High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.     

Imaging green fluorescent protein fusions in living fission yeast cells

The use of green fluorescent protein (GFP) fusions as biosensors for examining protein localization and dynamics has revolutionized cell biology. Here, we describe the methods developed for imaging of GFP-fusions in the fission yeast Schizosaccharomyces pombe using fluorescence microscopy, with a focus on the use of time-lapse imaging to analyze the dynamics of microtubules. We discuss the considerations in fluorescence microscopy, cell preparation, data acquisition, and image analysis appropriate for analysis of living cells.     

Quantifying subcellular distribution of fluorescent fusion proteins in cells migrating within tissues

The movement of proteins within cells can provide dynamic indications of cell signaling and cell polarity, but methods are needed to track and quantify subcellular protein movement within tissue environments. Here we present a semiautomated approach to quantify subcellular protein location for hundreds of migrating cells within intact living tissue using retrovirally expressed fluorescent fusion proteins and time-lapse two-photon microscopy of intact thymic lobes. We have validated the method using GFP-PKCζ, a marker for cell polarity, and LAT-GFP, a marker for T-cell receptor signaling, and have related the asymmetric distribution of these proteins to the direction and speed of cell migration. These approaches could be readily adapted to other fluorescent fusion proteins, tissues and biological questions.     

Using bleach-chase to measure protein half-lives in living cells

Protein removal has a central role in numerous cellular processes. Obtaining systematic measurements of multiple protein removal rates is necessary to understand the principles that govern these processes, but it is currently a major technical challenge. To address this, we developed 'bleach-chase', a noninvasive method for measuring the half-lives of multiple proteins at high temporal resolution in living cells. The method uses a library of annotated human reporter cell clones, each with a unique fluorescently tagged protein expressed from its native chromosomal location. In this protocol, we detail a simple procedure that bleaches the cells and uses time-lapse fluorescence microscopy and automated image analysis to systematically measure the half-life dynamics of multiple proteins. The duration of the protocol is 4-5 d. The method may be applicable to a wide range of fluorescently tagged proteins and cell lines.     

Dynamic proteomics in individual human cells uncovers widespread cell-cycle dependence of nuclear proteins

We examined cell cycle-dependent changes in the proteome of human cells by systematically measuring protein dynamics in individual living cells. We used time-lapse microscopy to measure the dynamics of a random subset of 20 nuclear proteins, each tagged with yellow fluorescent protein (YFP) at its endogenous chromosomal location. We synchronized the cells in silico by aligning protein dynamics in each cell between consecutive divisions. We observed widespread (40%) cell-cycle dependence of nuclear protein levels and detected previously unknown cell cycle-dependent localization changes. This approach to dynamic proteomics can aid in discovery and accurate quantification of the extensive regulation of protein concentration and localization in individual living cells.     

Dual-colour imaging with GFP variants

Green fluorescent protein (GFP) has become an important tool in cell biology and is widely used as a reporter for imaging intracellular proteins and structures in live cells. Recently, spectral variants of GFP with red- and blue-shifted fluorescence emissions have been characterized, opening the possibility of double labelling with two different-coloured GFP fusion proteins. This article reviews recent advances in this technique, with special emphasis on time-lapse imaging applications in living cells.     

Real-time Imaging of Plant Cell Surface Dynamics with Variable-angle Epifluorescence Microscopy

A plant’s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and cytoskeletal rearrangement. Real-time and high-resolution image analysis of such intracellular events will increase the understanding of plant cell biology at the molecular level. Variable angle epifluorescence microscopy (VAEM) is an emerging technique that provides high-quality, time-lapse images of fluorescently-labeled proteins on the plant cell surface. In this article, practical procedures are described for VAEM specimen preparation, adjustment of the VAEM optical system, movie capturing and image analysis. As an example of VAEM observation, representative results are presented on the dynamics of PATROL1. This is a protein essential for stomatal movement, thought to be involved in proton pump delivery to plasma membranes in the stomatal complex of Arabidopsis thaliana. VAEM real-time observation of guard cells and subsidiary cells in A. thaliana cotyledons showed that fluorescently-tagged PATROL1 appeared as dot-like structures on plasma membranes for several seconds and then disappeared. Kymograph analysis of VAEM movie data determined the time distribution of the presence (termed ‘residence time’) of the dot-like structures. The use of VAEM is discussed in the context of this example.     

