Fluorescence microscopy—A historical and technical perspective

Authors

  • Malte Renz

    Corresponding author
    1. Eunice Kennedy Shriver Institute of Child Health and Human Development, National Institutes of Health, Bethesda, Maryland 20892
    • Department of Obstetrics & Gynecology and Women's Health, Albert Einstein College of Medicine, Bronx, New York 10461
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Correspondence to: Department of Obstetrics & Gynecology and Women's Health, Albert Einstein College of Medicine, Bronx, NY 10461, USA. E-mail: mrenz@montefiore.org

Abstract

For a little more than a century, fluorescence microscopy has been an essential source of major discoveries in cell biology. Recent developments improved both visualization and quantification by fluorescence microscopy imaging and established a methodology of fluorescence microscopy. By outlining basic principles and their historical development, I seek to provide insight into and understanding of the ever-growing tools of fluorescence microscopy. Thereby, this synopsis may help the interested researcher to choose a fluorescence microscopic method capable of addressing a specific scientific question. © 2013 International Society for Advancement of Cytometry

In 1845, Sir Frederik William Herschel noted that a quinine solution, although itself colorless and transparent, exhibits a “vivid and beautiful celestial blue color,” when illuminated and observed under certain incidences of sunlight [1]. This is the first reported observation of fluorescence. In a quinine solution like tonic water (Fig. 1), the ultraviolet light from the sun excites quinine to emit blue light, most apparent when observed at a right angle relative to the incident sunlight. In 1852, George Stokes described Herschel's observation of fluorescence in greater detail [2]. Since Ernst Abbe demonstrated the limitations of microscopy using transmitted light [3], expectations were high for the implementation of fluorescence into microscopy. According to Helmholtz (1874), it was to be expected that an image can be better differentiated and fine structures are more easily discernible if the object itself emits light [4]. Baum even claimed that self-illuminating objects were not subject to diffraction [5]. Later in this article (see “Super-Resolution Imaging” section), I will refer back to Baum's postulation. At the beginning of the 20th century, the companies Carl Zeiss and Carl Reichert realized the first fluorescence microscopes [6-8]. For the examination of living organisms which may be big and opaque, Ellinger and Hirt devised “intravital microscopes” [9] using incident instead of transmitted light. They treated living organisms with fluorescent substances to place a source of light in the organism itself, used UV-light for illumination, and interposed filters between objective and eyepiece which reflected the exciting rays and transmitted the red-shifted fluorescent light. The approach Ellinger and Hirt took reminds in principle much of modern fluorescence microscopy. Certainly, the ways of labeling a specimen expanded over the decades; in the early 1940s, fluorescent antibody labeling was developed [10], since the early 1990s the cloning of the green fluorescent protein (GFP) [11, 12] and development of spectral variants [13] permit to label proteins of interest specifically by genetic encoding. Also, instrumentation and techniques have evolved considerably since the first realizations of fluorescence microscopy. Here, a synopsis will be provided of current instrumentations and techniques using fluorescence for obtaining images of cells, organelles, and proteins, and for extracting quantitative information about molecular dynamics and interactions. I will focus on basic principles and their historical development (application examples can be found in the literature [14-18]).

Figure 1.

(Ultra-) Violet light makes a colorless quinine solution such as tonic water fluoresce light blue. As Frederik William Herschel and George Stokes described, this phenomenon can be especially observed when viewed at a right angle relative to the incident light. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

This synopsis has two parts. The first part addresses fluorescence microscopy techniques for diffraction-limited imaging and methods of its quantification. The second part is dedicated to the recent developments of super-resolution microscopy. Instead of attempting to be encyclopedic, I will focus on developments, which may be considered major achievements in fluorescence microscopy fully aware that this can only be a selection and many important developments may remain unconsidered and undiscussed.

Diffraction-Limited Imaging

Because visualization has been an essential source for major discoveries in biology, researchers have always strived to enhance the quality of obtained fluorescence microscopic images. To this end, two general principles have been experimentally realized: selective detection and selective excitation of fluorescent molecules in a specimen. The techniques that will be discussed first, namely, confocal microscopy, spinning disk microscopy, and 4 pi microscopy, are illuminating a biological specimen in its entire depth while they are selectively detecting fluorescent molecules in only a thin image plane. Two-photon excitation, total internal reflection fluorescence microscopy (TIRF-M), and selective plane illumination microscopy (SPIM), on the other hand, are selectively exciting fluorescent dyes in a defined spot or thin plane of the biological specimen.

