Measurement of agonist-induced Ca2+ signals in human airway smooth muscle cells using excitation scan-based hyperspectral imaging and image analysis approaches.

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. To overcome this limitation, we utilized 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 measurements, and are thus well suited for the measurement of localized Ca2+ signals. We investigated the spatial spread and intensities of agonist-induced Ca2+ signals in primary human airway smooth muscle cells (HASMCs) using the Ca2+ indicator Cal520. We measured responses triggered by three agonists, carbachol, histamine, and chloroquine. We utilized custom software coded in MATLAB and Python to assess agonist induced changes in Ca2+ levels. Software algorithms removed the background and applied correction coefficients to spectral data prior to linear unmixing, spatial and temporal filtering, adaptive thresholding, and automated region of interest (ROI) detection. All three agonists triggered transient Ca2+ responses that were spatially and temporally complex. We are currently analyzing differences in both ROI area and intensity distributions triggered by these agonists. This work was supported by NIH awards P01HL066299, K25HL136869, and R01HL137030 and NSF award MRI1725937.

Extracellular vesicle-induced cyclic AMP signaling.

Second messenger signaling is required for cellular processes. We previously reported that extracellular vesicles (EVs) from stimulated cultured endothelial cells contain the biochemical second messenger, cAMP. In the current study, we sought to determine whether cAMP-enriched EVs induce second messenger signaling pathways in naïve recipient cells. Our results indicate that cAMP-enriched EVs increase cAMP content sufficient to stimulate PKA activity. The implications of our work are that EVs represent a novel intercellular mechanism for second messenger, specifically cAMP, signaling.

Measurement of 3-Dimensional cAMP Distributions in Living Cells using 4-Dimensional (x, y, z, and λ) Hyperspectral FRET Imaging and Analysis.

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 FÓ§rster 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.

Spectral imaging of FRET-based sensors reveals sustained cAMP gradients in three spatial dimensions.

Cyclic AMP is a ubiquitous second messenger that orchestrates a variety of cellular functions over different timescales. The mechanisms underlying specificity within this signaling pathway are still not well understood. Several lines of evidence suggest the existence of spatial cAMP gradients within cells, and that compartmentalization underlies specificity within the cAMP signaling pathway. However, to date, no studies have visualized cAMP gradients in three spatial dimensions (3D: x, y, z).This is in part due to the limitations of FRET-based cAMP sensors, specifically the low signal-to-noise ratio intrinsic to all intracellular FRET probes. Here, we overcome this limitation, at least in part, by implementing spectral imaging approaches to estimate FRET efficiency when multiple fluorescent labels are used and when signals are measured from weakly expressed fluorescent proteins in the presence of background autofluorescence and stray light. Analysis of spectral image stacks in two spatial dimensions (2D) from single confocal slices indicates little or no cAMP gradients formed within pulmonary microvascular endothelial cells (PMVECs) under baseline conditions or following 10 min treatment with the adenylyl cyclase activator forskolin. However, analysis of spectral image stacks in 3D demonstrates marked cAMP gradients from the apical to basolateral face of PMVECs. Results demonstrate that spectral imaging approaches can be used to assess cAMP gradients-and in general gradients in fluorescence and FRET-within intact cells. Results also demonstrate that 2D imaging studies of localized fluorescence signals and, in particular, cAMP signals, whether using epifluorescence or confocal microscopy, may lead to erroneous conclusions about the existence and/or magnitude of gradients in either FRET or the underlying cAMP signals. Thus, with the exception of cellular structures that can be considered in one spatial dimension, such as neuronal processes, 3D measurements are required to assess mechanisms underlying compartmentalization and specificity within intracellular signaling pathways.

5D imaging approaches reveal the formation of distinct intracellular cAMP spatial gradients.

