The authors would like to acknowledge Dr. Kees Jalink (The Netherlands Cancer Institute and van Leeuwenhoek Center for Advanced Microscopy, Amsterdam, the Netherlands) for providing us with the H188 cAMP FRET biosensor and Kenny Trinh (College of Engineering, University of South Alabama) for technical help in reducing the time taken to run our custom developed programming scripts.
The authors would like to acknowledge the funding sources: American Heart Association (16PRE27130004), National Science Foundation; (1725937) NIH, S100D020149, S10RR027535, R01HL058506, P01HL066299).

<ol>
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Drs. Leavesley and Rich disclose financial interest in a start-up company, SpectraCyte, LLC, that was formed to commercialize spectral imaging technologies. However, all procedures described in this protocol were conducted using commercially available products not associated with SpectraCyte, LLC.

The development of FRET biosensors has allowed the measurement and visualization of cyclic nucleotide signals in single cells, and there is great promise for visualizing subcellular signaling events13,22,37,38. However, the use of FRET biosensors presents several limitations, including the low signal-to-noise characteristics of many fluorescent protein-based FRET reporters and the weak transfec...

Cyclic adenosine monophosphate (cAMP) is a second messenger involved in key cellular and physiological processes including cell division, calcium influx, gene transcription, and signal transduction. A growing body of evidence suggests the existence of cAMP compartments in the cell through which signaling specificity is achieved1,2,3,4,5,6,7. Until recently, cAMP compartmentalization was inferred based upon distinct physiological or cellular effects induced by different G-coupled receptor agonists8,9,10,11. More recently, FRET based fluorescence imaging probes have provided new approaches for the direct measurement and observation of cAMP signals in a cell12,13,14.
F&#246;rster resonance energy transfer (FRET) is a physical phenomenon in which energy transfer occurs between donor and acceptor molecules in a non-radiative fashion when the molecules are in close proximity15,16. With the development of FRET based fluorescent indicators, this physical phenomenon has been used in biological applications to study protein-protein interactions17, protein co-localization18, Ca+2 signaling19, gene expression20, cell division21 and cyclic nucleotide signaling. FRET based cAMP indicators typically consist of a cAMP binding domain, a donor fluorophore and an acceptor fluorophore22. For example, the H188 cAMP sensor12,22 used in this methodology consists of a cAMP binding domain obtained from Epac, sandwiched between Turquoise (donor) and Venus (acceptor) fluorophores. At basal conditions (unbound), Turquoise and Venus are in an orientation such that FRET occurs between the fluorophores. Upon binding of cAMP to the binding domain, a conformational change occurs such that Turquoise and Venus move apart resulting in a decrease in FRET.
FRET based imaging approaches offer a promising tool for investigating and visualizing cAMP signals within a cell. However, current FRET based microscopic imaging techniques are often only partially successful in achieving sufficient signal strength to measure FRET with subcellular spatial clarity. This is due to several factors, including the limited signal strength of many FRET reporters, the high level of precision required to accurately quantify changes in FRET efficiency, and the presence of confounding factors, such as cellular autofluorescence23,24. The result is often a FRET image that is plagued by weak SNR, making visualization of subcellular changes in FRET very difficult. In addition, investigation of spatially localized cAMP signals has been almost exclusively performed in only two spatial dimensions and the axial cAMP distribution has been rarely considered25. This is likely because low SNR impeded the ability to measure and visualize cAMP gradients in three spatial dimensions. To overcome limitations of using FRET sensors with low SNR, we have implemented hyperspectral imaging and analysis approaches to measure FRET in single cells25,26,27.
Hyperspectral imaging approaches were developed by NASA to differentiate terrestrial objects present in satellite images28,29. These techniques have since been translated to the fluorescence microscopy field30, with several commercial confocal microscope systems offering spectral detectors. In traditional (non-spectral) fluorescence imaging, the sample is excited using a band-pass filter or a laser line, and the emission is collected using a second band-pass filter, often selected to match the peak emission wavelength of the fluorophore(s). By contrast, hyperspectral imaging approaches seek to sample a complete spectral profile of either the fluorescence emission26,31,32 or excitation33,34 at specific wavelength intervals. In our previous studies, we showed that hyperspectral imaging and analysis approaches can offer improved quantification of FRET signals in cells when compared to traditional filter-based FRET imaging techniques26. Here, we present a methodology for performing 4-dimensional (x, y, z, and &#955;) hyperspectral FRET imaging and analysis to measure and visualize cAMP distributions in three spatial dimensions. These approaches have allowed visualization of agonist-induced cAMP spatial gradients in single cells25. Interestingly, depending on the agonist, cAMP gradients may be apparent in cells. The methodology presented here utilizes spectral unmixing of non-uniform background and cellular autofluorescence to improve the accuracy of the FRET measurements. While this methodology is demonstrated in pulmonary microvascular endothelial cells (PMVECs) using a cAMP FRET biosensor, the methodology could easily be modified for use with alternative FRET reporters or alternative cell lines.

