- grid.7122.60000 0001 1088 8582Department of Biophysics and Cell Biology, Faculty of Medicine, University of Debrecen, Egyetem tér 1, Debrecen, 4032 Hungary ,grid.7122.60000 0001 1088 8582ELKH-DE Cell Biology and Signaling Research Group, Faculty of Medicine, University of Debrecen, Egyetem tér 1, Debrecen, 4032 Hungary
| | - György Vereb
- Department of Biophysics and Cell Biology, Faculty of Medicine, University of Debrecen, Egyetem tér 1, Debrecen, 4032, Hungary. .,ELKH-DE Cell Biology and Signaling Research Group, Faculty of Medicine, University of Debrecen, Egyetem tér 1, Debrecen, 4032, Hungary. .,Faculty of Pharmacy, University of Debrecen, Egyetem tér 1, Debrecen, 4032, Hungary.
| |
Collapse
|
6
|
Knighten JM, Aziz T, Pleshinger DJ, Annamdevula N, Rich TC, Taylor MS, Andrews JF, Macarilla CT, Francis CM. Algorithm for biological second messenger analysis with dynamic regions of interest. PLoS One 2023; 18:e0284394. [PMID: 37167308 PMCID: PMC10174521 DOI: 10.1371/journal.pone.0284394] [Citation(s) in RCA: 2] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/26/2022] [Accepted: 03/29/2023] [Indexed: 05/13/2023] Open
Abstract
Physiological function is regulated through cellular communication that is facilitated by multiple signaling molecules such as second messengers. Analysis of signal dynamics obtained from cell and tissue imaging is difficult because of intricate spatially and temporally distinct signals. Signal analysis tools based on static region of interest analysis may under- or overestimate signals in relation to region of interest size and location. Therefore, we developed an algorithm for biological signal detection and analysis based on dynamic regions of interest, where time-dependent polygonal regions of interest are automatically assigned to the changing perimeter of detected and segmented signals. This approach allows signal profiles to be rigorously and precisely tracked over time, eliminating the signal distortion observed with static methods. Integration of our approach with state-of-the-art image processing and particle tracking pipelines enabled the isolation of dynamic cellular signaling events and characterization of biological signaling patterns with distinct combinations of parameters including amplitude, duration, and spatial spread. Our algorithm was validated using synthetically generated datasets and compared with other available methods. Application of the algorithm to volumetric time-lapse hyperspectral images of cyclic adenosine monophosphate measurements in rat microvascular endothelial cells revealed distinct signal heterogeneity with respect to cell depth, confirming the utility of our approach for analysis of 5-dimensional data. In human tibial arteries, our approach allowed the identification of distinct calcium signal patterns associated with atherosclerosis. Our algorithm for automated detection and analysis of second messenger signals enables the decoding of signaling patterns in diverse tissues and identification of pathologic cellular responses.
Collapse
Affiliation(s)
- Jennifer M Knighten
- Department of Physiology and Cell Biology, University of South Alabama College of Medicine, Mobile, Alabama, United States of America
| | - Takreem Aziz
- Department of Physiology and Cell Biology, University of South Alabama College of Medicine, Mobile, Alabama, United States of America
| | - Donald J Pleshinger
- Department of Pharmacology, University of South Alabama College of Medicine, Mobile, Alabama, United States of America
| | - Naga Annamdevula
- Department of Pharmacology, University of South Alabama College of Medicine, Mobile, Alabama, United States of America
| | - Thomas C Rich
- Department of Pharmacology, University of South Alabama College of Medicine, Mobile, Alabama, United States of America
- Center for Lung Biology, University of South Alabama College of Medicine, Mobile, Alabama, United States of America
| | - Mark S Taylor
- Department of Physiology and Cell Biology, University of South Alabama College of Medicine, Mobile, Alabama, United States of America
| | - Joel F Andrews
- Bioimaging Core Facility, University of South Alabama College of Medicine, Mobile, Alabama, United States of America
| | - Christian T Macarilla
- Department of Physiology and Cell Biology, University of South Alabama College of Medicine, Mobile, Alabama, United States of America
| | - C Michael Francis
- Department of Physiology and Cell Biology, University of South Alabama College of Medicine, Mobile, Alabama, United States of America
- Center for Lung Biology, University of South Alabama College of Medicine, Mobile, Alabama, United States of America
| |
Collapse
|
7
|
Henderson J, Havranek O, Ma MCJ, Herman V, Kupcova K, Chrbolkova T, Pacheco-Blanco M, Wang Z, Comer JM, Zal T, Davis RE. Detecting Förster resonance energy transfer in living cells by conventional and spectral flow cytometry. Cytometry A 2022; 101:818-834. [PMID: 34128311 DOI: 10.1002/cyto.a.24472] [Citation(s) in RCA: 6] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/21/2020] [Revised: 05/31/2021] [Accepted: 06/07/2021] [Indexed: 01/27/2023]
Abstract
Assays based on Förster resonance energy transfer (FRET) can be used to study many processes in cell biology. Although this is most often done with microscopy for fluorescence detection, we report two ways to measure FRET in living cells by flow cytometry. Using a conventional flow cytometer and the "3-cube method" for intensity-based calculation of FRET efficiency, we measured the enzymatic activity of specific kinases in cells expressing a genetically-encoded reporter. For both AKT and protein kinase A, the method measured kinase activity in time-course, dose-response, and kinetic assays. Using the Cytek Aurora spectral flow cytometer, which applies linear unmixing to emission measured in multiple wavelength ranges, FRET from the same reporters was measured with greater single-cell precision, in real time and in the presence of other fluorophores. Results from gene-knockout studies suggested that spectral flow cytometry might enable the sorting of cells on the basis of FRET. The methods we present provide convenient and flexible options for using FRET with flow cytometry in studies of cell biology.
