Types of Imaging, Part 2: An Overview of Fluorescence Microscopy
2012; Wiley; Volume: 295; Issue: 10 Linguagem: Inglês
10.1002/ar.22548
ISSN1932-8494
Autores Tópico(s)Advanced Biosensing Techniques and Applications
ResumoThis is the second article in a series covering different types of microscopy (http://www.microscopy.info/Microscopy/ResourcesByCat/8). Fluorescence optical microscopy is a powerful imaging tool in biology used to collect spatial and functional information about both endogenous autofluorescent and exogenously labeled molecules and structures. Fluorescent molecules enable researchers to obtain spatial and functional information. The two main sources of light are mercury vapor or xenon arc lamps with an excitation filter, or lasers. In the fluorescence microscope, the high-energy light irradiates and excites fluorophores in the specimen. The excited fluorophore then emits lower energy fluorescent light. Filters separate the lower energy emitted light, which is seen by the eye or other detector. Fluorescence microscopy is the only type of microscopy where the specimen emits its own light (Davidson and Abramowitz, 1999) (http://www.microscopyu.com/). A sample needs to be fluorescent to be suitable for fluorescence microscopy. There are several different ways of creating a fluorescent sample. Most techniques involve labeling with fluorescent stains, and expression of a fluorescent protein is used for biological samples. Additionally, the intrinsic fluorescence of a sample (i.e., autofluorescence) can be used. Fluorescence microscopy is used to investigate live or fixed organisms, tissues, or cells (using in vitro or in vivo fluorochromes; e.g., fluorescein, Alexa Fluors, or DyLight 488) as well as organic and inorganic material. Immunofluorescence is a commonly used technique whereby a fluorescently labeled antibody is bound to its antigen to localize molecules in biological specimens. Fluorescent probes can also be used to examine physiological ion concentrations and pH values that rapidly change in living cells. Fluorochromes have enabled cells and cellular components to be identified with a high specificity among nonfluorescing material. Furthermore, the presence of fluorescing material can be shown with detailed sensitivity using fluorescence microscopy. Application of filter sets in the fluorescence technique can reduce background and provide high-quality images. One limitation of fluorescence microscopy is that fluorophores lose their capacity to fluoresce when illuminated due to photobleaching. Also, although use of fluorescent reporter proteins enables analysis of living cells, cells are prone to phototoxicity, especially when a short wavelength is used. Additionally, fluorescent molecules tend to generate reactive chemical species when being illuminated, increasing the phototoxic effect. A further limitation is that only specific structures that have been fluorescently labeled can be viewed. However, in many biological specimens, autofluorescence is a large problem that excitation/emission filters might not completely get rid of. This article discusses many of the different types of fluorescence microscopy used today. The scope of this article is not to provide a large amount of detail but to overview the most commonly used types. Future articles will go into more depth of particular issues of this type of microscopy. Most of the fluorescence microscopes in use are epifluorescence microscopes. In this type of microscopy, observation and excitation of the fluorescence occur from above the specimen. Fluorescence of the specimen results in emitted light, which is focused on the detector using the same objective used for excitation. Many attachments are available for epifluorescence microscopes that can enhance viewing. This microscopy technique has enabled researchers to identify the cellular and molecular components of each sample with a high amount of specificity (Fig. 1) and to examine impurities and disease conditions. A problem associated with epifluorescence microscopy is that secondary fluorescence is often produced by the specimen through the excited volume and hinders discrimination of features lying in the focal plane (Wilson, 1989). This issue is further compounded by thick specimens (> 2 μm), which often show high-fluorescence emission resulting in fine detail being lost. Epifluorescent image of a wide field image of rat artery in the heart labeled with phalloidin (red, labels actin) and von Willebrand factor (green, endothelial cell marker). Scale bar indicates 20 μm. The image was kindly provided by Dr. David Crossman, the University of Auckland. Because illumination results in fluorescence of the entire specimen thickness, in a two-dimensional (2D) image, more than 90% of the emitted light is out of focus. This can obscure detail and considerably reduce the remaining contrast (Conchello and Lichtman, 2005). Modern confocal microscopes were designed to address this issue. The optical microscope comprises the main part of this system that also includes one or more electronic detectors, multiple laser systems that are combined with wavelength selection devices and a beam scanning assembly, and a computer used for image display, output, processing, and storage (http://www.olympusconfocal.com). Confocal