<table><tbody><tr><td>Attofluor Cell Chamber</td><td>Invitrogen</td><td>A7816</td><td>Attofluor contains steel cell chambers and a rubber O-ring. Cell chamber holds the coverslip and O-ring provides a lock in mechanism to hold the buffer in cell chamber with out leakage</td></tr><tr><td>Dimethyl Sulfoxide (DMSO)</td><td>Fisher Scientific</td><td>BP231-100</td><td>Solvent used to prepare stock solution forskolin.</td></tr><tr><td>DRAQ5 Fluoroscent Probe Solution</td><td>Thermo Scientific</td><td>62252</td><td>Nuclear label</td></tr><tr><td>Dulbecco Modified Eagle Medium (DMEM)</td><td>Gibco</td><td>11965-092</td><td>Contains nutrients and growth factors for the cells to grow and divide in the culture dishes.</td></tr><tr><td>Fetal Bovine Serum (FBS)</td><td>Sigma</td><td>F6178</td><td>Growth factor suppliment that is added to culture medium, DMEM</td></tr><tr><td>Forskolin</td><td>Sigma</td><td>F3917</td><td>Adenyly cyclase activator.</td></tr><tr><td>H188 Cyclic AMP FRET biosensor</td><td>Netherland Cancer Institute, Dr. K. Jalink</td><td>Gift</td><td>Plasmid encoding Turquoise (donor fluorophore), Venus (acceptor fluorophore), and binding domain obtained from Epac.</td></tr><tr><td>Image J</td><td>image.net</td><td>Free download</td><td>Another image processing platform used to extact spectral information and image processing.</td></tr><tr><td>Integrating Sphere</td><td>Ocean Optics</td><td>FOIS-1</td><td>Used to measure illumination intensity of the laser line at different laser intensities (?).</td></tr><tr><td>Laminin Mouse Protein, Natural</td><td>Invitrogen</td><td>23017-015</td><td>Coverslips are coated with laminin and this helps in cell attachment, growth and motility of the cell.</td></tr><tr><td>Lipofectamine 3000 Transfection Kit</td><td>Invitrogen</td><td>L3000-015</td><td>Transfection reagent used to transfect cells with H188 FRET biosensor</td></tr><tr><td>MATLAB</td><td>Mathworks</td><td>R2019a</td><td>Image processing operations (linear unmixing and FRET efficiency calculations) are performed by writing custom programs in MATLAB programming environment</td></tr><tr><td>Nikon A1R confocal microscope</td><td>Nikon Instruments</td><td>Nikon A1R</td><td>Spectral image acquisition is performed using confocal microscope.</td></tr><tr><td>Nikon Elements Software</td><td>Nikon Instruments</td><td>Software dongle</td><td>used to export and handle nd2 image files (multidimensional image files) that are aquired using Nikon A1R</td></tr><tr><td>NIST-Traceable Calibration Lamp</td><td>Ocean Optics</td><td>LS-1-CAL-INT</td><td>A lamp with a known spectrum for use as a standard</td></tr><tr><td>PBS pH 7.4 (1X)</td><td>Gibco</td><td>10010-023</td><td>coomonly used buffer suring cell culture</td></tr><tr><td>Pulmonary Microvascular Endothelial Cells (PMVECs)</td><td>In house - Cell culture core, Univeristy of South Alabama</td><td>Isolated from Rat pulmonary microvasculature</td><td>PMVECs form inner lining of a blood vessel.</td></tr><tr><td>Penicillin-Streptomycin (10,000 U/ml)</td><td>Gibco</td><td>15140-122</td><td>antibiotics are added to culture medum to prevent contamination of the cells.</td></tr><tr><td>Pre-Cleaned Gold Seal Micro Slides</td><td>Clay Adams</td><td>3010</td><td>Microscope slides used for cell fixation</td></tr><tr><td>ProLong Diamond Antifade Mounting Media</td><td>Invitrogen</td><td>P36961</td><td>If samples are fixed using antifade mountant, then the later protects fluoroscent dyes and chromophores from fading.</td></tr><tr><td>Spectrometer</td><td>Ocean Optics</td><td>QE65000</td><td>Used to measure spectral response of the light source (?)</td></tr><tr><td>Trypsin-EDTA (0.25%)</td><td>Gibco</td><td>25200-056</td><td>Digests the protein-protein bond between the cell and cell matrix and helps to disscociate and lift the cells during cell plating.</td></tr><tr><td>Tyrodes Buffer</td><td>Made in-house</td><td>Made in-house</td><td>Tyrodes buffer is used to make working solutions and to maintain cells in aqueous solution during image acquisition.</td></tr><tr><td>6 Well Cell Culture Plate</td><td>Corning</td><td>3506</td><td>Laminin coated coverslips are placed in 6-well culture dish (one coverlisps/well). Cells along with medium is added into each well.</td></tr><tr><td>25 mm Round Microscope Cover Slips</td><td>Fisher Scientific</td><td>12545102</td><td>Cells were grown on round glass coverslips</td></tr><tr><td>60X Ojective</td><td>Nikon Instruments</td><td>Plan Apo VC 60X/1.2 WI &amp;infin;/0.15-0.18 WD 0.27</td><td>water immersion and commonly used objective for cells</td></tr></tbody></table>