Collapse
Affiliation(s)
- Jared Henderson
- Department of Lymphoma and Myeloma, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA
| | - Ondrej Havranek
- Department of Lymphoma and Myeloma, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA.,BIOCEV, First Faculty of Medicine, Charles University, Vestec, Czech Republic.,Department of Hematology, Charles University and General University Hospital, Prague, Czech Republic
| | - Man Chun John Ma
- Department of Lymphoma and Myeloma, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA
| | - Vaclav Herman
- BIOCEV, First Faculty of Medicine, Charles University, Vestec, Czech Republic.,Department of Hematology, Charles University and General University Hospital, Prague, Czech Republic
| | - Kristyna Kupcova
- BIOCEV, First Faculty of Medicine, Charles University, Vestec, Czech Republic
| | - Tereza Chrbolkova
- BIOCEV, First Faculty of Medicine, Charles University, Vestec, Czech Republic
| | | | - Zhiqiang Wang
- Department of Lymphoma and Myeloma, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA
| | - Justin M Comer
- Department of Lymphoma and Myeloma, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA
| | - Tomasz Zal
- Department of Leukemia, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA
| | - Richard Eric Davis
- Department of Lymphoma and Myeloma, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA.,Department of Translational Molecular Pathology, The University of Texas-MD Anderson Cancer Center, Houston, Texas, USA
| |
Collapse
|
8
|
Browning CM, Mayes S, Mayes SA, Rich TC, Leavesley SJ. Microscopy is better in color: development of a streamlined spectral light path for real-time multiplex fluorescence microscopy. BIOMEDICAL OPTICS EXPRESS 2022; 13:3751-3772. [PMID: 35991911 PMCID: PMC9352297 DOI: 10.1364/boe.453657] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Received: 02/09/2022] [Revised: 05/24/2022] [Accepted: 05/26/2022] [Indexed: 06/15/2023]
Abstract
Spectroscopic image data has provided molecular discrimination for numerous fields including: remote sensing, food safety and biomedical imaging. Despite the various technologies for acquiring spectral data, there remains a trade-off when acquiring data. Typically, spectral imaging either requires long acquisition times to collect an image stack with high spectral specificity or acquisition times are shortened at the expense of fewer spectral bands or reduced spatial sampling. Hence, new spectral imaging microscope platforms are needed to help mitigate these limitations. Fluorescence excitation-scanning spectral imaging is one such new technology, which allows more of the emitted signal to be detected than comparable emission-scanning spectral imaging systems. Here, we have developed a new optical geometry that provides spectral illumination for use in excitation-scanning spectral imaging microscope systems. This was accomplished using a wavelength-specific LED array to acquire spectral image data. Feasibility of the LED-based spectral illuminator was evaluated through simulation and benchtop testing and assessment of imaging performance when integrated with a widefield fluorescence microscope. Ray tracing simulations (TracePro) were used to determine optimal optical component selection and geometry. Spectral imaging feasibility was evaluated using a series of 6-label fluorescent slides. The LED-based system response was compared to a previously tested thin-film tunable filter (TFTF)-based system. Spectral unmixing successfully discriminated all fluorescent components in spectral image data acquired from both the LED and TFTF systems. Therefore, the LED-based spectral illuminator provided spectral image data sets with comparable information content so as to allow identification of each fluorescent component. These results provide proof-of-principle demonstration of the ability to combine output from many discrete wavelength LED sources using a double-mirror (Cassegrain style) optical configuration that can be further modified to allow for high speed, video-rate spectral image acquisition. Real-time spectral fluorescence microscopy would allow monitoring of rapid cell signaling processes (i.e., Ca2+ and other second messenger signaling) and has potential to be translated to clinical imaging platforms.