Selective photolabeling of proteins using photoactivatable GFP

Today's cell biologists rely on an assortment of advances in microscopy methods to study the inner workings of cells and tissues. Among these advances are fluorescent proteins which can be used to tag specifically and, in many cases, non-invasively proteins of interest within a living cell. Introduction of DNA encoding the fluorescently tagged protein of interest into a cell readily allows the visualization of the protein's localization and time-lapse imaging allows the movement of the structure or organelle to which the protein is localized to be observed. To monitor the movement of the protein within the population, researchers generally have to highlight a pool of molecules by perturbing the steady-state fluorescence. This perturbation has traditionally been performed by photobleaching the molecules within a selected region of the cell and monitoring the recovery of molecules into this region or the loss of molecules within other regions. Fluorescent proteins are now available, which allow a pool of molecules to be highlighted directly by photoactivation. Here, we discuss the technical aspects for using one of these recently developed photoactivatable fluorescent proteins, PA-GFP.     

Three-dimensional segmentation of nuclei and mitotic chromosomes for the study of cell divisions in live Drosophila embryos

Drosophila embryogenesis is an established model to investigate mechanisms and genes related to cell divisions in an intact multicellular organism. Progression through the cell cycle phases can be monitored in vivo using fluorescently labeled fusion proteins and time-lapse microscopy. To measure cellular properties in microscopic images, accurate and fast image segmentation methods are a critical prerequisite. To quantify static and dynamic features of interphase nuclei and mitotic chromosomes, we developed a three-dimensional (3D) segmentation method based on multiple level sets. We tested our method on 3D time-series images of live embryos expressing histone-2Av-green fluorescence protein. Our method is robust to low signal-to-noise ratios inherent to high-speed imaging, fluorescent signals in the cytoplasm, and dynamic changes of shape and texture. Comparisons with manual ground-truth segmentations showed that our method achieves more than 90% accuracy on the object as well as voxel levels and performs consistently throughout all cell cycle phases and developmental stages from syncytial blastoderm to postblastoderm mitotic domains.     

Sub-cellular localisation of GFP-tagged tobacco mitotic cyclins during the cell cycle and after spindle checkpoint activation

We have previously shown that the tobacco cyclin B1;1 protein accumulates during the G2 phase of the cell cycle and is subsequently destroyed during mitosis. Here, we investigated the sub-cellular localisation of two different B1-types and one A3-type cyclin during the cell cycle by using confocal imaging and differential interference contrast (DIC) microscopy. The cyclins were visualised as GFP-tagged fusion proteins in living tobacco cells. Both B1-type cyclins were found in the cytoplasm and in the nucleus during G2 but when cells entered into prophase, both cyclins became associated with condensing chromatin and remained on chromosomes until metaphase. As cells exited metaphase, the B1-type cyclins became degraded, as shown by time-lapse images. A stable variant of cyclin B1;1-GFP fusion protein, in which the destruction box had been mutated, maintained its association with the nuclear material at later phases of mitosis such as anaphase and telophase. Furthermore, we demonstrated that cyclin B1;1 protein is stabilised in metaphase-arrested cells after microtubule destabilising drug treatments. In contrast to the B1-type cyclins, the cyclin A3;1 was found exclusively in the nucleus in interphase cells and disappeared earlier than the cyclin B1 proteins during mitosis.     

Super-resolution imaging in live Caulobacter crescentus cells using photoswitchable EYFP

The commonly used, monomeric EYFP enabled imaging of intracellular protein structures beyond the optical resolution limit ('super-resolution' imaging) in living cells. By combining photoinduced activation of single EYFP fusions and time-lapse imaging, we obtained sub-40 nm resolution images of the filamentous superstructure of the bacterial actin protein MreB in live Caulobacter crescentus cells. These studies demonstrated that EYFP is a useful emitter for in vivo super-resolution imaging.     