measurement of 3-dimensional camp distributions in living cells using 4-dimensional (x, y, z, and &lambda;) 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 F&#1255;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.

This protocol describes the use of hyperspectral FRET imaging and analysis approaches to measure cAMP gradients in three spatial dimensions in living cells. There are several key steps involved in generating these results, for which careful attention is required while analyzing and quantifying the data. These key steps include construction of an appropriate spectral library, background spectral unmixing, thresholding to identify cell borders, and FRET efficiency calculations. Figure 1 illust...

Watch this Scientific Journal Video about Measurement of 3-Dimensional cAMP Distributions in Living Cells using 4-Dimensional x, y, z, and Î» Hyperspectral FRET Imaging and Analysis at JoVE.com

Measurement of 3-Dimensional cAMP Distributions in Living Cells using 4-Dimensional x, y, z, and &lambda; Hyperspectral FRET Imaging and Analysis

Due to inherent low signal-to-noise ratio (SNR) of F&#1255;rster resonance energy transfer (FRET) based sensors, measurement of cAMP signals has been challenging, especially in three spatial dimensions. Here, we describe a hyperspectral FRET imaging and analysis methodology that allows measurement of cAMP distribution in three spatial dimensions.

Measurement of 3-Dimensional cAMP Distributions in Living Cells using 4-Dimensional (x, y, z, and &lambda;) 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 ...

measurement-3-dimensional-camp-distributions-living-cells-using-4

Research

JoVE Journal

Biology

3.8K Views.  University of South Alabama. Vanwege de inherente lage signaal-ruisverhouding (SNR) van Fӧrster resonantie energieoverdracht (FRET) gebaseerde sensoren, is het meten van cAMP-signalen een uitdaging geweest, vooral in drie ruimtelijke dimensies. Hier beschrijven we een hyperspectrale FRET-beeldvormings- en analysemethodologie die het mogelijk maakt om de cAMP-verdeling in drie ruimtelijke dimensies te meten.

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

Live-cell Imaging of Migrating Cells Expressing Fluorescently-tagged Proteins in a Three-dimensional Matrix

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound healing, tissue regeneration, and cancer metastasis, cells must navigate through complex, three-dimensional extracellular tissue. To better understand the mechanisms behind these biological processes, it is important to examine the roles of the proteins responsible for driving cell migration. Here, we outline a protocol to study the mechanisms of cell migration using the epithelial cell line (MDCK), and a three-dimensional, fibrous, self-polymerizing matrix as a model system. This optically clear extracellular matrix is easily amenable to live-cell imaging studies and better mimics the physiological, soft tissue environment. This report demonstrates a technique for directly visualizing protein localization and dynamics, and deformation of the surrounding three-dimensional matrix. 
