Understanding the function of proteins within the context of the intact living cell is one of the main aims of contemporary biological research. The visualisation of specific cellular events has been greatly enhanced by modern microscopy. In addition to qualitatively viewing the images collected with a microscope, quantitative information can be gleaned from the images. The collection of meaningful measurements has been greatly facilitated by the advent of digital image processing. Subtle changes in intensity of probes of biochemical events can be detected with sensitive digital detectors. These technological advancements have allowed insight into the spatial aspects of molecular mechanisms.
Relatively simple measurements include counting features within a 2D image or measuring areas and lengths. Measurements of depth and volume can be made in 3D, 4D and 5D datasets. Images can be calibrated by collecting an image of a calibration grid at the same settings of the microscope that was used for collecting the images during the experiment. Many computer image-processing systems allow for a calibration factor to be added into the program, and all subsequent measurements will then be comparable.
The rapid development of fluorescence microscopy, together with digital imaging and, above all, the development of fluorescent probes of biological activity, such as GFP and DsRed, have enabled a new level of sophistication into quantitative imaging. Most of the measurements are based on the ability to accurately measure the bright ness of and the wavelength emitted from a fluorescent probe within a sample using a digital imaging system. This is also the basis of flow cytometry, which measures the individual brightness of a population of cells as they pass through a laser beam. Cells can be sorted into different populations using a related technique, fluorescence-activated cell sorting.
The brightness of the fluorescence emitted by the probe can be calibrated to the amount of probe present at any given location in the cell at high resolution. For example, the concentration of calcium is measured in different regions of living embryos using calcium indicator dyes, such as fluo-3, whose fluorescence increases in proportion to the amount of free calcium in the cell (Figure 1). Many probes have been developed for making such measurements in living tissues. Controls are a necessary part of such measurements, since photobleaching and various dye artefacts during the experiment can obscure the true measurements. This can be achieved by staining the sample with two ion-sensitive dyes, and comparing their measured brightness during the experiment. Such measurements are usually expressed as ratios (ratio imaging) and provide a control for dye loading problems, photobleaching and instrument variation.
Fig1. Calcium imaging in living cells. A fertilisation-induced calcium wave in the egg of the starfish. The egg was microinjected with the calcium-sensitive fluorescent dye fluo-3 and subsequently fertilised by the addition of sperm during observation using time-lapse confocal microscopy with a 40× water immersion lens and a laser-scanning confocal microscope. An optical section located near the egg equator was collected every 4 seconds using the normal scan mode accumulated for two frames. Afterwards, the images were corrected for offset and subjected to ratio generation by linearly dividing the initial pre-fertilisation image into each successive frame of the time-lapse run. After ratio generation, the images were prepared as a montage and rendered with a pseudo-colour look-up table in which blue regions represent low ratios of free calcium levels, and red areas depict high ratios of free calcium levels. Note that the wave sweeps through the entire ooplasm, rather than being cortically restricted. (Image kindly provided by Steve Stricker, University of New Mexico, USA.)
Fluorescence recovery after photobleaching (FRAP) uses the high light flux from a laser to locally destroy fluorophores labelling the macro molecules to create a bleached zone (photobleaching). The observation and recording of the subsequent movement of undamaged fluorophores into the bleached zone gives a measure of molecular mobility. This enables biochemical analysis within the living cell. A second technique related to FRAP, photoactivation, uses a probe whose fluorescence can be induced by a flash of short-wavelength light (typically UV). The method depends upon caged fluorescent probes that are locally activated (uncaged) by a pulse of UV light. Alternatively, variants of GFP can be expressed in cells and selectively photoactivated. The activated probe is imaged using a longer wavelength of light. Here, the signal-to-noise ratio of the images can be better than that for photo bleaching experiments.
A third method, fluorescence speckle microscopy was discovered as a chance observation while microinjecting fluorescently labelled proteins into living cells at a very low concentration. Since the fluorescently labelled protein participates in the same functions as the endogenous (and thus non-labelled) protein, the distribution of the fluorophore is ‘diluted’ and, when viewed in the microscope, structures inside cells that have been labelled in this way have a speckled appearance. The dark regions act as fiduciary marks for the observation of dynamics.
Fluorescence resonance energy transfer (FRET), is a fluorescence-based method that can take fluorescence microscopy past the theoretical resolution limit of the light microscope, allowing the observation of protein–protein interactions in vivo (Figure2). FRET occurs between two fluorophores when the emission of the first one (the donor) serves as the excitation source for the second one (the acceptor). This will only occur when two fluorophore molecules are very close to one another, at a distance of 60 Å or less.
Fig2. Fluorescence resonance energy transfer (FRET). In the situation shown in (a), the donor cyan fluorescent protein (CFP) and the acceptor yellow fluorescent protein (YFP) are not close enough for FRET to occur (more than 6 nm separation). Therefore, excitation with light of a wavelength of 430 nm (blue) results in the emission of light at 490 nm (green) – the fluorescence emission of CFP. In the situation shown in (b), CFP and YFP are close enough for energy transfer to occur (closer than 60 Å). Here, excitation with light at 430 nm (blue) results in fluorescence emission of CFP (490 nm; green) and of YFP (520 nm; red).
As an example, the complex formation between two proteins can be monitored using FRET. One of the two proteins is tagged with cyan fluorescent protein ( CFP), the other with yellow fluorescent protein (YFP). The excitation wavelength is chosen such that CFP is excited. If the two proteins of interest are spatially close, so are their fusion partners, and energy is transferred from the excited CFP to YFP. Fluorescence emission is then observed at the emission wavelength of YFP.
A more complex technique, fluorescence lifetime imaging (FLIM) measures the amount of time a fluorophore is fluorescent after excitation with a 10 ns pulse of laser light. FLIM is a method used for detecting multiple fluorophores with different fluorescent lifetimes and overlapping emission spectra.