To improve the resolution of fluorescence images, Marvin Minsky proposed the introduction of pinhole apertures into the beam path of a microscopic set-up; Minsky placed one pinhole in front of the light source and one in front of the detector. The light emanating from the first pinhole is focused by a lens upon the specimen to produce a point; a second lens focuses the illuminated specimen point upon a second pinhole aperture in front of the detector; because of these two lenses, Minsky called his Microscopy Apparatus “double focusing system.” With this tool, patented in 1961 [19], Minsky tried to obtain better images of the stained Golgi apparatus in thick brain tissue blocks. The “double focusing system” is the basis of modern confocal microscopes [20-22]. Here, instead of light from an electric bulb as in the microscopy apparatus, laser light is focused by the objective lens to a tight diffraction-limited spot (Fig. 2A). The fluorescence light from the focus of the excitation light is back projected. Because light source and detector cannot be in the same place, a second image plane is created by the use of a dichroic beam splitter. This wavelength dependent beam splitter reflects the excitation light, but transmits the red-shifted fluorescence light. The pinhole aperture placed in the second image plane blocks scattered and out-of-focus fluorescence light. There is no pinhole in the excitation pathway. The beam waist of the focused laser beam and pinhole aperture define the ellipsoidal confocal detection volume (see gray area in Fig. 2A inset). By blocking out out-of-focus light (orange lines in schematic), the axial resolution is significantly improved and optical sectioning of thick samples is made possible. The point-like excitation light is scanned across the specimen and thereby an image is being built up point by point. Image acquisition is, therefore, a relatively slow process.

Figure 2.

Schematic beam paths for fluorescence microscopy. (A) + (B) show microscope designs taking advantage of selectively detecting fluorophores in a small region of the biological specimen, whereas (C) + (D) illustrate microscopes using selective excitation of fluorophores in a confined region of the biological specimen. A: Confocal microscope. Based on Marvin Minsky's “double focusing system,” laser light is focused by the objective lens to a tight diffraction-limited spot. The fluorescence light from the focus of the excitation light is back projected. Because light source and detector cannot be in the same place, a second image plane is created by the use of a dichroic beam splitter. This wavelength dependent beam splitter reflects the excitation light, but transmits the red-shifted fluorescence light. The pinhole aperture placed in the second image plane blocks scattered and out-of-focus fluorescence light. As shown in the inset, the beam waist of the focused beam and pinhole aperture define the ellipsoidal confocal detection volume (gray area, ideally 200–250 nm laterally and 500–800 nm axially). By blocking out out-of-focus light (orange lines in schematic), especially the axial resolution is improved. The point like excitation light is scanned across the specimen and thereby an image is being built up point by point. B: Spinning disk microscope. A spread laser beam is passed through a rotating disc holding an array of microlenses which focus the laser light onto the pinholes of a second disk. Thereby, many spots in the biological specimen are illuminated simultaneously. This at the rapid rotation of the two disks allows for higher frame rates. The emitted fluorescence light is focused onto another or the same pinhole on the disk comprising an array of pinholes. As in a confocal microscope, out-of-focus light is blocked out. C: TIRF Microscope. In the objective method, the lateralized laser beam is being projected in a sufficiently oblique angle onto the interface of for example glass and water. When a light beam from a transparent medium of high refractivity, e.g., glass, hits a transparent medium of low refractivity, e.g., water, it will be deflected or totally reflected depending upon its angle of incidence. If the light beam gets totally reflected, an electromagnetic field of the same wavelength is induced. Its intensity decays exponentially with distance from the surface; it disappears, is evanescent. Therefore, it can only excite fluorophores in about 150-nm distance from the surface. D: Selective plane illumination microscope. The excitation light is focused by a cylindrical lens to a sheet of light that only irradiates the focal plane of the detecting optics. Thereby, only what is observed is illuminated resulting in less photobleaching and phototoxicity. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

Spinning disk microscopy, a variant of confocal microscopy, was developed to reduce image acquisition time and exposure to laser light. As shown in Figure 2B, a spread laser beam is passed through a rotating disk comprising a series of pinhole apertures (Nipkow [23]/Petráň disk [24]). Thereby, many spots in the biological specimen are illuminated simultaneously. This and the rapid disk rotation allowed for higher frame rates. The emitted fluorescence light is focused onto another or the same pinhole on the disk. Because the pinholes occupy just a small fraction of the disk, most of the light is blocked out making the use of the illumination light rather ineffective. Recently, Yokogawa Electric Japan introduced another disk holding an array of microlenses which focuses the laser light onto the pinholes and thereby increases the illumination efficiency roughly 10 times [25].