Collapse
Affiliation(s)
- Craig M. Browning
- Chemical and Biomolecular Engineering, University of South Alabama, AL 36688, USA
- Systems Engineering, University of South Alabama, AL 36688, USA
- These authors contributed equally to this work
| | - Samantha Mayes
- Chemical and Biomolecular Engineering, University of South Alabama, AL 36688, USA
- These authors contributed equally to this work
| | - Samuel A. Mayes
- Chemical and Biomolecular Engineering, University of South Alabama, AL 36688, USA
- Systems Engineering, University of South Alabama, AL 36688, USA
| | - Thomas C. Rich
- Pharmacology, University of South Alabama, AL 36688, USA
- Center for Lung Biology, University of South Alabama, AL 36688, USA
| | - Silas J. Leavesley
- Chemical and Biomolecular Engineering, University of South Alabama, AL 36688, USA
- Pharmacology, University of South Alabama, AL 36688, USA
- Center for Lung Biology, University of South Alabama, AL 36688, USA
| |
Collapse
|
9
|
Bene L, Damjanovich L. Spectral flow cytometric FRET: Towards a hyper dimensional flow cytometry. Cytometry A 2022; 101:468-473. [PMID: 35484961 DOI: 10.1002/cyto.a.24561] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/07/2022] [Revised: 04/11/2022] [Accepted: 04/19/2022] [Indexed: 11/07/2022]
Affiliation(s)
- László Bene
- Department of Surgery, Faculty of Medicine, University of Debrecen, Debrecen, Hungary
| | - László Damjanovich
- Department of Surgery, Faculty of Medicine, University of Debrecen, Debrecen, Hungary
| |
Collapse
|
10
|
Leavesley SJ, Annamdevula N, Johnson S, Pleshinger DJ, Rich TC. Automated Image Analysis of FRET Signals for Subcellular cAMP Quantification. Methods Mol Biol 2022; 2483:167-180. [PMID: 35286675 DOI: 10.1007/978-1-0716-2245-2_10] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 06/14/2023]
Abstract
A variety of FRET probes have been developed to examine cAMP localization and dynamics in single cells. These probes offer a readily accessible approach to measure localized cAMP signals. However, given the low signal-to-noise ratio of most FRET probes and the dynamic nature of the intracellular environment, there have been marked limitations in the ability to use FRET probes to study localized signaling events within the same cell. Here, we outline a methodology to dissect kinetics of cAMP-mediated FRET signals in single cells using automated image analysis approaches. We additionally extend these approaches to the analysis of subcellular regions. These approaches offer a unique opportunity to assess localized cAMP kinetics in an unbiased, quantitative fashion.
Collapse
Affiliation(s)
- Silas J Leavesley
- Department of Chemical and Biomolecular Engineering, University of South Alabama, Mobile, AL, USA.
- Department of Pharmacology, University of South Alabama, Mobile, AL, USA.
- Center for Lung Biology, University of South Alabama, Mobile, AL, USA.
| | - Naga Annamdevula
- Department of Pharmacology, University of South Alabama, Mobile, AL, USA
- Center for Lung Biology, University of South Alabama, Mobile, AL, USA
- Department of Physiology, University of South Alabama, Mobile, AL, USA
| | - Santina Johnson
- Department of Pharmacology, University of South Alabama, Mobile, AL, USA
- Center for Lung Biology, University of South Alabama, Mobile, AL, USA
| | - D J Pleshinger
- Department of Pharmacology, University of South Alabama, Mobile, AL, USA
- Center for Lung Biology, University of South Alabama, Mobile, AL, USA
| | - Thomas C Rich
- Department of Pharmacology, University of South Alabama, Mobile, AL, USA
- Center for Lung Biology, University of South Alabama, Mobile, AL, USA
- Department of Physiology, University of South Alabama, Mobile, AL, USA
- College of Engineering, University of South Alabama, Mobile, AL, USA
| |
Collapse
|
11
|
Warren R, Rich T, Leavesley S, Phan AV. A three-dimensional finite element model of cAMP signals. FORCES IN MECHANICS 2021; 4. [PMID: 35072121 PMCID: PMC8773462 DOI: 10.1016/j.finmec.2021.100041] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Grants] [Track Full Text] [Download PDF] [Figures] [Subscribe] [Scholar Register] [Indexed: 11/19/2022]
Affiliation(s)
- R. Warren
- Department of Mechanical, Aerospace and Biomedical Engineering, University of South Alabama, Mobile, AL 36688, USA
| | - T.C. Rich
- Center for Lung Biology & Department of Pharmacology, University of South Alabama, Mobile, AL 36688, USA
| | - S.J. Leavesley
- Center for Lung Biology & Department of Pharmacology, University of South Alabama, Mobile, AL 36688, USA
- Department of Chemical and Biomolecular Engineering, University of South Alabama, Mobile, AL 36688, USA
| | - A.-V. Phan
- Department of Mechanical, Aerospace and Biomedical Engineering, University of South Alabama, Mobile, AL 36688, USA
- Corresponding author. (A.-V. Phan)
| |
Collapse
|
12
|
cAMP Compartmentalization in Cerebrovascular Endothelial Cells: New Therapeutic Opportunities in Alzheimer's Disease. Cells 2021; 10:cells10081951. [PMID: 34440720 PMCID: PMC8392343 DOI: 10.3390/cells10081951] [Citation(s) in RCA: 15] [Impact Index Per Article: 5.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/01/2021] [Revised: 07/19/2021] [Accepted: 07/28/2021] [Indexed: 12/20/2022] Open
Abstract
The vascular hypothesis used to explain the pathophysiology of Alzheimer’s disease (AD) suggests that a dysfunction of the cerebral microvasculature could be the beginning of alterations that ultimately leads to neuronal damage, and an abnormal increase of the blood–brain barrier (BBB) permeability plays a prominent role in this process. It is generally accepted that, in physiological conditions, cyclic AMP (cAMP) plays a key role in maintaining BBB permeability by regulating the formation of tight junctions between endothelial cells of the brain microvasculature. It is also known that intracellular cAMP signaling is highly compartmentalized into small nanodomains and localized cAMP changes are sufficient at modifying the permeability of the endothelial barrier. This spatial and temporal distribution is maintained by the enzymes involved in cAMP synthesis and degradation, by the location of its effectors, and by the existence of anchor proteins, as well as by buffers or different cytoplasm viscosities and intracellular structures limiting its diffusion. This review compiles current knowledge on the influence of cAMP compartmentalization on the endothelial barrier and, more specifically, on the BBB, laying the foundation for a new therapeutic approach in the treatment of AD.
