Live-cell Imaging Techniques

Visualizing the Molecular Dynamics of Life

February 24, 2012

The understanding of complex and/or fast cellular dynamics is an important step for exploring biological processes. Therefore, today’s life science research is increasingly focusing on dynamic processes like cell migration, morphological changes of cells, organs or whole animals and physiological (e.g. changes of intracellular ion composition) events in living specimens in real time. One approach to address these challenging demands is to employ certain optical methods that are collectively called live-cell imaging. Live-cell imaging allows investigation of dynamical processes of living cells instead of giving a “snapshot” of a cell’s current state - it turns snapshots into movies. Live-cell imaging provides spatial and temporal information of dynamic events in single cells, cellular networks (in situ) or even whole organisms (in vivo). These features make live-cell imaging a requisite technique for addressing questions in cell biology, cancer research, developmental biology and neuroscience.

In recent years, substantial advances in electronics, optics and molecular biology have made live-cell imaging easily accessible for scientists.

Methods employed for live-cell imaging

The range of microscopic techniques applied for live-cell imaging is also extremely wide. Often, the growth of cells, cell aggregates or cell movement is observed over time using compound microscopes and contrasting methods like phase contrast and differential interference contrast (DIC). Also, time- lapse imaging of larger specimens, e.g. developing zebrafish embryos, is widely performed, usually with stereo microscopes or macroscopes. Advanced fluorescence techniques have become more and more important in the last decades. The rapid increase in the application of confocal microscopy has changed the perspective in biological investigation from planar to the third dimension. Here’s a short overview of commonly applied techniques.

Ion imaging – observing changes in ion concentrations

A commonly performed method is ion imaging (calcium, chloride, magnesium) using either fluorescent dyes or proteins especially designed to change their emission behavior upon calcium binding. This allows researchers to observe dynamic changes in the ion concentrations of cells. As the ion composition in the cytosol of cells determines many crucial functions of cells like excitability of neurons, gene transcription and cell movement (just to name a few) the regulation of intracellular ions in both spatial and temporal terms is of major interest for life science research. Also, the imaging of intracellular pH-levels or voltage is possible with special fluorescent dyes. A special technique for imaging changes in ion levels, pH levels or voltage are ratiometric imaging methods. These allow exact determination of e.g. the intracellular calcium concentration instead of monitoring relative changes as it is done in non-ratiometric methods.

FRET – quantifying protein-protein interactions

To detect dynamic protein interactions FRET (Förster resonance energy transfer) and BRET (bioluminescence resonance energy transfer) events can be imaged in live-cell experiments. FRET is a useful tool for quantifying molecular dynamics, such as protein-protein interactions, protein-DNA interactions, and protein conformational changes. FRET imaging normally uses derivatives of GFP (green fluorescent proteins), particularly CFP and YFP (cyan and yellow fluorescent protein, respectively), which are each attached to proteins of interest using molecular biology methods. Then the CFP molecule is excited with fluorescent light. As soon as the proteins of interest are in close spatial proximity (<20 nm) the CFP will act as a donor and transfer the energy which is emitted in the form of light to the YFP, which acts as an acceptor. The researcher will observe a shift from blue fluorescence emitted from the CFP to yellow fluorescence emitted from the YFP. In the case of BRET the donor is a bioluminescent molecule (e.g. luciferase derivatives) that acts as the donor and, like in FRET, GFP derivatives act as acceptor.

Fig. 1: Confocal live-cell image of Arabidopsis; endoplasmatic reticulum: GFP labeled in green, autofluorescent chloroplasts in red, transmission in blue. Based on such an image e.g. FRET or FRAP analysis can be done.

FRAP – monitoring protein and vesicle trafficking

A method often applied for monitoring protein or vesicle trafficking is fluorescence recovery after photo-bleaching (FRAP). Here a fluorescent protein (usually GFP) is attached to a protein of interest (i.e. a protein whose movement is to be monitored). Usually, the whole cell is initially fluorescent as the protein might be abundant in the whole cell. Then a certain region of the cell, often cell processes like axons or dendrites in neuronal cells, is exposed to high intensities of light (usually laser light) and the fluorescence in that particular region is destroyed (bleached). As the protein of interest is moving, proteins from other parts of the cell will reinvade the bleached region at a certain speed and the fluorescence in the bleached region recovers. This can give researchers insight into intracellular transport dynamics.

TIRF – observing processes close to the cell membrane

A special technique for observing events that are located in or close to the plasma membrane of a cell is TIRF (total internal reflection) microscopy. By using an evanescent field for fluorochrome excitation which only penetrates the cell 60-250 nm, TIRF microscopy provides an unmatched z-resolution which enables the imaging of events occurring in or close to the plasma membrane (e.g. molecule transport to the plasma membrane) without being outshone by fluorescence from molecules inside the cell.