4 Pi Microscopy, originally introduced as variant of confocal microscopy to increase further especially the axial resolution, uses two identical perfectly aligned juxtaposed objectives and creates excitation light at a common focal plane. Resulting interference, destructive and constructive, reduces axial resolution by fourfold to sevenfold and lateral resolution by 1.5-fold. First proposed in 1971 [26], it was first experimentally realized in 1994 [27].

In contrast to confocal microscopy in which fluorophores are excited by one photon, two-photon excitation microscopy relies on the near-simultaneous absorption of two photons. When two photons interact simultaneously with a fluorophore, i.e., within 10−18 s, energy combinations of the two photons can occur and trigger electronic transitions of energy twice that of each single photon. Thereby, two red photons can excite a UV-light electronic transition within a fluorophore. Unlike one-photon excitation, the excitation wavelength is longer than the emission wavelength and separated by some hundreds of nanometers. Based on quantum optics, two-photon absorption was theoretically predicted by Göppert-Mayer in 1931 [28] and experimentally realized in 1961 [29]. Denk et al. combined two-photon excitation with laser scanning microscopy for imaging chromosomes in live cells [30]. To permit simultaneous photon interactions, high photon density is required. Therefore, photons are crowded spatially by focusing the laser light as well as temporally by using pulsed lasers. Two-photon excitation produces fluorescence only at the focal plane where the photon density is sufficiently high. Because two-photon fluorescence falls off as the square of the distance from the focal plane, virtually no out-of-focus light is generated, and thus no pinhole needed. The axial and lateral resolution is not better than a well-aligned confocal microscope. In fact, the longer wavelengths used in two-photon excitation result in a slightly bigger illumination spot. However, because the fluorophores are excited only in the focal plane, there is less out-of-focus photobleaching and less out-of-focus photodamage. Because photons are not being absorbed, they can penetrate deeper into tissue, which has been proven especially useful for intravital tissue imaging. Although most fluorophores work well with two-photon excitation at twice the wavelength of their one-photon absorbance peak, two-photon spectra can be different from one-photon spectra and should be measured to take full advantage of two-photon excitation. On the other hand, simultaneous excitation of differently emitting fluorophores with only one excitation wavelength can be performed as shown for example for DAPI, Bodipy sphingomyelin (Golgi), rhodamine 123 (mito), and pyrene lysophatidylcholine (plasma membrane) [31].

In TIRF-M, only those fluorescent molecules, which are close to a surface, are selectively excited to fluoresce (Fig. 2C). TIRF-M exploits total reflection of oblique light beams at the interface of differentially refractive media and a thereby induced evanescent electromagnetic field. When a light beam from a transparent medium of high refractivity, e.g., glass, hits a transparent medium of low refractivity, e.g., water, it will be deflected or totally reflected depending upon its angle of incidence. If the light beam gets totally reflected, because electromagnetic fields cannot be discontinuous at a boundary, an electromagnetic field of the same wavelength is induced. Its intensity decays exponentially with distance from the surface; it disappears, is evanescent. Therefore, it can only excite fluorophores in about 150-nm distance from the surface. TIR was first introduced by Hirschfeld in 1965 for solid/liquid interfaces [32], Tweet et al. for liquid/air [33], and Carniglia et al. for liquid/solid interfaces [34]. Experimental realizations of TIRF-M [35] are prism- and objective-based approaches. Most of prism-based approaches, however, limit the accessibility of the probe. If the prism is positioned on the other side of the probe, the signal quality may suffer. If it is positioned between objective and probe, problems with the objective's working distance may arise. Given these restraints of the prism method, nowadays the objective method is predominantly used. In case of the objective method, a high numerical aperture (NA) is required to project the lateralized beam in a sufficiently oblique angle onto the interface (see Fig. 2C).

The “ultramicroscope” realized by Siedentopf and Zsigmondy in 1903 used a light sheet to illuminate the sample [36]. This ultramicroscope laid the ground for SPIM. Here, the excitation light is focused by a cylindrical lens to a sheet of light that only irradiates the focal plane of the detecting optics. Thus, irradiated objects are being observed perpendicular to the illumination plane (Fig. 2D). Thereby, only what is observed is illuminated which results in a higher signal-to-noise ratio and less photobleaching and phototoxicity [37]. Small free-floating fluorescent particles [38] and also large objects of up to 500 μm in size as for instance the developing zebrafish embryo have been thus observed [39].