Quantitative imaging of protein interactions in the cell nucleus

Over the past decade, genetically encoded fluorescent proteins have become widely used as noninvasive markers in living cells. The development of fluorescent proteins, coupled with advances in digital imaging, has led to the rapid evolution of live-cell imaging methods. These approaches are being applied to address biological questions of the recruitment, co-localization, and interactions of specific proteins within particular subcellular compartments. In the wake of this rapid progress, however, come important issues associated with the acquisition and analysis of ever larger and more complex digital imaging data sets. Using protein localization in the mammalian cell nucleus as an example, we will review some recent developments in the application of quantitative imaging to analyze subcellular distribution and co-localization of proteins in populations of living cells. In this report, we review the principles of acquiring fluorescence resonance energy transfer (FRET) microscopy measurements to define the spatial relationships between proteins. We then discuss how fluorescence lifetime imaging microscopy (FLIM) provides a method that is independent of intensity-based measurements to detect localized protein interactions with spatial resolution. Finally, we consider potential problems associated with the expression of proteins fused to fluorescent proteins for FRET-based measurements from living cells.     

N-Way FRET Microscopy of Multiple Protein-Protein Interactions in Live Cells

Fluorescence Resonance Energy Transfer (FRET) microscopy has emerged as a powerful tool to visualize nanoscale protein-protein interactions while capturing their microscale organization and millisecond dynamics. Recently, FRET microscopy was extended to imaging of multiple donor-acceptor pairs, thereby enabling visualization of multiple biochemical events within a single living cell. These methods require numerous equations that must be defined on a case-by-case basis. Here, we present a universal multispectral microscopy method (N-Way FRET) to enable quantitative imaging for any number of interacting and non-interacting FRET pairs. This approach redefines linear unmixing to incorporate the excitation and emission couplings created by FRET, which cannot be accounted for in conventional linear unmixing. Experiments on a three-fluorophore system using blue, yellow and red fluorescent proteins validate the method in living cells. In addition, we propose a simple linear algebra scheme for error propagation from input data to estimate the uncertainty in the computed FRET images. We demonstrate the strength of this approach by monitoring the oligomerization of three FP-tagged HIV Gag proteins whose tight association in the viral capsid is readily observed. Replacement of one FP-Gag molecule with a lipid raft-targeted FP allowed direct observation of Gag oligomerization with no association between FP-Gag and raft-targeted FP. The N-Way FRET method provides a new toolbox for capturing multiple molecular processes with high spatial and temporal resolution in living cells.     

HuC–eGFP mosaic labelling of neurons in zebrafish enables in vivo live cell imaging of growth cones

The field of axon guidance is taking advantage of the powerful genetic and imaging tools that are now available to visualise growth behaviour in living cells, both in vivo and in real time. We have developed a method to visualise individual neurons within the living zebrafish embryo which provides exceptional cellular resolution of growth cones and their filopodia. We generated a DNA construct in which the HuC promoter drives expression of eGFP. Injection of the plasmid into single cell fertilised zebrafish egg resulted in mosaic expression of eGFP in neurons throughout the developing embryo. By manipulating the concentration of injected plasmid, it was possible to optimise the numbers of neurons that expressed the construct so that individual growth cones could be easily visualised. We then used time-lapse high magnification widefield epifluorescence microscopy to visualise the growth cones as they were exploring their environment. Growth cones both near the surface of the embryo as well as deep within the developing brain of embryos at 20?h post fertilisation were clearly imaged. With time-lapse sequence imaging with intervals between frames as frequent as 1?s there was minimal loss of fluorescence intensity and the dynamic nature of the growth cones became evident. This method therefore provides high magnification, high resolution time-lapse imaging of living neurons in vivo and by use of widefield epifluorescence rather than confocal it is a relatively inexpensive microscopy method.     

