作者感谢Kees Jalink博士（荷兰癌症研究所和荷兰阿姆斯特丹范利乌文霍克高级显微镜中心）为我们提供了H188 cAMP FRET生物传感器和肯尼·特里恩（南阿拉巴马大学工程学院）的技术帮助，以减少运行我们自定义开发的编程脚本所花的时间。
作者要感谢资金来源:美国心脏协会（16PRE27130004），国家科学基金会:（1725937） NIH， S100D020149， S10RR027535， R01HL058506， P01HL066299）.）

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Leavesley博士和Rich博士披露了一家初创公司SpectraCyte的财务利益，该公司的成立是为了将光谱成像技术商业化。但是，本协议中描述的所有程序都是使用与 SpectraCyte 有限责任公司无关的商用产品进行的。

FRET生物传感器的研制使单细胞环核苷酸信号的测量和可视化，对13、22、37、38等亚细胞信号事件的可视化大有可为。然而，使用FRET生物传感器存在一些局限性，包括许多荧光蛋白基FRET记者的低信号到噪声特性，以及FRET记者的弱透射或表达效率（这在某些细胞系中可能特别具有挑战性，如PMVECs）23、24...

环腺苷单磷酸盐（cAMP）是参与关键细胞和生理过程的第二个信使，包括细胞分裂、钙的涌入、基因转录和信号转导。越来越多的证据表明，细胞中存在cAMP隔间，通过这些隔间可以达到1、2、3、4、5、6、7的信号特异性。直到最近，CAMP分隔断是根据不同的G耦合受体激动剂8，9，10，11诱导的独特的生理或细胞效应推断的。最近，基于FRET的荧光成像探测器为直接测量和观察12、13、14细胞中的CAMP信号提供了新的方法。
Fürster共振能量转移（FRET）是一种物理现象，当分子接近15、16时，能量转移以非辐射的方式发生在供体和接受分子之间。随着基于FRET的荧光指标的发展，这种物理现象已用于生物应用，研究蛋白质-蛋白质相互作用17，蛋白质共定位18，Ca+2信号19，基因表达20，细胞分裂21和环核苷酸信号。基于FRET的cAMP指标通常包括一个CAMP结合域，一个捐赠者氟和一个接受荧光22。例如，H188 cAMP 传感器12，22用于此方法包括从 Epac 获得的 cAMP 结合域，夹在绿松石（捐赠者）和金星（接受者）荧光之间。在基础条件下（未绑定），绿松石和金星处于一个方向，因此FRET发生在萤管之间。将 cAMP 与绑定域结合后，会发生构象变化，使绿松石和金星分开，导致 FRET 减少。
基于 FRET 的成像方法为研究细胞内的 cAMP 信号提供了一个很有前途的工具。然而，目前的基于FRET的微观成像技术往往只部分成功达到足够的信号强度，以测量具有亚细胞空间清晰度的FRET。这是由于几个因素，包括许多FRET记者的信号强度有限，准确量化FRET效率变化所需的高精度，以及存在混淆因素，如细胞自发光23，24。结果往往是受弱 SNR 困扰的 FRET 图像，使得 FRET 的子细胞变化的可视化变得非常困难。此外，空间定位 CAMP 信号的调查几乎只在两个空间维度中执行，轴向 CAMP 分布很少被视为25。这可能是因为低 SNR 妨碍了在三个空间维度中测量和可视化 cAMP 梯度的能力。为了克服使用低SNR的FRET传感器的限制，我们采用了高光谱成像和分析方法，在单个细胞25、26、27中测量FRET。
美国宇航局开发了超光谱成像方法，以区分卫星图像28、29中存在的地球物体。此后，这些技术被翻译成荧光显微镜场30，几个商业共聚焦显微镜系统提供光谱探测器。在传统的（非光谱）荧光成像中，样品使用带通滤镜或激光线激发，并且使用第二个带通滤波器收集发射，通常选择以匹配荧光的峰值发射波长。相比之下，高光谱成像方法寻求以特定波长间隔对荧光发射26、31、32或激发33、34的完整光谱剖面进行采样。在我们之前的研究中，我们发现，与传统的基于过滤器的FRET成像技术相比，高光谱成像和分析方法可以改善细胞中FRET信号的量化。在这里，我们提出了一种执行 4 维（x、y、z 和 +）高光谱 FRET 成像和分析的方法，以测量和可视化三个空间维度的 cAMP 分布。这些方法使单细胞25中激动剂诱导的CAMP空间梯度可视化。有趣的是，根据激动剂的不同，CAMP梯度在细胞中可能很明显。此处提供的方法利用不均匀背景和细胞自发光的光谱解密，以提高 FRET 测量的准确性。虽然这种方法在使用 cAMP FRET 生物传感器的肺微血管内皮细胞 （PMVECs） 中得到了证明，但该方法可以很容易地修改，用于替代 FRET 记者或替代细胞系。