Cyclic AMP (cAMP) is a ubiquitous second messenger known to differentially regulate many cellular functions. Several lines of evidence suggest that the distribution of cAMP within cells is not uniform. However, to date, no studies have measured the kinetics of 3D cAMP distributions within cells. This is largely due to the low signal-to-noise ratio of FRET-based probes. We previously reported that hyperspectral imaging improves the signal-to-noise ratio of FRET measurements. Here we utilized hyperspectral imaging approaches to measure FRET signals in five dimensions (5D) - three spatial (x, y, z), wavelength (λ), and time (t) - allowing us to visualize cAMP gradients in pulmonary endothelial cells. cAMP levels were measured using a FRET-based sensor (H188) comprised of a cAMP binding domain sandwiched between FRET donor and acceptor - Turquoise and Venus fluorescent proteins. We observed cAMP gradients in response to 0.1 or 1 µM isoproterenol, 0.1 or 1 µM PGE1, or 50 µM forskolin. Forskolin- and isoproterenol-induced cAMP gradients formed from the apical (high cAMP) to basolateral (low cAMP) face of cells. In contrast, PGE1-induced cAMP gradients originated from both the basolateral and apical faces of cells. Data suggest that 2D (x,y) studies of cAMP compartmentalization may lead to erroneous conclusions about the existence of cAMP gradients, and that 3D (x,y,z) studies are required to assess mechanisms of signaling specificity. Results demonstrate that 5D imaging technologies are powerful tools for measuring biochemical processes in discrete subcellular domains. This work was supported by NIH P01HL066299, R01HL058506, S10RR027535, AHA 16PRE27130004 and the Abraham Mitchell Cancer Research Fund.

Three dimensional measurement of cAMP gradients using hyperspectral confocal microscopy.

Cyclic AMP (cAMP) is a ubiquitous second messenger known to differentially regulate many cellular functions over a wide range of timescales. Several lines of evidence have suggested that the distribution of cAMP within cells is not uniform, and that cAMP compartmentalization is largely responsible for signaling specificity within the cAMP signaling pathway. However, to date, no studies have experimentally measured three dimensional (3D) cAMP distributions within cells. Here we use both 2D and 3D hyperspectral microscopy to visualize cAMP gradients in endothelial cells from the pulmonary microvasculature (PMVECs). cAMP levels were measured using a FRET-based cAMP sensor comprised of a cAMP binding domain from EPAC sandwiched between FRET donors and acceptors - Turquoise and Venus fluorescent proteins. Data were acquired using either a Nikon A1R spectral confocal microscope or custom spectral microscopy system. Analysis of hyperspectral image stacks from a single confocal slice or from summed images of all slices (2D analysis) indicated little or no cAMP gradients were formed within PMVECs under basal conditions or following agonist treatment. However, analysis of hyperspectral image stacks from 3D cellular geometries (z stacks) demonstrate marked cAMP gradients from the apical to basolateral membrane of PMVECs. These results strongly suggest that 2D imaging studies of cAMP compartmentalization - whether epifluorescence or confocal microscopy - may lead to erroneous conclusions about the existence of cAMP gradients, and that 3D studies are required to assess mechanisms of signaling specificity.

Estimating the magnitude of near-membrane PDE4 activity in living cells.

Recent studies have demonstrated that functionally discrete pools of phosphodiesterase (PDE) activity regulate distinct cellular functions. While the importance of localized pools of enzyme activity has become apparent, few studies have estimated enzyme activity within discrete subcellular compartments. Here we present an approach to estimate near-membrane PDE activity. First, total PDE activity is measured using traditional PDE activity assays. Second, known cAMP concentrations are dialyzed into single cells and the spatial spread of cAMP is monitored using cyclic nucleotide-gated channels. Third, mathematical models are used to estimate the spatial distribution of PDE activity within cells. Using this three-tiered approach, we observed two pharmacologically distinct pools of PDE activity, a rolipram-sensitive pool and an 8-methoxymethyl IBMX (8MM-IBMX)-sensitive pool. We observed that the rolipram-sensitive PDE (PDE4) was primarily responsible for cAMP hydrolysis near the plasma membrane. Finally, we observed that PDE4 was capable of blunting cAMP levels near the plasma membrane even when 100 µM cAMP were introduced into the cell via a patch pipette. Two compartment models predict that PDE activity near the plasma membrane, near cyclic nucleotide-gated channels, was significantly lower than total cellular PDE activity and that a slow spatial spread of cAMP allowed PDE activity to effectively hydrolyze near-membrane cAMP. These results imply that cAMP levels near the plasma membrane are distinct from those in other subcellular compartments; PDE activity is not uniform within cells; and localized pools of AC and PDE activities are responsible for controlling cAMP levels within distinct subcellular compartments.

Assessing FRET using spectral techniques.

