Name: measurement of 3-dimensional camp distributions in living cells using 4-dimensional (x, y, z, and &lambda;) hyperspectral fret imaging and analysis
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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).

This protocol follows procedures approved by the University of South Alabama Institutional Animal Care and Use Committee.
1. Cell, sample, and reagent preparation for imaging
<ol>
	<li>Isolate rat pulmonary microvascular endothelial cells (PMVECs) as described previously35. 
	NOTE: Cells were isolated and cultured by the Cell Culture Core at the University of South Alabama, Mobile, AL on 100 mm cell culture dishes.</li>
	<li>Seed isolated PMVECs on 25 mm round gl...

<ol>
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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 &#8734;/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 &amp;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 ...

This protocol is significant as it provides a method for measuring and visualizing FRET or cyclic AMP signals in three spatial dimensions in individual cells. Cyclic AMP distributions may occur axially in addition to laterally and it is important to acquire FRET cyclic AMP image data in three dimensions to sample these distributions. On the day of imaging, warm Tyrodes buffer to 37 degrees Celsius in a water bath and mount a cover slip seated with the transfected cells of interest into a cell chamber.Secure the top of the chamber with a mounting gasket to prevent leaking and use a delicate task wiper to remove any excess medium or adherent cells from the bottom of the cover slip. Add 800 microliters of working buffer and four microliters of five millimolar nuclear label to the cell chamber and gently rock the chamber for five to 10 seconds. Then cover the cell chamber with aluminum foil for a 10 minute incubation at room temperature.To image the cells, select the 60X water immersion objective on a confocal microscope equipped with a spectral detector and add a drop of water to the objective. Place the loaded cell chamber onto the microscope stage and tune the filter knob to select the appropriate filter set. Select the fluorescence widefield mode and use the eyepieces to select a field of view containing cells expressing the FRET cyclic AMP sensor.Open the imaging software, select the confocal mode, and unlock the laser interlock button. Open the A1 settings menu. Check the boxes corresponding to the 405 and 561 nanometer laser lines and set these spectral detections to SD, the resolution to 10, and the channels to 32.Select the start and end wavelength values to set the wavelength range. Open the A1 Settings menu to select the binning skip icon. Select the number 15 box and click OK.Set the laser intensities to eight percent for the 405 nanometer laser and to two percent for the 561 nanometer laser.Set the detector gain to 149 and the pinhole radius to 2.4 Airy disk units. Set the scan speed to 25 spectral frames per second, and select the unidirectional scan direction. Set the count to four and the scan size to 1024 by 1024.To define the z-stack acquisition parameters, click View, Acquisition Control, and ND Acquisition, and enter the file destination and name in the pop window to save the ND file. Click the z-series box and click Live in the A1 Plus Settings window to open a live viewing window. Adjust the focus knob to focus on the cells while looking in the preview window.Adjust the focus knob on the microscope to select the top of the cell and click Top in the ND Acquisition window to set the current position as the top. Adjust the focus knob on the microscope to select the bottom of the cell and click Bottom to set the current position as the bottom. Enter one micrometer for the step size.Select Top Bottom for the z-scan direction and click run to acquire a z-stack. When the z-stack acquisition is complete, use a pipette to gently add the reagent of interest to the chamber without disturbing the cells. After 10 minutes, change the file name and acquire a second z-stack image.After imaging, create new folders with the same file names as the spectral z-stack images and open the spectral image file of interest. Click File, Import/Export, and Export ND Document. In the pop-up window, select the newly created folder, TIFF for the file type, Mono image for each channel, and keep bit depth.Click export to export the individual files to the folder and open the linear spectral unmixing software. Open the Linear Unmixing. m file and click Run.Browse and select the folder containing the exported TIFF file sequence and click OK to open the wavelength and z-slice window. Copy the file name of the first file without the z-slice and channel number from the folder and paste it into the first step of the Enter of the imagename box, enter the number of channels in the Enter the number of wavelength bands box, and the number of z-slices in the Enter the number of Z Slices box and click OK.Then in the pop-up window, browse and select the Wavelength. mat file and click Open.Browse and select the Library. mat file in the new pop-up window and click Open again to initiate the slice unmixing. To calculate the FRET efficiency, open the multiFRRCF.m programming script and enter the number of experimental trials to analyze in the How many folders to reslice box. Click OK.Browse to select the unmixed folders and click OK again. In the new pop-up window, set the scaling factor to 12.4, the threshold to 56, the X, Y, and Z frequency to five, five, and one respectively, and the smoothing algorithm to Gaussian.Then click Run to perform the FRET measurements and reslicing. To map the FRET efficiency to cyclic AMP levels, open the Mapping_FRET.Efficiency_to_cAMP_concentration. m file and click Run.Navigate to and select the first grayscale FRET image and click OK.Then open the FRET cyclic AMP images to inspect the distribution of the cyclic AMP signals in three dimensions. In these images, three-dimensional views of false colored raw hyperspectral image data acquired using confocal microscopy at baseline and 10 minutes after forskolin treatment can be observed. In this analysis, an acceptor fluorophore was completely photo destructed allowing spectral signatures of the donor and acceptor with one-to-one stoichiometry to be obtained.Non-transfected cells labeled with a nuclear dye were utilized to obtain the pure spectrum of the dye fluorophore. Combining the spectra of the donor, acceptor, and nuclear dye fluorophores allows the creation of a three component library. Here, the sources of three background spectral signatures can be observed.Although these signals are distributed non-uniformly within the sample and cannot simply be subtracted out, adding the spectral signatures of these signals to the spectral library and using linear unmixing to separate the signals provides an approach for removing these confounding signals from the donor and acceptor signals prior to calculating their FRET efficiencies. The six component spectral library can then be used to perform linear spectral unmixing for each slice in the axial image. Here, the changes in FRET efficiency and cyclic AMP levels in different XY plane slices can be observed allowing a comparison between the baseline conditions and the conditions 10 minutes after forskolin treatment.This technique could also be used to quantify the FRET efficiency of other reporters in three dimensions in both cells and tissues.