technology is one of the most important achievements in optical microscopy. Confocal microscopy has a number of advantages compared with conventional widefield optical microscopy. The most important advantage is the ability to finely optically section, removing unwanted light above and below the focal plane, providing better contrast and allowing 3D reconstruction by combining the image data from a stack of images using a computer. Confocal scanning microscopy is the most common optical sectioning technique used for fluorescence imaging. Other advantages of confocal microscopy include being able to control depth of field, and the removal or reduction in background information away from the focal plane (Claxton et al., 2010). The basic concept behind the confocal approach is that spatial filtering techniques are used to remove light that is out of focus or glare in specimens when the thickness exceeds the immediate plane of focus. In contrast to epifluorescence microscopes, confocal microscopy excludes secondary fluorescence in regions removed from the focal plane. However, even though there is less degradation of signal, photobleaching is still a significant problem, especially for weaker signals. Resolution is considerably better in confocal microscopy than that in conventional wide field techniques, but resolution is still much less than that of transmission electron microscopy. Therefore, confocal microscopy provides a bridge between these common methods. In modern confocal microscopes, two different techniques used for beam scanning are: single beam scanning and multiple beam scanning. They are described below. In laser scanning confocal microscopy, a focused laser beam controlled by two high-speed oscillating mirrors powered with galvanometer motors is scanned across the region of interest a defined area in a raster pattern. (Claxton et al., 2010). Laser light that is scattered and reflected, and any fluorescent light from the illuminated spot, are received by the objective lens. A beam splitter separates a portion of the light and sends it through a filter that selectively allows emitted but not excitation wavelengths to pass. The filtered light passes a pinhole to a photodetection device, which then transforms the light signal into an electrical signal that is recorded by computer. Advantages of confocal laser scanning microscopes include a programmable sampling density and high resolution. Representative images are shown in Figs. 2 and 3. Laser scanning confocal image of connexin 43 gap junction plaques (green) that are colocalized with astrocytic processes (red) in the fetal sheep brain. The image was kindly proved by Dr. Joanne Davidson and Professor Colin Green, the University of Auckland. Laser scanning confocal image of fly larva expression using (Elav-Gal4) the Chameleon fluo protein calcium sensor. The image was kindly proved by Dr. Thomas Moninger, Central Microscopy Research Facility, University of Iowa, and Dr. Michael Welsh, University of Iowa. Spinning disc confocal microscopes contain a spinning Nipkow disk with an array of pinholes and microlenses (Nakano, 2002). These microscopes usually have arc-discharge lamps for illumination rather than lasers for reduction of damage to specimens and to increase detection of low levels of fluorescence while obtaining real time images. Another useful feature is that they can capture images with an array detector, for example, a charge-coupled device camera system. Laser scanning systems usually have very slow imaging frame rates (e.g., < 3 frames/sec), but spinning-disk confocal microscopes can obtain frame rates greater than 50 per sec. This is an important feature for dynamic observations such as imaging of live cells. Multiphoton fluorescence excitation is a powerful technique for optical sectioning microscopy. Multiphoton microscopes use high-energy lasers. As a result, nonlinear optical effects, such as multiphoton fluorescence can occur because the energy density is sufficiently large at the focal spot. These optical effects can be used to study biological material. Simultaneous multiphoton absorption mostly transpires in the focal region, and therefore, photodamage and photobleaching are confined to the focal plane. With multiphoton fluorescence microscopy, simultaneous absorption of two or three near-infrared (NIR) photons results in 3D resolution, which is similar to standard confocal microscopy (Straub and Stefan, 1998). Multiphoton microscopy is the most convenient approach to minimize bleaching of fluorophores or death of the specimen (Quercioli, 2011). Dye excitation only occurs at the observation point, and none of the fluorescence emission signal is lost. Additionally, the ultrafast laser's long wavelength emission (∼800 nm) results in less damage of living samples. Multiphoton fluorescence microscopy is also useful because of its inbuilt sectioning capability and deep penetration into scattering specimens. Additionally, a pinhole is not required to obtain 3D discrimination, and therefore, the efficiency of fluorescence collection can be enhanced (Piston, 1999). However, it is an expensive technique. Denk et al. (1990) published the first two-photon excited fluorescence imaging of live specimens using an NIR dye laser. Currently, two-photon microscopy