Quantification of Diffraction-Limited Imaging

In the following, methods of fluorescence microcopy are discussed that aim to reveal and analyze what is not obvious in the simple fluorescence image. These methods aim to determine molecular dynamics beyond the steady-state distribution and underlying molecular interactions. I will describe methods to assess molecular dynamics, which address the dynamics of many, only a few, or even single molecules. Also, tools will be discussed to assess molecular interactions in live cells read out on different distance-scales ranging from co-localization within a 300-nm spot to molecular vicinity of only a few nanometers.

Fluorescence recovery after photobleaching (FRAP) or fluorescence photobleaching recovery (FPR) analyzes the ensemble dynamics of many molecules on the timescale of seconds to minutes. FRAP, or FPR, has been introduced in the mid 1970s to determine the mobility of fluorescent molecules in the plasma membrane of live cells [40, 41]. In this technique, the equilibrium of fluorescent molecules is initially perturbed, and it is subsequently registered how the equilibrium recovers over time. Using high laser intensity, fluorescent molecules in a defined region of interest are bleached. Thereby, spatially separated populations of still fluorescent and bleached molecules are generated. Then, the exchange and redistribution of these molecule populations are visualized by taking a picture series with low light illumination (Fig. 3A). After background subtraction, correction for laser fluctuations and bleaching during image acquisition, and normalization to prebleach pictures [42], the increase in fluorescence intensity in the initially bleached area relative to the total cell intensity, provides information about the molecular dynamics. The slope of the recovery curve yields the characteristic recovery time of the underlying process, while the amplitude of the recovery curve helps determine the fraction of molecules immobile on the time scale of the experiment. To extract quantitative data, FRAP curves have been fit to exponential functions [43], or more accurately to a Bessel function [44]. Some related bleaching techniques have been described, including inverse FRAP [45] and fluorescence loss in photobleaching (FLIP) [46]. For FLIP, a defined region is repetitively bleached causing loss of fluorescent molecules that move into the bleached region. Thus, it is especially suitable to prove connectivity of cellular compartments. If bleaching power and frequency are kept constant, FLIP can also provide quantitative information about molecular dynamics. Due to advances in laser scanning microscopy, the development of fluorescent marker proteins and data analysis, photobleaching approaches have been broadly applicable since the mid 1990s [43]. Bulk photoactivation (PA) is a similar while in the experimental set-up complementary approach, which might be advantageous over photobleaching approaches. Photoactivatable or photoswitchable proteins shift their spectral properties in response to irradiation with light of specific wavelength and intensity; that is, they become either bright or change color upon being irradiated [47-50]. The intracellular distribution of these fluorophores can then be monitored over time. PA may be more instantaneous than photobleaching, easier to detect, and is not diluted by concurrent protein synthesis. It also permits the analysis of protein degradation and synthesis.

Figure 3.

Principles of making visible and quantifying what is not obvious in a simple picture of a fluorescent specimen. A: Fluorescence recovery after photobleaching (FRAP) analyzes ensemble dynamics of many molecules. Using high-laser power, the equilibrium of fluorescent molecules is initially perturbed, and it is subsequently registered how the fluorescence equilibrium recovers over time. After background subtraction, correction for laser fluctuations and bleaching during image acquisition, and normalization to prebleach pictures (Fpb), the slope of the fluorescence recovery curve, F(t), yields the characteristic recovery time of the underlying process, while the amplitude of the recovery curve (F) helps determine the fraction of molecules immobile on the time scale of the experiment immobile fraction, IF. B: Fluorescence correlation spectroscopy (FCS) addresses the dynamics of only a few molecules by analyzing the spontaneous intensity fluctuations of a fluorescence signal. These intensity fluctuations are typically generated by a change in the number of fluorescent molecules in a detection volume. Temporal autocorrelation provides a measure of the time-dependent relaxation of the fluctuations to the mean. Thereby, diffusional behavior of different molecular species can be identified. In case of rapidly diffusing molecules, intensity fluctuations increase and decrease rapidly, while slowly diffusing molecules remain longer in observation volume and fluctuations occur more slowly. In addition to the dynamics information, the amplitude of the autocorrelation function, G(τ), is a measure of the number of independently diffusing particles (1/N) in the detection volume. C: Single-particle tracking (SPT) helps discriminate different populations of molecules comprising different modes of motion. In SPT, multiple particles in a field of view are identified; then, the individual particle displacements between successive frames are monitored. D–F: For studying molecular interactions, various methods of co-localization (D), fluorescence cross-correlation spectroscopy (FCCS) (E), or fluorescence resonance energy transfer (FRET) (F) may be used. These approaches help determine interactions at different distance-scales ranging from co-localization within a 300 nm spot to molecular vicinity of only a few nanometers. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

Figure 4.