Total internal reflection fluorescence microscopy for single-molecule imaging in living cells

Marvelous background rejection in total internal reflection fluorescence microscopy (TIR-FM) has made it possible to visualize single-fluorophores in living cells. Cell signaling proteins including peptide hormones, membrane receptors, small G proteins, cytoplasmic kinases as well as small signaling compounds have been conjugated with single chemical fluorophore or tagged with green fluorescent proteins and visualized in living cells. In this review, the reasons why single-molecule analysis is essential for studies of intracellular protein systems such as cell signaling system are discussed, the instrumentation of TIR-FM for single-molecule imaging in living cells is explained, and how single molecule visualization has been used in cell biology is illustrated by way of two examples: signaling of epidermal growth factor in mammalian cells and chemotaxis of Dictyostelium amoeba along a cAMP gradient. Single-molecule analysis is an ideal method to quantify the parameters of reaction dynamics and kinetics of unitary processes within intracellular protein systems. Knowledge of these parameters is crucial for the understanding of the molecular mechanisms underlying intracellular events, thus single-molecule imaging in living cells will be one of the major technologies in cellular nanobiology.     

High-throughput RNAi screening by time-lapse imaging of live human cells

RNA interference (RNAi) is a powerful tool to study gene function in cultured cells. Transfected cell microarrays in principle allow high-throughput phenotypic analysis after gene knockdown by microscopy. But bottlenecks in imaging and data analysis have limited such high-content screens to endpoint assays in fixed cells and determination of global parameters such as viability. Here we have overcome these limitations and developed an automated platform for high-content RNAi screening by time-lapse fluorescence microscopy of live HeLa cells expressing histone-GFP to report on chromosome segregation and structure. We automated all steps, including printing transfection-ready small interfering RNA (siRNA) microarrays, fluorescence imaging and computational phenotyping of digital images, in a high-throughput workflow. We validated this method in a pilot screen assaying cell division and delivered a sensitive, time-resolved phenoprint for each of the 49 endogenous genes we suppressed. This modular platform is scalable and makes the power of time-lapse microscopy available for genome-wide RNAi screens.     

Spectral imaging and its applications in live cell microscopy

In biological microscopy, the ever expanding range of applications requires quantitative approaches that analyze several distinct fluorescent molecules at the same time in the same sample. However, the spectral properties of the fluorescent proteins and dyes presently available set an upper limit to the number of molecules that can be detected simultaneously with common microscopy methods. Spectral imaging and linear unmixing extends the possibilities to discriminate distinct fluorophores with highly overlapping emission spectra and thus the possibilities of multicolor imaging. This method also offers advantages for fast multicolor time-lapse microscopy and fluorescence resonance energy transfer measurements in living samples. Here we discuss recent progress on the technical implementation of the method, its limitations and applications to the imaging of biological samples.     

Four-color, 4-D time-lapse confocal imaging of chick embryos

Our understanding of the molecular mechanisms that direct cell motility, cell division, and cell shaping has benefited from innovations in cell labeling and the ability to resolve intracellular dynamics with multispectral, high-resolution imaging. However, due to difficulties with in vivo cell marking and monitoring, most studies have been restricted to fixed tissue or cells in culture. Here, we report the delivery of multiple (up to four), multicolor fluorescent protein (FP) constructs and four-dimensional (4-D), multispectral time-lapse confocal imaging of cell movements in living chick embryos. Cell cytoskeletal components are fluorescently tagged after microinjection and electroporation of a cocktail of FP constructs into specific regions of chick embryos. We tested 11 different FP constructs in various two-, three-, and four-color combinations using multispectral imaging and linear unmixing to limit the crosstalk between different emission spectra. We monitored intracellular dynamics in individual multicolored migrating cells in vivo and developed a set of advantageous imaging parameters for 4-D time-lapse confocal microscopy. We find that the number of four-color labeled cells in a typical embryo is approximately 10% of the total number of fluorescently labeled cells; this value consistently increases showing that approximately 50% of the total labeled cells have only one-color. We find that multicolored cells are photostable for time-lapses of approximately 2-3 h. Thus, cell labeling with up to four FP color schemes combined with multispectral, 4-D confocal time-lapse imaging offers a powerful tool to simultaneously analyze cellular and molecular dynamics during chick embryogenesis.