Collapse
|
13
|
Margarido AS, Bornes L, Vennin C, van Rheenen J. Cellular Plasticity during Metastasis: New Insights Provided by Intravital Microscopy. Cold Spring Harb Perspect Med 2020; 10:cshperspect.a037267. [PMID: 31615867 DOI: 10.1101/cshperspect.a037267] [Citation(s) in RCA: 9] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/10/2023]
Abstract
Metastasis is a highly dynamic process during which cancer and microenvironmental cells undergo a cascade of events required for efficient dissemination throughout the body. During the metastatic cascade, tumor cells can change their state and behavior, a phenomenon commonly defined as cellular plasticity. To monitor cellular plasticity during metastasis, high-resolution intravital microscopy (IVM) techniques have been developed and allow us to visualize individual cells by repeated imaging in animal models. In this review, we summarize the latest technological advancements in the field of IVM and how they have been applied to monitor metastatic events. In particular, we highlight how longitudinal imaging in native tissues can provide new insights into the plastic physiological and developmental processes that are hijacked by cancer cells during metastasis.
Collapse
Affiliation(s)
- Andreia S Margarido
- Molecular Pathology, Oncode Institute, The Netherlands Cancer Institute, 1066CX Amsterdam, The Netherlands
| | - Laura Bornes
- Molecular Pathology, Oncode Institute, The Netherlands Cancer Institute, 1066CX Amsterdam, The Netherlands
| | - Claire Vennin
- Molecular Pathology, Oncode Institute, The Netherlands Cancer Institute, 1066CX Amsterdam, The Netherlands
| | - Jacco van Rheenen
- Molecular Pathology, Oncode Institute, The Netherlands Cancer Institute, 1066CX Amsterdam, The Netherlands
| |
Collapse
|
14
|
Annamdevula NS, Sweat R, Gunn H, Griswold JR, Britain AL, Rich TC, Leavesley SJ. Measurement of 3-Dimensional cAMP Distributions in Living Cells using 4-Dimensional (x, y, z, and λ) Hyperspectral FRET Imaging and Analysis. J Vis Exp 2020. [PMID: 33191928 DOI: 10.3791/61720] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/19/2023] Open
Abstract
Cyclic AMP is a second messenger that is involved in a wide range of cellular and physiological activities. Several studies suggest that cAMP signals are compartmentalized, and that compartmentalization contributes to signaling specificity within the cAMP signaling pathway. The development of Fӧrster resonance energy transfer (FRET) based biosensors has furthered the ability to measure and visualize cAMP signals in cells. However, these measurements are often confined to two spatial dimensions, which may result in misinterpretation of data. To date, there have been only very limited measurements of cAMP signals in three spatial dimensions (x, y, and z), due to the technical limitations in using FRET sensors that inherently exhibit low signal to noise ratio (SNR). In addition, traditional filter-based imaging approaches are often ineffective for accurate measurement of cAMP signals in localized subcellular regions due to a range of factors, including spectral crosstalk, limited signal strength, and autofluorescence. To overcome these limitations and allow FRET-based biosensors to be used with multiple fluorophores, we have developed hyperspectral FRET imaging and analysis approaches that provide spectral specificity for calculating FRET efficiencies and the ability to spectrally separate FRET signals from confounding autofluorescence and/or signals from additional fluorescent labels. Here, we present the methodology for implementing hyperspectral FRET imaging as well as the need to construct an appropriate spectral library that is neither undersampled nor oversampled to perform spectral unmixing. While we present this methodology for measurement of three-dimensional cAMP distributions in pulmonary microvascular endothelial cells (PMVECs), this methodology could be used to study spatial distributions of cAMP in a range of cell types.