Examination of protein localization and dynamics during cellular processes provides key insight into protein functions. Genetically encoded fluorescent tags provide a unique method for observing protein localization and dynamics. Using this technique, we can analyze the subcellular accumulation of key, force-generating cytoskeletal components in real-time as the cell maneuvers through the matrix. In addition, using multiple fluorescent tags with different wavelengths, we can examine the localization of multiple proteins simultaneously, thus allowing us to test, for example, whether different proteins have similar or divergent roles. Furthermore, the dynamics of fluorescently tagged proteins can be quantified using Fluorescent Recovery After Photobleaching (FRAP) analysis. This measurement assays the protein mobility and how stably bound the proteins are to the cytoskeletal network.
By combining live-cell imaging with the treatment of protein function inhibitors, we can examine in real-time the changes in the distribution of proteins and morphology of migrating cells. Furthermore, we also combine live-cell imaging with the use of fluorescent tracer particles embedded within the matrix to visualize the matrix deformation during cell migration. Thus, we can visualize how a migrating cell distributes force-generating proteins, and where the traction forces are exerted to the surrounding matrix. Through these techniques, we can gain valuable insight into the roles of specific proteins and their contributions to the mechanisms of cell migration.

Cellular processes such as cell migration have traditionally been studied on two-dimensional, stiff plastic surfaces. This report describes a technique for directly visualizing protein localization and analyzing protein dynamics in cells migrating in a more physiologically relevant, three-dimensional matrix.

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound ...

Bioengineering

Visualization of Endosome Dynamics in Living Nerve Terminals with Four-dimensional Fluorescence Imaging

Four-dimensional (4D) light imaging has been used to study behavior of small structures within motor nerve terminals of the thin transversus abdominis muscle of the garter snake. Raw data comprises time-lapse sequences of 3D z-stacks. Each stack contains 4-20 images acquired with epifluorescence optics at focal planes separated by 400-1,500 nm. Steps in the acquisition of image stacks, such as adjustment of focus, switching of excitation wavelengths, and operation of the digital camera, are automated as much as possible to maximize image rate and minimize tissue damage from light exposure. After acquisition, a set of image stacks is deconvolved to improve spatial resolution, converted to the desired 3D format, and used to create a 4D "movie" that is suitable for variety of computer-based analyses, depending upon the experimental data sought. One application is study of the dynamic behavior of two classes of endosomes found in nerve terminals-macroendosomes (MEs) and acidic endosomes (AEs)-whose sizes (200-800 nm for both types) are at or near the diffraction limit. Access to 3D information at each time point provides several advantages over conventional time-lapse imaging. In particular, size and velocity of movement of structures can be quantified over time without loss of sharp focus. Examples of data from 4D imaging reveal that MEs approach the plasma membrane and disappear, suggesting that they are exocytosed rather than simply moving vertically away from a single plane of focus. Also revealed is putative fusion of MEs and AEs, by visualization of overlap between the two dye-containing structures as viewed in each three orthogonal projections.