is usually performed with expensive pulsed lasers; for example, the mode-locked titanium:sapphire (Ti:Sa) laser provides pico- or subpicosecond pulses at ∼80 MHz. Two-photon excitation occurs when two photons are simultaneously absorbed in a single quantized event. This excitation is produced by a single pulsed laser being focused through the optics of the microscope. The photons become more crowded as the laser beam is focused, and the chance increases for two of them simultaneously interacting with a single fluorophore. This excitation relies on simultaneous absorption and the resultant fluorescence emission is dependent on the square of the excitation intensity. This quadratic dependence is responsible for many advantages related to two-photon excitation microscopy. Very high-laser powers are necessary to produce a large amount of two-photon-excited fluorescence. Two-photon excitation microscopy can be used as an alternative option to confocal microscopy and it has many advantages for 3D imaging. In particular, two-photon excitation microscopy is superior for live cell imaging, particularly in thick, multicellular preparations, including developing embryos, brain slices, and other intact tissue (Piston, 1999). The localization of two-photon excitation to the focal point (see next paragraph) can explain a lot of the advantages over confocal microscopy; no background is present and no pinhole is required. Three-photon excitation functions in a way similar to that of two-photon excitation, except three photons need to interact with the fluorophore at the same time. The quantum mechanics of fluorescence absorption mean that only an ∼10-fold greater photon density is needed for three photon excitation than is required for two-photon absorption. Three photon excitation is useful for extending the area of useful imaging into the deep ultraviolet (UV) (i.e., 720 nm light is used to excite a fluorophore that normally absorbs at 240 nm), which is useful because UV wavelengths below ∼300 nm can cause problems for regular microscope optics. Total internal reflection fluorescence (TIRF) microscopy was developed to overcome the problem of background fluorescence when studying fixed molecules in a fluid environment containing large amounts of unbound molecules (Mattheyses et al., 2010). TIRF microscopy uses the evanescent wave that occurs when light is totally reflected at an interface, typically glasswater. The internally reflected light produces a thin electromagnetic evanescent field in the water with the same wavelength as the incident light. This can excite fluorophores close to the glass surface. Because the evanescent field intensity decays exponentially with the distance from the glass surface, fluorophores that are further from the surface do not become excited, thereby greatly reducing the background noise. TIRF enables thin regions (usually less than 200 nm) of a specimen to be observed. TIRF enables direct monitoring of biomolecules at the single-molecule level in vitro and in living cells (Wazawa and Ueda, 2005). This technique has led to a new era in bioscience called single-molecule nanobioscience. Using TIRF, it is possible to follow biochemical reactions, structural changes, and movements of biomolecules at the level of single molecules in real time, and the results from this technique have given new insights on molecular mechanisms of biomolecules. TIRF is useful for observing fluorophores attached to biomolecules and living cells in aqueous solution near a glass surface. TIRF greatly reduces background noise by limiting the depth of fluorophore excitation at an interface. Other advantages of TIRF include production of high-contrast images of fluorophores near plasma membranes, and reduced cellular photodamage and exposure times (Mattheyses et al., 2010). Fluorescence correlation spectroscopy (FCS) is a versatile technique that operates at the single-molecule level (Kohl and Schwille, 2005). Because of poor signal-to-noise ratios, this technique did not become extensively used until FCS was combined with confocal detection. FCS analyzes very small spontaneous fluctuations in the fluorescence emission of small molecular complexes to study underlying inter- and intramolecular dynamics. FCS is used for the study of chemical kinetics, conformation dynamics, and molecular diffusion in solution and on membranes. Because FCS can be used to observe fluorescent molecules at nanomolar or lower concentrations, it provides quantitative data under physiological conditions and monitors dynamic equilibria in vivo (Fig. 4). FCS time lapse images showing calcium flux in KC-1 (koala cell) cells. Cells were labeled with the calcium indicator fluo-3. The image was kindly proved by Dr. Thomas Moninger, Central Microscopy Research Facility, University of Iowa. In contrast to conventional single molecule setups, FCS does not rely on the intensity of emission, but instead on spontaneous fluctuations of fluorescence intensity caused by deviations from a mean. An FCS setup consists of a laser line, which is reflected into the objective of a microscope using a dichroic mirror. The laser beam is then focused on the sample, which has such a high dilution of fluorescent particles that only a few are within the focal spot. The particles fluoresce when