Principles of super-resolution microscopy. A: Structured illumination (SIM). By passing laser light through a moveable grating, a periodically patterned illumination can be generated and projected onto the sample. When an object that contains fine structures, i.e., high-frequency spatial information, is thus illuminated, coarser interference patterns (Moiré fringes) arise and high-frequency features of the sample are thereby shifted to lower frequencies that are detectable by microscope optics. If multiple images using illumination patterns of different phases and orientations are acquired, a final image of super-resolution can be mathematically reconstructed. B: STED microscopy. The excitation beam producing a diffraction-limited excitation volume is superimposed by a doughnut-shaped laser beam with zero intensity in its center, the STED beam. Fluorophores are excited and in the periphery instantly de-excited using stimulated emission depletion. With a thus sharpened illumination spot, probes are scanned and an image is being built up point by point. C: In PALM and STORM, only a few sparsely distributed fluorophores are stochastically photoactivated to fluoresce (green stars), whereas neighboring molecules remain dark (green open circles). The individual fluorophores are imaged, localized, and bleached. Then, in the next cycle, adjacent fluorophores can be switched on. Merging all obtained single-molecule localizations yields the final super-resolution image. Thus, repeated activation and sampling permit densely expressed fluorescent proteins to be resolved in time, even though they are spatially inseparable when viewed all together at the same time. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

In contrast to aforementioned methods, fluctuation analyses such as fluorescence correlation spectroscopy (FCS) [51] allow to address the dynamics of only a few molecules in the micro- to milli-second range. FCS analyzes the spontaneous intensity fluctuations of a fluorescence signal. These intensity fluctuations are typically generated by a change in the number of fluorescent molecules in the detection volume (Fig. 3B). Numbers of molecules can change by diffusion, i.e., molecules moving in and out of the detection volume, and by photochemical or photophysical reactions, e.g., blinking of fluorescent molecules. Single fluorophores cause distinct fluctuations of the mean fluorescence intensity only when few fluorescent molecules are simultaneously present in the detection volume. Therefore, it is favorable to measure for example cells, which express only a few GFP-tagged proteins. In thermodynamic equilibrium and an open detection volume, the fluorescence intensity fluctuations exhibit variations around a mean, which is determined by the surrounding medium and its thermodynamics. The stochastic character of the fluctuations requires the statistical analysis of multiple fluctuations. Temporal autocorrelation provides a measure of the time-dependent relaxation of the intensity fluctuations to the mean. In case of rapidly diffusing molecules, intensity fluctuations increase and decrease rapidly, while slowly diffusing molecules remain longer in the observation volume and fluctuations occur more slowly. To derive characteristic diffusion and reaction constants, it is essential to choose a theoretical function to be fit with the experimental data which reflects best the experimental conditions, e.g., free three-dimensional diffusion of a single fluorophore species in an approximately ellipsoidal detection volume for analyzing diffusion of proteins in the cytoplasm. Thus, both FCS and FRAP can be used to determine rates of simple transport processes or chemical reactions and their theoretical bases are indeed similar [52]. According to Onsager, the regression of spontaneous microscopic fluctuations back to the equilibrium and the regression of induced macroscopic perturbations of the equilibrium are described by the same phenomenological laws [53]. In addition to the dynamic information, the amplitude of the autocorrelation function in FCS provides the number of independently diffusing particles in the detection volume. From this, the fluorescence per diffusing particle can be determined and thereby, in comparison with the freely diffusing dye, the oligomerization state of the diffusing particles of interest can be estimated for monodisperse systems. Photon counting histogram provides a statistical tool for the in-depth analysis of diffusing molecular species of one and two colors [54, 55]. Variants of FCS move the detection volume relative to the probe as for example realized in scanning FCS [56] and thus shift the resolvable time window to slower processes and reduce photobleaching. Number and brightness analysis [57], image correlation spectroscopy (ICS) [58], and Raster ICS [59, 60] allow fluctuation analyses of entire confocal images and not only a single diffraction-limited spot. Furthermore, fluctuation analyses have been successfully implemented into two-photon excitation microscopy [61], TIRF microscopy [62], and SPIM [63].

To examine the dynamics of single molecules at the plasma membrane with a typical time resolution of milliseconds to seconds, Barak and Webb introduced in the 1980s single-particle tracking (Fig. 3C) using fluorescently labeled low-density lipoprotein [64]. In complex processes, a particle may switch between different modes of motion. Then, single-particle tracking is a versatile tool to discriminate different populations of molecules comprising different modes of motion. In general, SPT approaches include two steps. First, multiple particles in a field of view are identified. Then, the individual particle displacements between successive frames are monitored. Tracking algorithms include cross correlation of subsequent images [65], sum-absolute difference approaches [66], center-of-mass (centroid) determination [67], and Gaussian fitting [68].