Four-dimensional (4D) imaging is utilized to study the behavior and interactions among two types of endosomes in living vertebrate nerve terminals. Movement of these small structures is characterized in three dimensions, permitting confirmation of events such as endosome fusion and exocytosis.

Four-dimensional (4D) light imaging has been used to study behavior of small structures within motor nerve terminals of the thin transversus abdominis ...

Neuroscience

Cortical Actin Flow in T Cells Quantified by Spatio-temporal Image Correlation Spectroscopy of Structured Illumination Microscopy Data

Filamentous-actin plays a crucial role in a majority of cell processes including motility and, in immune cells, the formation of a key cell-cell interaction known as the immunological synapse. F-actin is also speculated to play a role in regulating molecular distributions at the membrane of cells including sub-membranous vesicle dynamics and protein clustering. While standard light microscope techniques allow generalized and diffraction-limited observations to be made, many cellular and molecular events including clustering and molecular flow occur in populations at length-scales far below the resolving power of standard light microscopy. By combining total internal reflection fluorescence with the super resolution imaging method structured illumination microscopy, the two-dimensional molecular flow of F-actin at the immune synapse of T cells was recorded. Spatio-temporal image correlation spectroscopy (STICS) was then applied, which generates quantifiable results in the form of velocity histograms and vector maps representing flow directionality and magnitude. This protocol describes the combination of super-resolution imaging and STICS techniques to generate flow vectors at sub-diffraction levels of detail. This technique was used to confirm an actin flow that is symmetrically retrograde and centripetal throughout the periphery of T cells upon synapse formation.

To investigate flow velocities and directionality of filamentous-actin at the T cell immunological synapse, live-cell super-resolution imaging is combined with total internal reflection fluorescence and quantified with spatio-temporal image correlation spectroscopy.

Filamentous-actin plays a crucial role in a majority of cell processes including motility and, in immune cells, the formation of a key cell-cell ...

Immunology and Infection

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called mechanotransduction. A crucial step in mechanotransduction is the transmission of forces between the external and internal environments. To transmit forces, there must be a sustained, unbroken physical linkage created by a series of protein-protein interactions. For a given protein-protein interaction, mechanical load can either have no effect on the interaction, lead to faster disassociation of the interaction, or even stabilize the interaction. Understanding how molecular load dictates protein turnover in living cells can provide valuable information about the mechanical state of a protein, in turn elucidating its role in mechanotransduction. Existing techniques for measuring force-sensitive protein dynamics either lack direct measurements of protein load or rely on the measurements performed outside of the cellular context. Here, we describe a protocol for the F&#246;rster resonance energy transfer-fluorescence recovery after photobleaching (FRET-FRAP) technique, which enables the measurement of force-sensitive protein dynamics within living cells. This technique is potentially applicable to any FRET-based tension sensor, facilitating the study of force-sensitive protein dynamics in variety of subcellular structures and in different cell types.

Here, we present a protocol for the simultaneous use of F&#246;rster resonance energy transfer-based tension sensors to measure protein load and fluorescence recovery after photobleaching to measure protein dynamics enabling the measurement of force-sensitive protein dynamics within living cells.

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called ...