Collapse
Affiliation(s)
- Naga S Annamdevula
- Department of Pharmacology, University of South Alabama; Center for Lung Biology, University of South Alabama
| | - Rachel Sweat
- Department of Chemical and Biomolecular Engineering, University of South Alabama
| | - Hayden Gunn
- Department of Pharmacology, University of South Alabama
| | - John R Griswold
- Department of Chemical and Biomolecular Engineering, University of South Alabama
| | - Andrea L Britain
- Department of Pharmacology, University of South Alabama; Center for Lung Biology, University of South Alabama
| | - Thomas C Rich
- Department of Pharmacology, University of South Alabama; Center for Lung Biology, University of South Alabama
| | - Silas J Leavesley
- Department of Pharmacology, University of South Alabama; Center for Lung Biology, University of South Alabama; Department of Chemical and Biomolecular Engineering, University of South Alabama;
| |
Collapse
|
15
|
Deal J, Pleshinger DJ, Johnson SC, Leavesley SJ, Rich TC. Milestones in the development and implementation of FRET-based sensors of intracellular signals: A biological perspective of the history of FRET. Cell Signal 2020; 75:109769. [PMID: 32898611 DOI: 10.1016/j.cellsig.2020.109769] [Citation(s) in RCA: 12] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/12/2020] [Revised: 08/28/2020] [Accepted: 08/31/2020] [Indexed: 01/24/2023]
Abstract
Fӧrster resonance energy transfer (FRET) has been described for more than a century. FRET has become a mainstay for the study of protein localization in living cells and tissues. It has also become widely used in the fields that comprise cellular signaling. FRET-based probes have been developed to monitor second messenger signals, the phosphorylation state of peptides and proteins, and subsequent cellular responses. Here, we discuss the milestones that led to FRET becoming a widely used tool for the study of biological systems: the theoretical description of FRET, the insight to use FRET as a molecular ruler, and the isolation and genetic modification of green fluorescent protein (GFP). Each of these milestones were critical to the development of a myriad of FRET-based probes and reporters in common use today. FRET-probes offer a unique opportunity to interrogate second messenger signals and subsequent protein phosphorylation - and perhaps the most effective approach for study of cAMP/PKA pathways. As such, FRET probes are widely used in the study of intracellular signaling pathways. Yet, somehow, the potential of FRET-based probes to provide windows through which we can visualize complex cellular signaling systems has not been fully reached. Hence we conclude by discussing the technical challenges to be overcome if FRET-based probes are to live up to their potential for the study of complex signaling networks.
Collapse
Affiliation(s)
- J Deal
- Basic Medical Sciences Graduate Program, University of South Alabama, Mobile, AL 36688, USA; Center for Lung Biology, Departments of Biomolecular Engineering, University of South Alabama, Mobile, AL 36688, USA
| | - D J Pleshinger
- Center for Lung Biology, Departments of Biomolecular Engineering, University of South Alabama, Mobile, AL 36688, USA; Pharmacology and Biomolecular Engineering, University of South Alabama, Mobile, AL 36688, USA
| | - S C Johnson
- Basic Medical Sciences Graduate Program, University of South Alabama, Mobile, AL 36688, USA; Pharmacology and Biomolecular Engineering, University of South Alabama, Mobile, AL 36688, USA
| | - S J Leavesley
- Basic Medical Sciences Graduate Program, University of South Alabama, Mobile, AL 36688, USA; Center for Lung Biology, Departments of Biomolecular Engineering, University of South Alabama, Mobile, AL 36688, USA; Pharmacology and Biomolecular Engineering, University of South Alabama, Mobile, AL 36688, USA; Chemical and Biomolecular Engineering, University of South Alabama, Mobile, AL 36688, USA
| | - T C Rich
- Basic Medical Sciences Graduate Program, University of South Alabama, Mobile, AL 36688, USA; Center for Lung Biology, Departments of Biomolecular Engineering, University of South Alabama, Mobile, AL 36688, USA; Pharmacology and Biomolecular Engineering, University of South Alabama, Mobile, AL 36688, USA.
| |
Collapse
|
16
|
Szabó Á, Nagy P. I Am the Alpha and the …Gamma, and the G. Calibration of Intensity-Based FRET Measurements. Cytometry A 2020; 99:369-371. [PMID: 32790096 DOI: 10.1002/cyto.a.24206] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/24/2020] [Revised: 08/05/2020] [Accepted: 08/10/2020] [Indexed: 12/22/2022]
Affiliation(s)
- Ágnes Szabó
- Department of Biophysics and Cell Biology, Faculty of Medicine, University of Debrecen, Debrecen, Hungary.,MTA-DE Cell Biology and Signaling Research Group, Faculty of Medicine, University of Debrecen, Debrecen, Hungary
| | - Peter Nagy
- Department of Biophysics and Cell Biology, Faculty of Medicine, University of Debrecen, Debrecen, Hungary
| |
Collapse
|
17
|
Deal J, Annamdevula N, Pleshinger DJ, Griswold JR, Odom A, Tayara A, Lall M, Browning C, Parker M, Rich TC, Leavesley SJ. Comparison of spectral FRET microscopy approaches for single-cell analysis. PROCEEDINGS OF SPIE--THE INTERNATIONAL SOCIETY FOR OPTICAL ENGINEERING 2020; 11243:112430Y. [PMID: 34035557 PMCID: PMC8142325 DOI: 10.1117/12.2546308] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 11/14/2022]
Abstract
Förster resonance energy transfer (FRET) is a valuable tool for measuring molecular distances and the effects of biological processes such as cyclic nucleotide messenger signaling and protein localization. Most FRET techniques require two fluorescent proteins with overlapping excitation/emission spectral pairing to maximize detection sensitivity and FRET efficiency. FRET microscopy often utilizes differing peak intensities of the selected fluorophores measured through different optical filter sets to estimate the FRET index or efficiency. Microscopy platforms used to make these measurements include wide-field, laser scanning confocal, and fluorescence lifetime imaging. Each platform has associated advantages and disadvantages, such as speed, sensitivity, specificity, out-of-focus fluorescence, and Z-resolution. In this study, we report comparisons among multiple microscopy and spectral filtering platforms such as standard 2-filter FRET, emission-scanning hyperspectral imaging, and excitation-scanning hyperspectral imaging. Samples of human embryonic kidney (HEK293) cells were grown on laminin-coated 28 mm round gridded glass coverslips (10816, Ibidi, Fitchburg, Wisconsin) and transfected with adenovirus encoding a cAMP-sensing FRET probe composed of a FRET donor (Turquoise) and acceptor (Venus). Additionally, 3 FRET "controls" with fixed linker lengths between Turquoise and Venus proteins were used for inter-platform validation. Grid locations were logged, recorded with light micrographs, and used to ensure that whole-cell FRET was compared on a cell-by-cell basis among the different microscopy platforms. FRET efficiencies were also calculated and compared for each method. Preliminary results indicate that hyperspectral methods increase the signal-to-noise ratio compared to a standard 2-filter approach.