For studying molecular interactions, co-localization, fluorescence cross-correlation spectroscopy (FCCS), or fluorescence resonance energy transfer (FRET) may be used. These methods help to determine interactions on different distance-scales. A similar distribution of molecules, i.e., their co-localization, may give a first hint at potential molecular interactions (Fig. 3D). Most co-localization methods consider pixel intensities independent of their position. Creating dot plots of, e.g., red versus green intensities of each pixel gives a good idea about the extent of colocalization and can be quantified with Pearson's correlation coefficient. Because this approach is biased by many low-intensity background pixels, the overlap coefficient as a weighted co-localization coefficient insensitive to coincident small intensities has been used [69]. Co-localization can also be characterized with regard to each channel; e.g., as the proportion of the total intensity in one channel coinciding with nonzero intensity of the other or considering if the intensity in co-localizing pixels tends to be above or below the mean pixel intensity [70-72].

FCCS, a two-color variant of FCS, analyzes the fluctuation in the fluorescence signals of two different fluorophores. The fluorescence signals of a fluorophore are not correlated with themselves as in the case of FCS, rather they are cross correlated with signals of the other fluorophore; the greater is the fraction of observed molecules moving together, the higher is the registered cross-correlation amplitude [73]. FCCS detects the joint movement of distinctly labeled molecules (Fig. 3E) and not, like FRET, their actual distance.

In FRET or Förster resonance energy transfer, energy is transferred from an excited donor molecule to an acceptor molecule (Fig. 3F). FRET is a radiationless quantum mechanical process. Preconditions for the energy to be transferred are proper orientation and distance of the molecules as well as significantly overlapping donor emission and acceptor absorbance spectra. In 1948, Theodor Förster laid the theoretical foundation of FRET [74]. Because the rate of energy transfer is inversely proportional to the sixth power of the donor–acceptor distance, FRET has been mainly applied as a spectroscopic ruler to measure distances between a donor and an acceptor molecule in the range of 1–10 nm [75]. FRET competes with all other de-excitation processes of the donor molecule and results in a reduction of emitted donor photons, the so-called donor quenching, and a corresponding increase in the amount of emitted acceptor photons, the so-called sensitized emission of the acceptor. In acceptor photobleaching [76], the acceptor is photodestructed by intense laser light, and thereby the donor released from quenching which results in an increase in measured donor fluorescence. Because FRET de-excites the donor, the donor molecule spends less time in the excited state and is, therefore, less prone to photobleaching, another phenomenon which has been used to determine FRET efficiency (donor photobleaching [76]). Inversely, the rate of acceptor photobleaching is increased in the presence of an excited donor molecule (FRET-sensitized photobleaching of the acceptor [77, 78]). All these approaches use fluorescence intensity measurements to determine FRET efficiencies. Because FRET, however, reduces the time a donor molecule spends in the excited state, i.e., its fluorescence lifetime, detection of a decreased donor lifetime is also a reliable FRET indicator. In contrast to intensity-based FRET measurements, fluorescence lifetime-based FRET measurements are independent of fluorophore concentrations and insensitive to spectral bleed-through. They require, however, rather sophisticated equipment that provides sufficiently high-temporal resolution in the picosecond range (FRET-FLIM [79]). Contrasting aforementioned hetero-FRET, energy can also be transferred between spectroscopically identical molecules, a phenomenon called homo-FRET. In these chain-like FRET interactions, fluorescence intensity, lifetime, and photobleaching rate of the molecules do not change. Hence, homo-FRET can only be assessed through measurements of fluorescence polarization [80, 81].