Label-Free Identification of Lymphocyte Subtypes Using Three-Dimensional Quantitative Phase Imaging and Machine Learning

We describe here a protocol for the label-free identification of lymphocyte subtypes using quantitative phase imaging and machine learning. Identification of lymphocyte subtypes is important for the study of immunology as well as diagnosis and treatment of various diseases. Currently, standard methods for classifying lymphocyte types rely on labeling specific membrane proteins via antigen-antibody reactions. However, these labeling techniques carry the potential risks of altering cellular functions. The protocol described here overcomes these challenges by exploiting intrinsic optical contrasts measured by 3D quantitative phase imaging and a machine learning algorithm. Measurement of 3D refractive index (RI) tomograms of lymphocytes provides quantitative information about 3D morphology and phenotypes of individual cells. The biophysical parameters extracted from the measured 3D RI tomograms are then quantitatively analyzed with a machine learning algorithm, enabling label-free identification of lymphocyte types at a single-cell level. We measure the 3D RI tomograms of B, CD4+ T, and CD8+ T lymphocytes and identified their cell types with over 80% accuracy. In this protocol, we describe the detailed steps for lymphocyte isolation, 3D quantitative phase imaging, and machine learning for identifying lymphocyte types.

We describe a protocol for the label-free identification of lymphocyte subtypes using quantitative phase imaging and a machine learning algorithm. Measurements of 3D refractive index tomograms of lymphocytes present 3D morphological and biochemical information for individual cells, which is then analyzed with a machine-learning algorithm for identification of cell types.

We describe here a protocol for the label-free identification of lymphocyte subtypes using quantitative phase imaging and machine learning. Identification ...

An Analytical Tool that Quantifies Cellular Morphology Changes from Three-dimensional Fluorescence Images

The most common software analysis tools available for measuring fluorescence images are for two-dimensional (2D) data that rely on manual settings for inclusion and exclusion of data points, and computer-aided pattern recognition to support the interpretation and findings of the analysis. It has become increasingly important to be able to measure fluorescence images constructed from three-dimensional (3D) datasets in order to be able to capture the complexity of cellular dynamics and understand the basis of cellular plasticity within biological systems. Sophisticated microscopy instruments have permitted the visualization of 3D fluorescence images through the acquisition of multispectral fluorescence images and powerful analytical software that reconstructs the images from confocal stacks that then provide a 3D representation of the collected 2D images. Advanced design-based stereology methods have progressed from the approximation and assumptions of the original model-based stereology1 even in complex tissue sections2. Despite these scientific advances in microscopy, a need remains for an automated analytic method that fully exploits the intrinsic 3D data to allow for the analysis and quantification of the complex changes in cell morphology, protein localization and receptor trafficking. 
Current techniques available to quantify fluorescence images include Meta-Morph (Molecular Devices, Sunnyvale, CA) and Image J (NIH) which provide manual analysis. Imaris (Andor Technology, Belfast, Northern Ireland) software provides the feature MeasurementPro, which allows the manual creation of measurement points that can be placed in a volume image or drawn on a series of 2D slices to create a 3D object. This method is useful for single-click point measurements to measure a line distance between two objects or to create a polygon that encloses a region of interest, but it is difficult to apply to complex cellular network structures. Filament Tracer (Andor) allows automatic detection of the 3D neuronal filament-like however, this module has been developed to measure defined structures such as neurons, which are comprised of dendrites, axons and spines (tree-like structure). This module has been ingeniously utilized to make morphological measurements to non-neuronal cells3, however, the output data provide information of an extended cellular network by using a software that depends on a defined cell shape rather than being an amorphous-shaped cellular model. To overcome the issue of analyzing amorphous-shaped cells and making the software more suitable to a biological application, Imaris developed Imaris Cell. This was a scientific project with the Eidgen&ouml;ssische Technische Hochschule, which has been developed to calculate the relationship between cells and organelles. While the software enables the detection of biological constraints, by forcing one nucleus per cell and using cell membranes to segment cells, it cannot be utilized to analyze fluorescence data that are not continuous because ideally it builds cell surface without void spaces. To our knowledge, at present no user-modifiable automated approach that provides morphometric information from 3D fluorescence images has been developed that achieves cellular spatial information of an undefined shape (Figure 1). 
We have developed an analytical platform using the Imaris core software module and Imaris XT interfaced to MATLAB (Mat Works, Inc.). These tools allow the 3D measurement of cells without a pre-defined shape and with inconsistent fluorescence network components. Furthermore, this method will allow researchers who have extended expertise in biological systems, but not familiarity to computer applications, to perform quantification of morphological changes in cell dynamics.