Förster resonance energy transfer (FRET) techniques have proven invaluable for probing the complex nature of protein-protein interactions, protein folding, and intracellular signaling events. These techniques have traditionally been implemented with the use of one or more fluorescence band-pass filters, either as fluorescence microscopy filter cubes, or as dichroic mirrors and band-pass filters in flow cytometry. In addition, new approaches for measuring FRET, such as fluorescence lifetime and acceptor photobleaching, have been developed. Hyperspectral techniques for imaging and flow cytometry have also shown to be promising for performing FRET measurements. In this study, we have compared traditional (filter-based) FRET approaches to three spectral-based approaches: the ratio of acceptor-to-donor peak emission, linear spectral unmixing, and linear spectral unmixing with a correction for direct acceptor excitation. All methods are estimates of FRET efficiency, except for one-filter set and three-filter set FRET indices, which are included for consistency with prior literature. In the first part of this study, spectrofluorimetric data were collected from a CFP-Epac-YFP FRET probe that has been used for intracellular cAMP measurements. All comparisons were performed using the same spectrofluorimetric datasets as input data, to provide a relevant comparison. Linear spectral unmixing resulted in measurements with the lowest coefficient of variation (0.10) as well as accurate fits using the Hill equation. FRET efficiency methods produced coefficients of variation of less than 0.20, while FRET indices produced coefficients of variation greater than 8.00. These results demonstrate that spectral FRET measurements provide improved response over standard, filter-based measurements. Using spectral approaches, single-cell measurements were conducted through hyperspectral confocal microscopy, linear unmixing, and cell segmentation with quantitative image analysis. Results from these studies confirmed that spectral imaging is effective for measuring subcellular, time-dependent FRET dynamics and that additional fluorescent signals can be readily separated from FRET signals, enabling multilabel studies of molecular interactions. © 2013 International Society for Advancement of Cytometry.

Hyperspectral imaging of FRET-based cGMP probes.

In recent years a variety of fluorescent probes for measurement of cGMP signals have been developed (Nikolaev et al., Nat. Methods 3:23-25, 2006; Honda et al., Proc Natl Acad Sci USA 98:2437-42, 2001; Nausch et al., Proc Natl Acad Sci USA 105:365-70, 2008). The probes are comprised of known cGMP binding sites-e.g., from phosphodiesterase type 5 (PDE5) or protein kinase G (PKG)-attached to fluorescent proteins. Binding of cGMP triggers conformational changes that alter the emitted fluorescence. In the case of Förster resonance energy transfer (FRET)-based probes, binding of cGMP alters the distance between the donor and acceptor fluorophores and thus alters FRET. However, FRET-based probes inherently have low signal-to-noise ratios, limiting the utility of these probes. Here we describe the use of hyperspectral imaging and analysis approaches to increase the signal-to-noise ratio of FRET-based cGMP measurements. These approaches are appropriate for monitoring changes in cGMP signals either in cell populations using a spectrofluorimeter or in single cells using spectral microscope systems with appropriate spectral filtering capabilities.

Calcineurin regulates homologous desensitization of natriuretic peptide receptor-A and inhibits ANP-induced testosterone production in MA-10 cells.

Receptor desensitization is a ubiquitous regulatory mechanism that defines the activatable pool of receptors, and thus, the ability of cells to respond to environmental stimuli. In recent years, the molecular mechanisms controlling the desensitization of a variety of receptors have been established. However, little is known about the molecular mechanisms that underlie desensitization of natriuretic peptide receptors, including natriuretic peptide receptor-A (NPR-A). Here we report that calcineurin (protein phosphatase 2B, PP2B, PPP3C) regulates homologous desensitization of NPR-A in murine Leydig tumor (MA-10) cells. We demonstrate that both pharmacological inhibition of calcineurin activity and siRNA-mediated suppression of calcineurin expression potentiate atrial natriuretic peptide (ANP)-induced cGMP synthesis. Treatment of MA-10 cells with inhibitors of other phosphoprotein phosphatases had little or no effect on ANP-induced cGMP accumulation. In addition, overexpression of calcineurin blunts ANP-induced cGMP synthesis. We also present data indicating that the inhibition of calcineurin potentiates ANP-induced testosterone production. To better understand the contribution of calcineurin in the regulation of NPR-A activity, we examined the kinetics of ANP-induced cGMP signals. We observed transient ANP-induced cGMP signals, even in the presence of phosphodiesterase inhibitors. Inhibition of both calcineurin and phosphodiesterase dramatically slowed the decay in the response. These observations are consistent with a model in which calcineurin mediated dephosphorylation and desensitization of NPR-A is associated with significant inhibition of cGMP synthesis. PDE activity hydrolyzes cGMP, thus lowering intracellular cGMP toward the basal level. Taken together, these data suggest that calcineurin plays a previously unrecognized role in the desensitization of NPR-A and, thereby, inhibits ANP-mediated increases in testosterone production.