<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

循环AMP是第二个信使，参与广泛的细胞和生理活动。一些研究表明，cAMP 信号是分隔的，隔间化有助于 cAMP 信号通路中的信号特异性。基于Fürster共振能量转移（FRET）的生物传感器的发展，进一步增强了测量和可视化细胞中CAMP信号的能力。但是，这些测量通常局限于两个空间维度，这可能导致对数据的误解。迄今为止，由于使用本身信号与噪声比 （SNR） 较低的 FRET 传感器的技术限制，在三个空间维度（x、y 和 z）中对 CAMP 信号的测量非常有限。此外，由于光谱相声、信号强度有限和自发光等一系列因素，传统的基于滤波器的成像方法通常对精确测量局部子细胞区域的 CAMP 信号无效。为了克服这些限制，并允许基于FRET的生物传感器与多个荧光素一起使用，我们开发了高光谱FRET成像和分析方法，为计算FRET效率提供光谱特异性，并能够将FRET信号与混淆的自发光和/或来自其他荧光标签的信号隔离开来。在这里，我们介绍了实施高光谱FRET成像的方法，以及需要构建一个适当的光谱库，既不采样不足，也不采样过采，以执行光谱未混合。虽然我们提出了测量肺微血管内皮细胞（PMVECs）中三维CAMP分布的方法，但这种方法可用于研究CAMP在一系列细胞类型的空间分布。

本协议描述了使用高光谱 FRET 成像和分析方法测量活细胞中三个空间维度的 cAMP 梯度。生成这些结果涉及几个关键步骤，在分析和量化数据时需要仔细注意这些步骤。这些关键步骤包括构建适当的光谱库、背景光谱未混合、识别细胞边框的阈值以及 FRET 效率计算。 图1 说明了测量活细胞中FRET效率和cAMP水平所涉及的所有步骤的示意图流。如果执行得当，这些成像和分析步骤?...

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

由于基于 Fürster 共振能量转移 （FRET） 传感器的固有的低信号与噪声比 （SNR），CAMP 信号的测量一直具有挑战性，尤其是在三个空间维度方面。在这里，我们描述了一个高光谱FRET成像和分析方法，允许测量三个空间尺寸的CAMP分布。

使用 4 维（x、y、z 和 +）高光谱 FRET 成像和分析测量活细胞中的三维 cAMP 分布

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

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

3.8K 次观看.  University of South Alabama. 由于基于 Fürster 共振能量转移 （FRET） 传感器的固有的低信号与噪声比 （SNR），CAMP 信号的测量一直具有挑战性，尤其是在三个空间维度方面。在这里，我们描述了一个高光谱FRET成像和分析方法，允许测量三个空间尺寸的CAMP分布。

视频: 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.

活细胞成像的移植细胞在一个三维矩阵表示荧光标记的蛋白质

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

科研

生物工程

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.

皮层肌动蛋白流通过结构照明显微镜数据的时空图像相关光谱量化性T细胞

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 ...

免疫与感染

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.