Collapse
Affiliation(s)
- Joshua Deal
- Department of Chemical & Biomolecular Engineering, University of South Alabama
- Center for Lung Biology, University of South Alabama
- Department of Pharmacology, University of South Alabama
| | - Naga Annamdevula
- Center for Lung Biology, University of South Alabama
- Department of Pharmacology, University of South Alabama
| | - Donald John Pleshinger
- Center for Lung Biology, University of South Alabama
- Department of Pharmacology, University of South Alabama
| | | | - Aliyah Odom
- Department of Chemical & Biomolecular Engineering, University of South Alabama
| | - Alia Tayara
- Department of Chemical & Biomolecular Engineering, University of South Alabama
| | - Malvika Lall
- College of Medicine, University of South Alabama
| | - Craig Browning
- Department of Chemical & Biomolecular Engineering, University of South Alabama
- Systems Engineering, University of South Alabama
| | - Marina Parker
- Department of Chemical & Biomolecular Engineering, University of South Alabama
- Systems Engineering, University of South Alabama
| | - Thomas C Rich
- Center for Lung Biology, University of South Alabama
- Department of Pharmacology, University of South Alabama
| | - Silas J Leavesley
- Department of Chemical & Biomolecular Engineering, University of South Alabama
- Center for Lung Biology, University of South Alabama
- Department of Pharmacology, University of South Alabama
| |
Collapse
|
18
|
Stone N, Shettlesworth S, Rich TC, Leavesley SJ, Phan AV. A two-dimensional finite element model of cyclic adenosine monophosphate (cAMP) intracellular signaling. SN APPLIED SCIENCES 2019; 1. [PMID: 33615142 DOI: 10.1007/s42452-019-1757-9] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 10/25/2022] Open
Abstract
In this work, we present a two-dimensional finite element analysis (FEA) model that describes fundamental intracellular signals of cyclic adenosine monophosphate (cAMP) in a general fashion. The model was subsequently solved numerically and the results were displayed in forms of time-course plots of cAMP concentration at a cellular location or color-filled contour maps of cAMP signal distribution within the cell at specific time points. Basic intracellular cAMP signaling was described in this model so it can be numerically validated by verifying its numerical results against available analytical solutions and against results obtained from other numerical techniques reported in the literature. This is the first important step before the model can be expanded in future work. Model simulations demonstrate that under certain conditions, sustained cAMP concentrations can be formed within endothelial cells (ECs), similar to those observed in rat pulmonary microvascular ECs. Spatial and temporal cAMP dynamic simulations indicated that the proposed FEA model is an effective tool for the study of the kinetics and spatial spread of second messenger signaling and can be expanded to simulate second messenger signals in the pulmonary vasculature.
Collapse
Affiliation(s)
- N Stone
- William B. Burnsed, Jr. Department of Mechanical Engineering, University of South Alabama, Mobile, AL 36688, USA
| | - S Shettlesworth
- William B. Burnsed, Jr. Department of Mechanical Engineering, University of South Alabama, Mobile, AL 36688, USA
| | - T C Rich
- Center for Lung Biology & Department of Pharmacology, University of South Alabama, Mobile, AL 36688, USA
| | - S J Leavesley
- Department of Chemical and Biomolecular Engineering & Department of Pharmacology, University of South Alabama, Mobile, AL 36688, USA
| | - A-V Phan
- William B. Burnsed, Jr. Department of Mechanical Engineering, University of South Alabama, Mobile, AL 36688, USA
| |
Collapse
|
19
|
Deal J, Britain A, Rich T, Leavesley S. Excitation-Scanning Hyperspectral Imaging Microscopy to Efficiently Discriminate Fluorescence Signals. J Vis Exp 2019:10.3791/59448. [PMID: 31498305 PMCID: PMC6800214 DOI: 10.3791/59448] [Citation(s) in RCA: 7] [Impact Index Per Article: 1.4] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/15/2023] Open
Abstract
Several techniques rely on detection of fluorescence signals to identify or study phenomena or to elucidate functions. Separation of these fluorescence signals were proven cumbersome until the advent of hyperspectral imaging, in which fluorescence sources can be separated from each other as well as from background signals and autofluorescence (given knowledge of their spectral signatures). However, traditional, emission-scanning hyperspectral imaging suffers from slow acquisition times and low signal-to-noise ratios due to the necessary filtering of both excitation and emission light. It has been previously shown that excitation-scanning hyperspectral imaging reduces the necessary acquisition time while simultaneously increasing the signal-to-noise ratio of acquired data. Using commercially available equipment, this protocol describes how to assemble, calibrate, and use an excitation-scanning hyperspectral imaging microscopy system for separation of signals from several fluorescence sources in a single sample. While highly applicable to microscopic imaging of cells and tissues, this technique may also be useful for any type of experiment utilizing fluorescence in which it is possible to vary excitation wavelengths, including but not limited to: chemical imaging, environmental applications, eye care, food science, forensic science, medical science, and mineralogy.