We developed a software platform that utilizes Imaris Neuroscience, ImarisXT and MATLAB to measure the changes in morphology of an undefined shape taken from three-dimensional confocal fluorescence of single cells. This novel approach can be used to quantify changes in cell shape following receptor activation and therefore represents a possible additional tool for drug discovery.

The most common software analysis tools available for measuring fluorescence images are for two-dimensional (2D) data that rely on manual settings for ...

Automated Two-dimensional Spatiotemporal Analysis of Mobile Single-molecule FRET Probes

Single-molecule F&#246;rster resonance energy transfer (smFRET) is a versatile technique reporting on distances in the sub-nanometer to nanometer range. It has been used in a wide range of biophysical and molecular biological experiments, including the measurement of molecular forces, characterization of conformational dynamics of biomolecules, observation of intracellular colocalization of proteins, and determination of receptor-ligand interaction times. In a widefield microscopy configuration, experiments are typically performed using surface-immobilized probes. Here, a method combining single-molecule tracking with alternating excitation (ALEX) smFRET experiments is presented, permitting the acquisition of smFRET time traces of surface-bound, yet mobile probes in plasma membranes or glass-supported lipid bilayers. For the analysis of recorded data, an automated, open-source software collection was developed supporting (i) the localization of fluorescent signals, (ii) single-particle tracking, (iii) determination of FRET-related quantities including correction factors, (iv) stringent verification of smFRET traces, and (v) intuitive presentation of the results. The generated data can conveniently be used as input for further exploration via specialized software, e.g., for the assessment of the diffusional behavior of probes or the investigation of FRET transitions.

This article presents a method for spatiotemporal analysis of mobile, single-molecule F&#246;rster resonance energy transfer (smFRET)-based probes using widefield fluorescence microscopy. The newly developed software toolkit allows the determination of smFRET time traces of moving probes, including the correct FRET efficiency and the molecular positions, as functions of time.

Single-molecule F&#246;rster resonance energy transfer (smFRET) is a versatile technique reporting on distances in the sub-nanometer to nanometer range. ...

Comprehensive Protocol for Cell Identification and Metabolic Analysis Using OPTIR-FISH

Optical Photothermal Infrared-Fluorescence In Situ Hybridization (OPTIR-FISH)

Understanding the metabolic activities of individual cells within complex communities is critical for unraveling their role in human disease. Here, we present a comprehensive protocol for simultaneous cell identification and metabolic analysis with the OPTIR-FISH platform by combining rRNA-tagged FISH probes and isotope-labeled substrates. Fluorescence imaging provides cell identification by the specific binding of rRNA-tagged FISH probes, while OPTIR imaging provides metabolic activities within single cells by isotope-induced red shift on OPTIR spectra. Using bacteria cultured with 13C-glucose as a test bed, the protocol outlines microbial culture with isotopic labeling, fluorescence in situ hybridization (FISH), sample preparation, optimization of the OPTIR-FISH imaging setup, and data acquisition. We also demonstrate how to perform image analysis and interpret spectral data at the single-cell level with high throughput. This protocol&#39;s standardized and detailed nature will greatly facilitate its adoption by researchers from diverse backgrounds and disciplines within the broad single-cell metabolism research community.

Author Spotlight: Integrated OPTIR-FISH for Single-Cell Metabolic and Identity Analysis in Complex Environments

Here, we present a protocol using optical photothermal infrared-fluorescence in situ hybridization (OPTIR-FISH), also known as mid-infrared photothermal-FISH (MIP-FISH), to identify individual cells and understand their metabolism. This methodology can be applied broadly for diverse applications, including mapping cellular metabolism with single-cell resolution.

Understanding the metabolic activities of individual cells within complex communities is critical for unraveling their role in human disease. Here, we ...