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

Research

JoVE Journal

Biology

Pulse-chase Analysis of N-linked Sugar Chains from Glycoproteins in Mammalian Cells

Attachment of the Glc3Man9GlcNAc2 precursor oligosaccharide to nascent polypeptides in the ER is a common modification for secretory proteins. Although this modification was implicated in several biological processes, additional aspects of its function are emerging, with recent evidence of its role in the production of signals for glycoprotein quality control and trafficking. Thus, phenomena related to N-linked glycans and their processing are being intensively investigated. Methods that have been recently developed for proteomic analysis have greatly improved the characterization of glycoprotein N-linked glycans. Nevertheless, they do not provide insight into the dynamics of the sugar chain processing involved. For this, labeling and pulse-chase analysis protocols are used that are usually complex and give very low yields. We describe here a simple method for the isolation and analysis of metabolically labeled N-linked oligosaccharides. The protocol is based on labeling of cells with [2-3H] mannose, denaturing lysis and enzymatic release of the oligosaccharides from either a specifically immunoprecipitated protein of interest or from the general glycoprotein pool by sequential treatments with endo H and N-glycosidase F, followed by molecular filtration (Amicon). In this method the isolated oligosaccharides serve as an input for HPLC analysis, which allows discrimination between various glycan structures according to the number of monosaccharide units comprising them, with a resolution of a single monosaccharide. Using this method we were able to study high mannose N-linked oligosaccharide profiles of total cell glycoproteins after pulse-chase in normal conditions and under proteasome inhibition. These profiles were compared to those obtained from an immunoprecipitated ER-associated degradation (ERAD) substrate. Our results suggest that most NIH 3T3 cellular glycoproteins are relatively stable and that most of their oligosaccharides are trimmed to Man9-8GlcNAc2. In contrast, unstable ERAD substrates are trimmed to Man6-5GlcNAc2 and glycoproteins bearing these species accumulate upon inhibition of proteasomal degradation.

We describe a method for analysis of the alteration of N-linked glycans through the early life of glycoproteins after their biosynthesis in mammalian cells. This is achieved by pulse-chase analysis of metabolically labeled glycans, enzymatic release from glycoproteins and examination by HPLC.

Attachment of the Glc3Man9GlcNAc2 precursor oligosaccharide to nascent polypeptides in the ER is a common modification for secretory proteins. Although ...

Immunocytochemical Analysis of Human Pluripotent Stem Cells using a Self-Made Cytospin Apparatus

Human pluripotent stem cells (hPSCs) that include human embryonic stem cells (hESCs) and human induced pluripotent stem cells (hiPSCs) are exciting cell sources due to their limitless self-renewal capabilities and their potential to differentiate into multiple cell types. The pluripotent state of hPSCs is typically assessed by techniques such as qPCR, immunocytochemistry, and by other in vitro and in vivo differentiation strategies into multiple cell types. Among these, immunocytochemical techniques have been developed for routine characterization of the undifferentiated state of hPSCs based on analysis of candidate intracellular and cell-surface biomarkers. Given the fact that hPSCs grow as colonies, problems arise in quantifying the expression of these markers at the individual cell level on a routine basis. Flow cytometry analyses serve to address this issue but require cell numbers and use of reagents that are not normally conducive for routine quality control assessment of hPSC cultures. Thus, the development of practical and reproducible means of creating monolayer cell samples with preserved integrity for marker evaluation has many advantages in stem cell research. This greatly benefits immunocytochemical analysis because individual cells from the monolayer can be easily observed and quantified for the expression of specific markers. Towards this goal, a self-made cytospin apparatus was constructed and optimized for use with immunocytochemical staining. Two cell-surface markers (SSEA3/SSEA4) expression were analyzed in a variant BG01 stem cell line for the purpose of this protocol.