Collapse
Affiliation(s)
- Joshua Deal
- Department of Chemical and Biomolecular Engineering, University of South Alabama; Center for Lung Biology, University of South Alabama; Department of Pharmacology, University of South Alabama
| | - Andrea Britain
- Center for Lung Biology, University of South Alabama; Department of Pharmacology, University of South Alabama
| | - Thomas Rich
- Center for Lung Biology, University of South Alabama; Department of Pharmacology, University of South Alabama
| | - Silas Leavesley
- Department of Chemical and Biomolecular Engineering, University of South Alabama; Center for Lung Biology, University of South Alabama; Department of Pharmacology, University of South Alabama;
| |
Collapse
|
20
|
Bene L, Damjanovich L. Förster Resonance Energy Transfer Pioneers Biomechanics: Molecular-Scale Force Meters for Visualizing Intracellular Stress Fields. Cytometry A 2019; 95:819-822. [PMID: 31034713 DOI: 10.1002/cyto.a.23780] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.4] [Reference Citation Analysis] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/26/2019] [Revised: 04/08/2019] [Accepted: 04/13/2019] [Indexed: 12/29/2022]
Affiliation(s)
- László Bene
- Department of Surgery, Faculty of Medicine, University of Debrecen, Debrecen, Hungary
| | - László Damjanovich
- Department of Surgery, Faculty of Medicine, University of Debrecen, Debrecen, Hungary
| |
Collapse
|
21
|
Rich TC, Griswold JR, Deal J, Annamdevula N, McAlister K, Mayes S, Browning C, Parker M, Leavelsey SJ. Hyperspectral imaging microscopy for measurement of localized second messenger signals in single cells. PROCEEDINGS OF SPIE--THE INTERNATIONAL SOCIETY FOR OPTICAL ENGINEERING 2019; 10881:108811F. [PMID: 34045781 PMCID: PMC8151147 DOI: 10.1117/12.2508052] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 11/14/2022]
Abstract
Ca2+ and cAMP are ubiquitous second messengers known to differentially regulate a variety of cellular functions over a wide range of timescales. Studies from a variety of groups support the hypothesis that these signals can be localized to discrete locations within cells, and that this subcellular localization is a critical component of signaling specificity. However, to date, it has been difficult to track second messenger signals at multiple locations within a single cell. This difficulty is largely due to the inability to measure multiplexed florescence signals in real time. To overcome this limitation, we have utilized both emission scan- and excitation scan-based hyperspectral imaging approaches to track second messenger signals as well as labeled cellular structures and/or proteins in the same cell. We have previously reported that hyperspectral imaging techniques improve the signal-to-noise ratios of both fluorescence and FRET measurements, and are thus well suited for the measurement of localized second messenger signals. Using these approaches, we have measured near plasma membrane and near nuclear membrane cAMP signals, as well as distributed signals within the cytosol, in several cell types including airway smooth muscle, pulmonary endothelial, and HEK-293 cells. We have also measured cAMP and Ca2+ signals near autofluorescent structures that appear to be golgi. Our data demonstrate that hyperspectral imaging approaches provide unique insight into the spatial and kinetic distributions of cAMP and Ca2+ signals in single cells.
Collapse
Affiliation(s)
- Thomas C Rich
- Pharmacology, University of South Alabama, AL 36688
- Center for Lung Biology, University of South Alabama, AL 36688
| | - J R Griswold
- Chemical and Biomolecular Engineering, University of South Alabama, AL 36688
| | - Joshua Deal
- Pharmacology, University of South Alabama, AL 36688
- Center for Lung Biology, University of South Alabama, AL 36688
| | - Naga Annamdevula
- Pharmacology, University of South Alabama, AL 36688
- Center for Lung Biology, University of South Alabama, AL 36688
| | | | - Samuel Mayes
- Chemical and Biomolecular Engineering, University of South Alabama, AL 36688
| | - Craig Browning
- Chemical and Biomolecular Engineering, University of South Alabama, AL 36688
| | - Marina Parker
- Chemical and Biomolecular Engineering, University of South Alabama, AL 36688
| | - Silas J Leavelsey
- Pharmacology, University of South Alabama, AL 36688
- Center for Lung Biology, University of South Alabama, AL 36688
- Chemical and Biomolecular Engineering, University of South Alabama, AL 36688
| |
Collapse
|
22
|
Deal J, Rich TC, Leavesley SJ. Optimizing channel selection for excitation-scanning hyperspectral imaging. PROCEEDINGS OF SPIE--THE INTERNATIONAL SOCIETY FOR OPTICAL ENGINEERING 2019; 10881:108811B. [PMID: 34045784 PMCID: PMC8151237 DOI: 10.1117/12.2510784] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/12/2023]
Abstract
A major benefit of fluorescence microscopy is the now plentiful selection of fluorescent markers. These labels can be chosen to serve complementary functions, such as tracking labeled subcellular molecules near demarcated organelles. However, with the standard 3 or 4 emission channels, multiple label detection is restricted to segregated regions of the electromagnetic spectrum, as in RGB coloring. Hyperspectral imaging allows the user to discern many fluorescence labels by their unique spectral properties, provided there is significant differentiation of their emission spectra. The cost of this technique is often an increase in gain or exposure time to accommodate the signal reduction from separating the signal into many discrete excitation or emission channels. Recent advances in hyperspectral imaging have allowed the acquisition of more signal in a shorter time period by scanning the excitation spectra of fluorophores. Here, we explore the selection of optimal channels for both significant signal separation and sufficient signal detection using excitation-scanning hyperspectral imaging. Excitation spectra were obtained using a custom inverted microscope (TE-2000, Nikon Instruments) with a Xe arc lamp and thin film tunable filter array (VersaChrome, Semrock, Inc.) Tunable filters had bandwidths between 13 and 17 nm. Scans utilized excitation wavelengths between 340 nm and 550 nm. Hyperspectral image stacks were generated and analyzed using ENVI and custom MATLAB scripts. Among channel consideration criteria were: number of channels, spectral range of scan, spacing of center wavelengths, and acquisition time.