Multicellular Spheroid Formation for FLIM-Based Metabolic and Hypoxia Analysis

Production and Multi-Parameter Live Cell Fluorescence Lifetime Imaging Microscopy (FLIM) of Multicellular Spheroids

Multicellular tumor spheroids are a popular 3D tissue microaggregate&#160;model for reproducing tumor microenvironment, testing and optimizing drug therapies&#160;and using bio- and nanosensors in a 3D context. Their ease of production, predictable size, growth, and observed nutrient and metabolite gradients are important to recapitulate the 3D niche-like&#160;cell microenvironment. However, spheroid heterogeneity and variability of their production methods can influence overall cell metabolism, viability, and drug response. This makes it difficult to choose the most appropriate&#160;methodology, considering the requirements in size, variability, needs of biofabrication, and use as in vitro 3D tissue models in stem and cancer cell biology. In particular, spheroid production can influence their compatibility with quantitative live microscopies, such as optical metabolic imaging, fluorescence lifetime imaging microscopy (FLIM), monitoring of spheroid&#160;hypoxia with nanosensors, or viability. Here, a number of conventional spheroid formation protocols are presented, highlighting their compatibility with the live widefield, confocal, and two-photon microscopies. The follow-up imaging to analysis&#160;pipeline with multiplexed autofluorescence FLIM and, using various types of cancer and stem cell spheroids, is also presented.

Author Spotlight: Standardizing Spheroid Formation Methods for Metabolic and Oxygenation Analysis Using Fluorescence Lifetime Imaging Microscopy

Here, we describe different multicellular spheroid formation methods to perform follow-up multi-parameter live cell microscopy. Using fluorescence lifetime imaging microscopy (FLIM), cellular autofluorescence, staining dyes, and nanoparticles, the approach for analysis of cell metabolism, hypoxia, and cell death in live three-dimensional (3D) cancer and stem cell-derived spheroids is demonstrated.

Multicellular tumor spheroids are a popular 3D tissue microaggregate&#160;model for reproducing tumor microenvironment, testing and optimizing drug ...

FRET Imaging in Three-dimensional Hydrogels

Imaging of F&#246;rster resonance energy transfer (FRET)&#160;is a powerful tool for examining cell biology in real-time. Studies utilizing FRET commonly employ two-dimensional (2D) culture, which does not mimic the three-dimensional (3D) cellular microenvironment. A method to perform quenched emission FRET imaging using conventional widefield epifluorescence microscopy of cells within a 3D hydrogel environment is presented. Here an analysis method for ratiometric FRET probes that yields linear ratios over the probe activation range is described. Measurement of intracellular cyclic adenosine monophosphate (cAMP) levels is demonstrated in chondrocytes under forskolin stimulation using a probe for EPAC1 activation (ICUE1) and the ability to detect differences in cAMP signaling dependent on hydrogel material type, herein a photocrosslinking hydrogel (PC-gel, polyethylene glycol dimethacrylate) and a thermoresponsive hydrogel (TR-gel). Compared with 2D FRET methods, this method requires little additional work. Laboratories already utilizing FRET imaging in 2D can easily adopt this method to perform cellular studies in a 3D microenvironment. It can further be applied to high throughput drug screening in engineered 3D microtissues. Additionally, it is compatible with other forms of FRET imaging, such as anisotropy measurement and fluorescence lifetime imaging (FLIM), and with advanced microscopy platforms using confocal, pulsed, or modulated illumination.

F&#246;rster resonance energy transfer (FRET) imaging is a powerful tool for real-time cell biology studies. Here a method for FRET imaging cells in physiologic three-dimensional (3D) hydrogel microenvironments using conventional epifluorescence microscopy is presented. An analysis for ratiometric FRET probes that yields linear ratios over the activation range is described.

Imaging of F&#246;rster resonance energy transfer (FRET)&#160;is a powerful tool for examining cell biology in real-time. Studies utilizing FRET commonly ...

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

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