Suspension immunocytochemical staining of human pluripotent stem cells (hPSCs) for cell-surface markers (SSEA-3/SSEA-4) was achieved based on use of a self-made cytospin apparatus to create a monolayer of cells for observation and quantification.

Human pluripotent stem cells (hPSCs) that include human embryonic stem cells (hESCs) and human induced pluripotent stem cells (hiPSCs) are exciting cell ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

Rapid Genetic Analysis of Epithelial-Mesenchymal Signaling During Hair Regeneration

Hair follicle morphogenesis, a complex process requiring interaction between epithelia-derived keratinocytes and the underlying mesenchyme, is an attractive model system to study organ development and tissue-specific signaling. Although hair follicle development is genetically tractable, fast and reproducible analysis of factors essential for this process remains a challenge. Here we describe a procedure to generate targeted overexpression or shRNA-mediated knockdown of factors using lentivirus in a tissue-specific manner. Using a modified version of a hair regeneration model 5, 6, 11, we can achieve robust gain- or loss-of-function analysis in primary mouse keratinocytes or dermal cells to facilitate study of epithelial-mesenchymal signaling pathways that lead to hair follicle morphogenesis. We describe how to isolate fresh primary mouse keratinocytes and dermal cells, which contain dermal papilla cells and their precursors, deliver lentivirus containing either shRNA or cDNA to one of the cell populations, and combine the cells to generate fully formed hair follicles on the backs of nude mice. This approach allows analysis of tissue-specific factors required to generate hair follicles within three weeks and provides a fast and convenient companion to existing genetic models.

Tissue-specific analysis of a hair follicle regeneration model using lentivirus to mediate gain- or loss-of-function.

Hair follicle morphogenesis, a complex process requiring interaction between epithelia-derived keratinocytes and the underlying mesenchyme, is an ...

Simultaneous pH Measurement in Endocytic and Cytosolic Compartments in Living Cells using Confocal Microscopy

Intracellular pH is tightly regulated and differences in pH between the cytoplasm and organelles have been reported1. Regulation of cellular pH is crucial for homeostatic control of physiological processes that include: protein, DNA and RNA synthesis, vesicular trafficking, cell growth and cell division. Alterations in cellular pH homeostasis can lead to detrimental functional changes and promote progression of various diseases2. Various methods are available for measuring intracellular pH but very few of these allow simultaneous measurement of pH in the cytoplasm and in organelles. Here, we describe in detail a rapid and accurate method for the simultaneous measurement of cytoplasmic and organellar pH by using confocal microscopy on living cells3. This goal is achieved with the use of two pH-sensing ratiometric dyes that possess selective cellular compartment partitioning. For instance, SNARF-1 is compartmentalized inside the cytoplasm whereas HPTS is compartmentalized inside endosomal/lysosomal organelles. Although HPTS is commonly used as a cytoplasmic pH indicator, this dye can specifically label vesicles along the endosomal-lysosomal pathway after being taken up by pinocytosis3,4. Using these pH-sensing probes, it is possible to simultaneously measure pH within the endocytic and cytoplasmic compartments. The optimal excitation wavelength of HPTS varies depending on the pH while for SNARF-1, it is the optimal emission wavelength that varies. Following loading with SNARF-1 and HPTS, cells are cultured in different pH-calibrated solutions to construct a pH standard curve for each probe. Cell imaging by confocal microscopy allows elimination of artifacts and background noise. Because of the spectral properties of HPTS, this probe is better suited for measurement of the mildly acidic endosomal compartment or to demonstrate alkalinization of the endosomal/lysosomal organelles. This method simplifies data analysis, improves accuracy of pH measurements and can be used to address fundamental questions related to pH modulation during cell responses to external challenges.

Several methods are available for measuring intracellular pH but very few of these allow simultaneous measurement of cytoplasmic and organellar pH. Here, we describe in detail a rapid and accurate methodology to simultaneously measure cytoplasmic and vesicular pH by ratiometric imaging of living cells.