Collapse
Affiliation(s)
- Joshua Deal
- Department of Chemical & Biomolecular Engineering, University of South Alabama
- Center for Lung Biology, University of South Alabama
- Department of Pharmacology, University of South Alabama
| | - Thomas C Rich
- Center for Lung Biology, University of South Alabama
- Department of Pharmacology, University of South Alabama
| | - Silas J Leavesley
- Department of Chemical & Biomolecular Engineering, University of South Alabama
- Center for Lung Biology, University of South Alabama
- Department of Pharmacology, University of South Alabama
| |
Collapse
|
23
|
Leavesley SJ, Griswold JR, Deal J, McAlister K, Mayes S, Browning C, Parker M, Mayes SG, Rich TC. Hyperspectral imaging fluorescence excitation scanning (HIFEX) microscopy for live cell imaging. PROCEEDINGS OF SPIE--THE INTERNATIONAL SOCIETY FOR OPTICAL ENGINEERING 2019; 10883:108831A. [PMID: 34045785 PMCID: PMC8151178 DOI: 10.1117/12.2510562] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 11/14/2022]
Abstract
In the past two decades, spectral imaging technologies have expanded the capacity of fluorescence microscopy for accurate detection of multiple labels, separation of labels from cellular and tissue autofluorescence, and analysis of autofluorescence signatures. These technologies have been implemented using a range of optical techniques, such as tunable filters, diffraction gratings, prisms, interferometry, and custom Bayer filters. Each of these techniques has associated strengths and weaknesses with regard to spectral resolution, spatial resolution, temporal resolution, and signal-to-noise characteristics. We have previously shown that spectral scanning of the fluorescence excitation spectrum can provide greatly increased signal strength compared to traditional emission-scanning approaches. Here, we present results from utilizing a Hyperspectral Imaging Fluorescence Excitation Scanning (HIFEX) microscope system for live cell imaging. Live cell signaling studies were performed using HEK 293 and rat pulmonary microvascular endothelial cells (PMVECs), transfected with either a cAMP FRET reporter or a Ca2+ reporter. Cells were further labeled to visualize subcellular structures (nuclei, membrane, mitochondria, etc.). Spectral images were acquired using a custom inverted microscope (TE2000, Nikon Instruments) equipped with a 300W Xe arc lamp and tunable excitation filter (VF-5, Sutter Instrument Co., equipped with VersaChrome filters, Semrock), and run through MicroManager. Timelapse spectral images were acquired from 350-550 nm, in 5 nm increments. Spectral image data were linearly unmixed using custom MATLAB scripts. Results indicate that the HIFEX microscope system can acquire live cell image data at acquisition speeds of 8 ms/wavelength band with minimal photobleaching, sufficient for studying moderate speed cAMP and Ca2+ events.
Collapse
Affiliation(s)
- Silas J Leavesley
- Department of Chemical and Biomolecular Engineering, University of South Alabama
- Department of Pharmacology, University of South Alabama
- Center for Lung Biology, University of South Alabama
| | - John Robert Griswold
- Department of Chemical and Biomolecular Engineering, University of South Alabama
| | - Joshua Deal
- Department of Pharmacology, University of South Alabama
| | | | - Sam Mayes
- Department of Chemical and Biomolecular Engineering, University of South Alabama
- Systems Engineering, University of South Alabama
| | - Craig Browning
- Department of Chemical and Biomolecular Engineering, University of South Alabama
- Systems Engineering, University of South Alabama
| | - Marina Parker
- Department of Chemical and Biomolecular Engineering, University of South Alabama
- Systems Engineering, University of South Alabama
| | - Samantha Gunn Mayes
- Department of Chemical and Biomolecular Engineering, University of South Alabama
| | - Thomas C Rich
- Department of Pharmacology, University of South Alabama
- Center for Lung Biology, University of South Alabama
| |
Collapse
|