Super-Resolution Imaging

Allowing a beam of sunlight to pass through a small aperture in a screen, Francesco Maria Grimaldi noticed that the beam was diffused in the form of a cone. In experiments like these, Grimaldi was the first to describe the phenomena of diffraction and coined the term “diffraction” in 1665 [82]. Diffraction refers to phenomena which occur when waves encounter obstacles and include bending of the waves around obstacles and their spreading past small openings. These phenomena were further characterized by Huygens [83], Young [84], Fresnel [85], and Fraunhofer [86-88] and found to ultimately limit the resolution of lens-based imaging systems. The best focused spot of light that a perfect lens with a circular aperture can make is in its lateral dimension often referred to as Airy Disk (after Airy [89]), while its axial dimension is not round disk-shaped, but rather elongated hourglass-shaped. The size of the aperture and the wavelength of light determine the size of such a spot; the smaller the aperture and the longer the wavelength, the larger the spot size at a given distance. In 1873, Ernst Abbe described the lateral resolution limitations of lens-based optical microscopes as the inability to discern details closer than roughly half the wavelength of the illumination light. The maximal relative angle between different wavefronts leaving the specimen defines the smallest details and periodicity that can be resolved [3]. The reader may allow here a small technical excursion describing different historical concepts of resolution: According to Abbe, the lateral resolution is given by math formula, and the axial resolution by math formula, where λ is the wavelength of light, n the refractive index of the imaging medium, and the term math formula known as NA. Rayleigh described objects as indistinguishable when they are closer than the radius of their Airy disks and redefined the formula for lateral resolution as math formula [90, 91]. For Sparrow, the resolution limit was reached when there is no longer a dip in brightness between two objects (defined as math formula) [92]. Here, the concept of contrast and its importance for resolution become evident; contrast is defined as difference between the maximum intensity of an Airy disk and the minimum intensity between Airy disks of adjacent objects [93].

Because high-frequency spatial information of fine objects is lost, when light propagates a distance larger than its wavelength, one possibility to circumvent the limitations imposed by diffraction is to approximate object and imaging optics such as realized in near-field scanning optical microscopy (NSOM/SNOM [94-98]) and total internal fluorescence reflection microscopy (see above). Recent developments in (far-field) fluorescence microscopy come closer to what Helmholtz and Baum were hoping for a century ago (see “Introduction”). These methods of super-resolution microscopy can be described as illumination- and single-molecule- (or probe-) based. Following the resolution that can be practically achieved, first the illumination-based techniques structured illumination (SIM) and stimulated emission depletion (STED) will be addressed, and then the single-molecule based approaches photoactivated localization microscopy (PALM) and stochastic optical reconstruction microscopy (STORM).

By passing laser light through a moveable grating, a periodically patterned or structured illumination can be generated and projected onto the sample. When an object that contains fine structures, i.e., high-frequency spatial information, is thus illuminated, coarser interference patterns (Moiré fringes) arise and high-frequency features of the sample are thereby shifted to lower frequencies that are detectable by the microscope optics. If multiple images using illumination patterns of different phases and orientations are acquired, a final image of super-resolution can be reconstructed (Fig. 4A). First, realizations resulted in a twofold increase in the lateral resolution [99-101]. Using excitation light modulated also in the z-axis, an additional twofold increase in axial resolution can be achieved [102, 103]. However, the illumination pattern itself is diffraction limited. The summation of the highest spatial frequency in the illumination pattern and highest detectable frequency in the sample gives the spatial frequency that is maximally resolvable and results in a twofold resolution improvement. To disrupt the linear dependence of fluorophore emission on illumination power, saturated structured illumination (SSIM) uses high saturating illumination power and thereby enables resolution of about 50 nm [104, 105]. In structured illumination, almost any fluorophore and fluorescent protein can be used to obtain multicolor images. During the multiple image acquisitions of different phases and orientations, however, the sample needs to be stable in order to avoid imaging and reconstruction artifacts. Nevertheless, live cell measurements have been realized [106, 107].

Stimulated emission depletion microscopy (STED) can also be described as an illumination-based super-resolution technique. In STED, two laser beams are used to illuminate the sample. The excitation beam is superimposed by a doughnut-shaped laser beam with zero intensity in its center, the so-called STED beam. The described spatial illumination pattern, however, is diffraction limited. To achieve super-resolution, a nonlinear fluorophore response is used. In contrast to SSIM, STED microscopy exploits nonlinear fluorophore saturation for de-excitation. An excited fluorescent dye can return to its ground state by spontaneous fluorescence emission or it can be transferred back to the ground state by stimulated emission [108]. Stimulated emission requires relatively intense light pulses of wavelength similar to the emitted fluorescence light of the dye that hit the excited fluorescent dye before spontaneous emission can occur. Even though both laser pulses, excitation and STED beam, are diffraction limited, the STED pulse is modified to feature zero-intensity at its center and strong intensity at the periphery. Thereby, fluorophores are excited by the excitation beam and when located in the periphery of the illuminated spot, they are instantly de-excited by the red doughnut-shaped STED beam. With a thus sharpened illumination spot, the probes are scanned (Fig. 4B). The concept of STED microscopy was proposed in 1994 by Hell and Wichmann [109], and in 2000, experimentally realized [110]. The routinely achieved lateral resolution is 30–80 nm. Three-dimensional STED microscopy has been achieved by using a z-depletion pattern generated by two opposing objectives [111] and by combining STED microscopy with two-photon [112, 113]. STED can be used for two-color [114], and live cell imaging [115] which may be combined with FCS [116]. Because the fluorophore should not absorb the red STED light to prevent excitation and photobleaching, the choice of the proper fluorophore is important.