they cross the focal volume. The electronic signal from this process is stored directly as an intensity-versus-time trace that is able to be analyzed later, or it can be computed to generate the autocorrelation directly. Advances in instrument design and optical approaches have enabled FCS to be useful for complex problems that are not solved by other imaging methods. One of the key advantages of FCS is that it is an in situ, nondestructive, and nonperturbing technique. Specific advantages of FCS include a high temporal and spatial resolution and short measurement time (∼msec) (Tian et al., 2011). FCS can be combined with two-photon excitation leading to further advantages, such as excitation of two or more spectrally different dyes with one laser line, and reducing cell damage and sample consumption by photobleaching. However, FCS is possible only within a limited concentration range and is strongly reliant on the correctness of the models used in evaluation of data (Enderlein et al., 2004). Novel techniques have been developed as extensions of FCS to overcome these problems, such as fluorescence cross-correlation spectroscopy, numerical FCS, fluorescence lifetime correlation spectroscopy combined FCS with time-correlated single-photon counting, and TIRF illumination in conjugation with FCS. Loss of fluorescent signal after the breakdown of fluorescent molecules following their interaction with oxygen is termed photobleaching. In most experiments with live cells, imaging conditions are optimized to reduce photobleaching, as described above, but deliberate photobleaching can be a useful technique. Fluorescence recovery after photobleaching (FRAP) allows investigation of the diffusion and motion of biological macromolecules in a uniformly labeled region of interest. First, fluorescently tagged molecules are observed at low light intensity to define a region of interest. The molecules in the area are then photobleached using high-intensity light. Emission intensity is monitored as the bleached dye diffuses out and new dye diffuses in until a uniform intensity is restored. FRAP is commonly used to study membrane diffusion, membrane–protein interactions, and protein binding (http://www.trinklelab.com/pubpdf/04/rms04.pdf). The basic set-up comprises an optical microscope, a light source, and a fluorescent probe. The green fluorescent protein found in the jellyfish Aequorea victoria has led to a revolution in cell and molecular cell biology, which has boosted the application of FRAP. Confocal microscopes are commonly used for FRAP experiments (Houtsmuller, 2005). Fluorescence loss in photobleaching (FLIP) is a related technique that is used to determine the decrease of fluorescence in a defined area that lies adjacent to a photobleached area. FLIP is similar to FRAP in that this technique can be used to investigate molecular mobility and dynamics in live cells. FRAP has a number of variations, including FRAP for immobilization measurement (FRAP-FIM; developed to study the immobilization of nuclear repair factors), combined strip-FRAP and FLIP-FRAP (used to study transient immobilization), and inverse FRAP (developed to estimate the rate of dissociation of molecules from the nucleolus). Difficulty may arise when fluorophore levels are low requiring monitoring at relatively high excitation intensity, which may lead to erroneous results. Problems may also occur when proteins under observation accumulate in one or multiple foci or when they exchange with neighboring areas of the cell. Another potential issue is that many fluorescent proteins rapidly switch between a dark nonfluorescent state and a fluorescent state. Advantages of FRAP are that it is noninvasive and can be easier and faster than approaches such as dye transfer by microinjection. In fluorescence lifetime imaging (FLIM), the nanosecond decay kinetics of the excited state of chromophores are spatially mapped by a microscope equipped with a detector capable of high-frequency modulation or fast gating (Bastiaens and Squire, 1999; Van Munster and Gadella, 2005). This can involve a single decay time to an entire decay profile, in two or three dimensions. Lifetime is independent of dye concentration, photobleaching, light scattering and excitation light intensity. Factors affecting the fluorescence lifetime include ion intensity, hydrophobic properties, oxygen concentration, molecular binding, and molecular interaction by energy transfer when two proteins approach each other. Therefore, FLIM makes fluorescence resonance energy transfer (FRET) possible. There are many types of FLIM instruments but they all fall into two categories based on whether measurements are made using the time-domain or the frequency-domain method. They are mostly used for biomedical applications. The properties of fluorescence lifetimes enable determination of the molecular environment of labeled macromolecules inside cells. Imaging of fluorescence lifetimes allows biochemical reactions to be examined within the cell. An advantage of FLIM is that lifetime depends on excited-state reactions but is independent of light-path length and chromophore concentration, which are difficult to control inside cells. FLIM acquisition is also fast enough (hundreds of milliseconds to seconds) to allow measurements in living