Intracellular pH is tightly regulated and differences in pH between the cytoplasm and organelles have been reported1. Regulation of cellular pH is crucial ...

Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the development of diverse techniques to visualize individual RNA transcripts in single living cells. One promising technique that has recently been described utilizes an oligonucleotide-based optical probe, ratiometric bimolecular beacon (RBMB), to detect RNA transcripts that were engineered to contain at least four tandem repeats of the RBMB target sequence in the 3&#8217;-untranslated region. RBMBs are specifically designed to emit a bright fluorescent signal upon hybridization to complementary RNA, but otherwise remain quenched. The use of a synthetic probe in this approach allows photostable, red-shifted, and highly emissive organic dyes to be used for imaging. Binding of multiple RBMBs to the engineered RNA transcripts results in discrete fluorescence spots when viewed under a wide-field fluorescent microscope. Consequently, the movement of individual RNA transcripts can be readily visualized in real-time by taking a time series of fluorescent images. Here we describe the preparation and purification of RBMBs, delivery into cells by microporation and live-cell imaging of single RNA transcripts.

Ratiometric bimolecular beacons (RBMBs) can be used to image single engineered RNA transcripts in living cells. Here, we describe the preparation and purification of RBMBs, delivery of RBMBs into cells by microporation and fluorescent imaging of single RNA transcripts in real-time.

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Fluorescence Imaging with One-nanometer Accuracy (FIONA)

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in the x-y plane. Here a summary of the FIONA technique is reported and examples of research that have been performed using FIONA are briefly described. First, how to set up the required equipment for FIONA experiments, i.e., a total internal reflection fluorescence microscopy (TIRFM), with details on aligning the optics, is described. Then how to carry out a simple FIONA experiment on localizing immobilized Cy3-DNA single molecules using appropriate protocols, followed by the use of FIONA to measure the 36 nm step size of a single truncated myosin Va motor labeled with a quantum dot, is illustrated. Lastly, recent effort to extend the application of FIONA to thick samples is reported. It is shown that, using a water immersion objective and quantum dots soaked deep in sol-gels and rabbit eye corneas (&#62;200 &#181;m), localization precision of 2-3 nm can be achieved.

Fluorescence Imaging with One-nanometer Accuracy FIONA

Single fluorophores can be localized with nanometer precision using FIONA. Here a summary of the FIONA technique is reported, and how to carry out FIONA experiments is described.

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in ...

Measurement of Calcium Fluctuations Within the Sarcoplasmic Reticulum of Cultured Smooth Muscle Cells Using FRET-based Confocal Imaging

Maintenance of steady-state calcium (Ca2+) levels in the sarcoplasmic reticulum (SR) of vascular smooth muscle cells (VSMCs) is vital to their overall health. A significant portion of intracellular Ca2+ content is found within the SR stores in VSMCs. As the only intracellular organelle with a close association to the surrounding extracellular space through plasma membrane-SR junctions, the SR can be considered to constitute the first line of response to any irregularity in Ca2+ transients, or stress experienced by the cell. Among its many functions, one of the most important is its role in the transmission of Ca2+-regulated signals throughout the cell to induce further cell-wide reactions downstream. The more common use of cytoplasmic Ca2+ indicators in this regard is overall insufficient for research into the highly dynamic changes to the intraluminal SR Ca2+ store that have yet to be fully characterized. Here, we provide a detailed protocol for the direct and clear measurement of luminal SR Ca2+. This tool is useful for investigation into the nuanced changes in SR Ca2+ that have significant subsequent effects on the normal function and health of the cell. Fluctuations in SR Ca2+ content specifically can provide us with a significant amount of information pertaining to cellular responses to disease or stress conditions experienced by the cell. In this method, a modified version of a SR-targeted Ca2+ indicator, D1SR, is used to detect Ca2+ fluctuations in response to the introduction of agents to cultured rat aortic smooth muscle cells (SMCs). Following incubation with the D1SR indicator, confocal fluorescence microscopy and fluorescence resonance energy transfer (FRET)-based imaging are used to directly observe changes to intraluminal SR Ca2+ levels under control conditions and with the addition of agonist agents that function to induce intracellular Ca2+ movement.

Currently, most available calcium indicators are used to quantify cytoplasmic calcium transients as indirect measures of calcium released from the sarcoplasmic reticulum in cultured smooth muscle cells. This protocol describes the use of a specific FRET-based indicator that allows direct measurement of calcium signals within the sarcoplasmic reticulum lumen.

Maintenance of steady-state calcium (Ca2+) levels in the sarcoplasmic reticulum (SR) of vascular smooth muscle cells (VSMCs) is vital to their overall ...

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 ...