Single-molecule-based super-resolution microscopy has been realized using photoactivatable fluorescent proteins in photoactivated localization microscopy (PALM [117]) and fluorescence photoactivation localization microscopy (FPALM [118]) and similarly exploiting the on-off blinking behavior of organic dyes as in stochastic optical reconstruction microscopy (STORM [119]) and direct stochastic optical reconstruction microscopy (dSTORM [120]). In these techniques, only a few sparsely distributed fluorophores are stochastically photoactivated to fluoresce while neighboring molecules remain dark (Fig. 4C). The individual fluorophores are imaged, localized, and bleached. The emitted light from a single fluorophore of about 2.5 nm in size as in the case of PA-GFP will be blurred and spread by the imaging optics due to diffraction (see above). The resulting intensity profile can be described with the so-called point spread function. Such a blurred spot is about 250–300 nm in lateral and 500–800 nm in axial dimension. This prevents details in the spot to be resolved. However, the prior knowledge that just a single molecule is being viewed allows the mathematical localization of the xy intensity center of the spot with a much greater precision. For localization, Gaussian fitting has been used as in previous single-particle tracking methods [121]. Determinants of the localization precision are the number of detected photons per fluorophore, pixel size of a pixilated detector and background noise [122] and can routinely reach 10–20 nm. Localization precision is often reported as the standard deviation of localization measurements or as full-width at half maximum (FWHM), which gives the width of a distribution at half of its maximum and equals 2.35× the standard deviation for a Gaussian distribution. FWHM illustrates the closest distance between objects to be resolved (see Rayleigh and Sparrow). Then, in the next cycle, adjacent fluorophores can be switched on. Merging all obtained single-molecule localizations yields the final pointillistic super-resolution image. Thus, repeated activation and sampling permit densely expressed fluorescent proteins or densely labeling with organic dyes to be resolved in time, even though they are spatially inseparable when viewed all together at once. Beside the localization precision of the single fluorophore, the amount of thus detected molecules per area, i.e., the labeling density, specifies the resolution of the final image. The average distance between two molecules should be half of the desired resolution (known as Nyquist criterion) [123, 124]. For confocal laser scanning approaches, the Nyquist criterion correspondingly requires a sampling interval twice the desired resolution.

First realizations of PALM and STORM used TIRF illumination to reduce background. To increase the axial resolution beyond the 150–200 nm of the TIRF zone, a weak cylindrical lens has been introduced into the detection path to create differential ellipticity in xy depending upon a point source emitter's z-position [125]. In a different approach, interferometry has been applied to achieve the best z-resolution in single-molecule-based imaging so far of about 10 nm [126]. While both of these techniques use TIRF or illumination at a highly oblique incident angle, biplane FPALM [127] has been demonstrated for wide-field fluorescence illumination. Here, a 50:50 beam splitter creates two emission paths of different length and thereby two image planes about 350 nm apart in z direction. Alternative approaches include engineering of the point spread function to a double helix [128] or obtaining an axial view of the point spread function [129] to derive information about the axial position of a fluorophore. Whole-cell PALM has been realized using two-photon illumination for PA [130, 131]. To analyze further the acquired single-molecule positions in single-molecule based images, various tools of point-pattern statistics as used in forestry have been applied. These include cluster analysis with Ripley's k-function [132] and pair correlation [133]. Recently, a method using calibration probes of directly coupled spectrally distinct photoactivatable proteins has been introduced that permits quantification of the relative amount of two proteins in a given membrane area and within a protein cluster [134]. Single-molecule-based imaging is applicable to live cells. Examples include PALM imaging of entire structures [135] and STORM of labeled ligands or proteins labeled with various small tags [136-138]. Live cell PALM has also been used to gain insight into the diffusional behavior of single molecules at the plasma membrane, with [139] and without [132] the identification of individual tracks.

Conclusions

Ever since its first realizations, the development of fluorescence microscopy has been motivated by aspirations to illuminate and understand the functional anatomy of the cell. Thereby, fluorescence microscopy has permitted fundamental insight into the steady-state organization of a cell and its dynamics as well as the structure, turnover, mobility, and function of its components. “Just as GFP and other fluorescent molecules absorb light of one wavelength and convert it to light of a different wavelength, we, too, take in what others have learned about the world, add our observations and insights, and produce additional gains in human knowledge” (Martin Chalfie, Nobel Lecture, December 8, 2008).

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2011

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Ancillary