荧光技术结合测定活细胞中的力敏感蛋白动力学

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

Monitoring Kinase and Phosphatase Activities Through the Cell Cycle by Ratiometric FRET

F&ouml;rster resonance energy transfer (FRET)-based reporters1 allow the assessment of endogenous kinase and phosphatase activities in living cells. Such probes typically consist of variants of CFP and YFP, intervened by a phosphorylatable sequence and a phospho-binding domain. Upon phosphorylation, the probe changes conformation, which results in a change of the distance or orientation between CFP and YFP, leading to a change in FRET efficiency (Fig 1). Several probes have been published during the last decade, monitoring the activity balance of multiple kinases and phosphatases, including reporters of PKA2, PKB3, PKC4, PKD5, ERK6, JNK7, Cdk18, Aurora B9 and Plk19. Given the modular design, additional probes are likely to emerge in the near future10. 
Progression through the cell cycle is affected by stress signaling pathways 11. Notably, the cell cycle is regulated differently during unperturbed growth compared to when cells are recovering from stress12.Time-lapse imaging of cells through the cell cycle therefore requires particular caution. This becomes a problem particularly when employing ratiometric imaging, since two images with a high signal to noise ratio are required to correctly interpret the results. Ratiometric FRET imaging of cell cycle dependent changes in kinase and phosphatase activities has predominately been restricted to sub-sections of the cell cycle8,9,13,14.
Here, we discuss a method to monitor FRET-based probes using ratiometric imaging throughout the human cell cycle. The method relies on equipment that is available to many researchers in life sciences and does not require expert knowledge of microscopy or image processing.

FRET-based reporters are increasingly used to monitor kinase and phosphatase activities in live cells. Here we describe a method on how to use FRET-based reporters to assess cell cycle-dependent changes in target phosphorylation.

通过细胞周期激酶和磷酸酶活性成比例的FRET监测

F&ouml;rster resonance energy transfer (FRET)-based reporters1 allow the assessment of endogenous kinase and phosphatase activities in living cells. Such ...

生物学

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.

用三维定量相位成像和机器学习技术对淋巴细胞亚型进行无标签鉴定

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.

细胞形态变化，从三维荧光图像量化的分析工具，

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.

移动式单分子FRET探头的自动化二维时空分析

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

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. 
We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisi&#230;, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.

Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

通过电穿孔内在和生物分子荧光观察生活中的微生物

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing ...

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.

FRET成像的三维水凝胶

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 ...

Open Source High Content Analysis Utilizing Automated Fluorescence Lifetime Imaging Microscopy

We present an open source high content analysis instrument utilizing automated fluorescence lifetime imaging (FLIM) for assaying protein interactions using F&#246;rster resonance energy transfer (FRET) based readouts of fixed or live cells in multiwell plates. This provides a means to screen for cell signaling processes read out using intramolecular FRET biosensors or intermolecular FRET of protein interactions such as oligomerization or heterodimerization, which can be used to identify binding partners. We describe here the functionality of this automated multiwell plate FLIM instrumentation and present exemplar data from our studies of HIV Gag protein oligomerization and a time course of a FRET biosensor in live cells. A detailed description of the practical implementation is then provided with reference to a list of hardware components and a description of the open source data acquisition software written in &#181;Manager. The application of FLIMfit, an open source MATLAB-based client for the OMERO platform, to analyze arrays of multiwell plate FLIM data is also presented. The protocols for imaging fixed and live cells are outlined and a demonstration of an automated multiwell plate FLIM experiment using cells expressing fluorescent protein-based FRET constructs is presented. This is complemented by a walk-through of the data analysis for this specific FLIM FRET data set.

We present an open source high content analysis (HCA) instrument utilizing automated fluorescence lifetime imaging (FLIM) for assaying protein interactions using F&#246;rster resonance energy transfer (FRET) based readouts. Data acquisition for this openFLIM-HCA instrument is controlled by software written in &#181;Manager and data analysis is undertaken in FLIMfit.

开源高内涵分析利用自动荧光寿命成像显微镜

We present an open source high content analysis instrument utilizing automated fluorescence lifetime imaging (FLIM) for assaying protein interactions ...