TIRF images: Transport of Galectin-3 vesicles close to the membrane along actin filaments.

Fig. 2A: Overview image with epifluorescence,
Fig. 2B: Overview image with TIRF, labeled section is shown in C,
Fig. 2C: Time sequence of section of TIRF; time given in seconds. A Galectin-3 vesicle (marked with YFP) close to the membrane (arrow) is first transported along an actin filament (from the bottom upwards), switches to another filament (88 s), moves to the left (109 s), is transported to the right again, switches filament again and is then transported upwards (246 s).
YFP: red; CFP: green; scale of overview images: 20 µm; section: 6 µm; penetration depth TIRF: 110 nm. Courtesy of Ralf Jacob, University of Marburg, Germany

Photoactivation – monitoring gene expression and protein transport

A very recently developed method called photoactivation selectively labels certain regions or areas of interest within a cell or a whole organism. For photoactivation especially designed dyes or fluorescent proteins like photoactivatable green fluorescent protein (paGFP) or Kaede are used. These fluorophores are not fluorescent in their normal state. However, after illumination with light of certain wavelengths, these fluorophores can be activated to fluoresce like conventional fluorophores. In many cases, these proteins are genetically fused to certain proteins of interest whose expression or transport can then be monitored. Methods like FRAP or particle tracking can then be applied to further investigate the protein of interest.

MPE – investigating processes in depth

Modern biological research demands real in vivo investigation to complement the “quasi-in-vivo”, in cell culture experiments. But it is difficult to investigate processes occurring inside living organisms like a mouse. Multiphoton excitation (MPE) microscopy enables deeper penetration into tissue, because the near-infrared excitation light, having a longer wavelength, is less scattered compared to the short-wavelength light used for single-photon excitation. The non-linear nature of the MPE technique restricts photobleaching and phototoxicity to the area in the focus. This is tremendously beneficial for long-term investigation, because both the fluorescent proteins and the organism suffer from these problems. Using proper preparation and microscopic setup, imaging up to a few millimeters into the tissue has been reported. The precise localization of the excitation makes it suitable for photon manipulation as well.  This method has found wide application in neurobiology.

STED – studying cellular dynamics at nanometer scale

Stimulated emission depletion (STED) microscopy allows scientists to investigate structures beyond the optical resolution limit. This technique uses the property of fluorescent dyes, being stimulated to emission, to eliminate detectable signals. Intracellular structures down to 50-70 nm have been successfully imaged. Increased resolution is very important for investigating small intracellular structures. Especially for those who want to look into colocalization events, the improvement of resolution could produce a more realistic result. The stochastic independence of STED enables very fast imaging compared to other super-resolution techniques. Video rate STED-acquisition has been achieved, which allows cellular dynamics to be studied in real time.

Fig. 3: STED live-cell imaging with snap-tag technology: Vero cells, structure: EB3,  transient transfection; label: Oregon Green 488.

FLIM – spatial measurements in living cells

Fluorescent lifetime imaging has the advantage that the data is not dependent on the intensity of the signal. Therefore it is not influenced by common artifacts like photobleaching and concentration variation. Using time-correlated single photon counting, FLIM images are reconstructed from data of single molecule detection. The minimal change of the fluorescence lifetime in sub-nanoseconds can be registered and analyzed. The method is versatile for investigation of any kinds of extra- and intracellular environmental modification which lead to an alteration of the fluorescence lifetime. FLIM- based FRET analysis is insensitive to the intensity of the emission and thereby increases the precision of the quantitative data.

CARS and SRS – label-free method using vibrational contrast

Almost all of the fluorescent imaging methods in living cells are based on genetic expression of fluorescent proteins. This involves substantial technical effort and considerable expense. In addition, the expression of external genes may change the microenvironment, which leads to variation of data from the physiological reality. Coherent anti-Stokes Raman Scattering (CARS) microscopy and stimulated Raman Scattering (SRS) microscopy are non-linear confocal methods which are not dependent on fluorescent dyes. These label-free methods image the vibration states of specific chemical bonds in the sample. The accumulation of specific chemical bonds in living organisms, such as the lipid in myelin around the axons, can be imaged in high resolution and excellent signal-to-noise quality, without any need for staining.

The future is quantitative

Biological research has left the age of descriptive investigation and entered an era of quantitative analysis. New live cell imaging techniques are evolving in the direction of higher resolution, both in space and time. Technological developments are now concentrating on the quantitative study of single molecules in the nanometer range and molecular reactions as short as a few picoseconds.

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