cells. FLIM applied to fixed and archived tissues can be used to determine the earlier functional state of proteins involved in the pathology of diseased tissues. The nanosecond fluorescence lifetimes enable an extra spectroscopic dimension as well as a steady-state fluorescence image. This enables the resolution of parameters reporting on the activation state of proteins in situ without disrupting the cellular components. The lifetime of the fluorophore signal is used to create the image in FLIM, which is advantageous for minimizing the effect of photon scattering in thick layers of a sample. FRET is a distance-dependent interaction that occurs between the electronic excited states of two dye molecules, where excitation is transferred from a donor to an acceptor molecule and there is no emission of photons (Clegg, 1992). The dependence of the energy transfer efficiency on the donor-acceptor separation enables this phenomenon to be used for the study of cell component interactions. Optical microscopy combined with FRET allows obtainment of quantitative temporal and spatial information regarding the binding and interaction of proteins, lipids, enzymes, DNA, and RNA in vivo. FRET improves spatial resolution because it relies on the close physical interaction of two fluorophores (called the donor and the acceptor). Methods routinely used for measuring FRET include FLIM-FRET, acceptor photobleaching, and sensitized emission (http://www.invitrogen.com/site/us/en/home/References/Molecular-Probes-The-Handbook/Technical-Notes-and-Product-Highlights/Fluorescence-Resonance-Energy-Transfer-FRET.html). Advantages of FRET include a spatial resolution of the fluorescence microscope to below 10 nm. FRET can be used as a contrast mechanism, and colocalization of proteins and other molecules can then be imaged with a spatial resolution that exceeds that of conventional optical microscopy. Limitations of FRET are that steady-state FRET microscopy measurements can be affected by distortion, which has to be corrected, and the number of donor/acceptor pairs suffers significant cross talk (Gordon et al., 1998). In the 1990s, new microscopies revolutionized the imaging field, and broke the lateral resolution diffraction limit for the first time. These techniques are collectively called super-resolution imaging techniques (Stepanenko et al., 2011). All super-resolution imaging methods have successfully addressed different biological problems. A comprehensive description of all the different types of super-resolution imaging is beyond the scope of this article. However, some of the most common types are briefly discussed below. There are two types of super-resolution imaging (Fernandez-Suarez and Ting, 2008). Near-field super-resolution imaging overcomes the diffraction limit by removal of the lenses, thus eliminating the need to focus. Although near-field super-resolution imaging is used to study the nanoscale organization of several membrane proteins, this type of imaging is technically difficult. The aperture probe is difficult to construct, and the requirement for feedback to keep a constant distance from an irregular sample restricts the speed of image acquisition. Additionally, near-field super-resolution imaging is not suitable for intracellular imaging. Therefore, these factors have limited the use of near-field super-resolution imaging in cell biology. The second type of super-resolution imaging is far-field super-resolution imaging (Stepanenko et al., 2011, Fernandez-Suarez and Ting, 2008). In contrast to near-field super-resolution imaging, far-field microscopy uses lenses, which are placed at a distance from the sample. To overcome the diffraction limit, it is important to spatially and/or temporally modulate the transition between two molecular states of a fluorophore. Some of the techniques obtain super resolution by narrowing the point spread function of an image containing many fluorophores. These techniques include ground-state depletion, stimulated emission depletion, and saturated structured illumination microscopy as well as its combination with I5M (I5S). Other techniques can detect single molecules and are based on the concept that a single emitter can be accurately localized if a sufficient number of photons is obtained. Examples of these techniques include stochastic optical reconstruction microscopy (STORM), photoactivated localization microscopy (PALM) (Fig. 5), and fluorescence photoactivation localization microscopy. Direct STORM (variant of PALM/STORM) super-resolution image depicts clusters of Ryanodine receptors adjacent to the surface membrane of rat ventricular myocytes imaged using localization microscopy, demonstrating the resolution enhancement available with the technique. Outside the circle, clusters are shown as they appear at conventional, diffraction limited, resolution (∼30 nm resolution within the circle). The scale bar = 2 μm. The image was kindly provided by Dr. David Baddeley, the University of Auckland. I wish to thank the following people for the images in this article: Dr. Joanne Davidson, Professor Colin Green, Dr. David Crossman, Dr. David Baddeley, Dr. Thomas Moninger and Dr. Michael Welsh. I also thank Jacqui Ross for advice on confocal microscopy.
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