The noted German physicist and instrument designer Ernst Abbe realized in 1873 that the resolution of optical imaging in microscopy was fundamentally limited by the diffraction of light as spherical wavefronts pass through the circular apertures of objectives and conjugate focal planes. Abbe's research had revealed that the ultimate resolution of a microscope was not constrained by the quality of the lenses; rather it was restricted by the wavelength of light passing through the instrument as well as the aperture angle of the objective. In formalizing the relationship, Abbe noted that a microscope could not resolve two objects positioned closer than λ/2NA, where λ is the imaging wavelength and NA (numerical aperture) is defined by the objective aperture angle and refractive index of the imaging medium. This hallmark equation defines the lateral resolution in the x-y plane, which is perpendicular to the optical axis of the microscope. Resolution in the axial dimension is at least twice as poor. The diffraction barrier hindered the performance of optical microscopes for several centuries and was considered a physical limitation that could not be overcome with the use of standard glass-based objectives. In recent years, however, several new approaches have emerged that circumvent the limits of diffraction in optical microscopes based on the reversible photoswitching of fluorescent probes to achieve what is now commonly referred to as superresolution imaging. Thus, a number of exciting and creative new techniques have been advanced that apparently have no fundamental limit in achieving high spatial resolution, therefore rendering it possible to resolve specimen features in terms of just a few nanometers.

The new superresolution techniques can be roughly divided into two categories: those that are able to image ensembles of fluorophores using spatially modulated, focused light with saturating intensities, and those that sequentially image single molecule emitters that are spread to distances greater than the Abbe diffraction limit. The latter methods include photoactivated localization microscopy (PALM), stochastic optical reconstruction microscopy (STORM), and fluorescence photoactivation localization microscopy (FPALM), technologies that are differentiated primarily by the character of probes used for labeling the specimen. However, regardless of the minor technical differences between PALM, STORM, and FPALM, these single-molecule localization techniques all rely on the common principle of stochastically activating, localizing, and then photobleaching synthetic or genetically-encoded photoswitchable fluorophores. PALM and FPALM originally used photoactivatable or photoconvertible fluorescent proteins targeted to sub-cellular structures at high density to achieve the localization precision necessary to generate images at more than 10 times the spatial resolution afforded by diffraction-limited fluorescence microscopy. In contrast, STORM was developed using synthetic photoswitchable carbocyanine dyes to produce similar resolution.
The principle of single-molecule localization microscopy is illustrated in Figure 1 with a series of cartoons detailing the sequence of events for determining the precise location of a single set of photoactivated fluorescent probes. Initially all molecules in the specimen are inactive (native non-emissive state; dark circles) as shown in Figure 1(a). A violet laser is used to photoactivate a sparse subset of molecules, while a green laser is used for readout of the resulting fluorescent molecules. In Figure 1(b), a 405-nanometer laser is used to photoactivate a small number of molecules in the specimen (those surrounded by boxes). This number is maintained at a very low level by ensuring the laser intensity is sufficiently weak at the focal plane. Photoactivation of the molecules occurs stochastically where the probability of activation is proportional to location and the intensity of the activation laser. After photoactivation, the 561-nanometer readout laser is used to detect and record the position of the photoactivated molecules within the illuminated area (Figure 1(c)). Digital images of the photoactivated molecules are then analyzed to identify and localize molecules (Figure 1(d)) for as long as they remain fluorescent. During readout, the photoactivated molecules spontaneously photobleach (Figure 1(e)), eventually reducing the number of active molecules in the specimen. A new set of molecules is photoactivated (Figure 1(f)) to repeat the sequence, which is reiterated until all molecules in the specimen have been exhausted.
A number of practical considerations are mission-critical in achieving the best single-molecule localization images using PALM and related methodology. The primary concern is choice of photoactivatable probe, which will govern the maximum achievable resolution. This topic will be addressed in detail in the following sections. Second, although PALM instrumentation is relatively straightforward, some additional knowledge in the basic requirements for single-molecule imaging and other aspects of PALM are necessary to achieve successful results. Among the other important variables to understand for PALM imaging are the basic detector characteristics (pixel size, noise levels, intermediate magnification, readout speed), ensuring that a sufficient number of pixels are used to image single molecules, and maintaining the density of labeled features in the specimen at a level high enough to reconstruct the features of interest. Finally, careful attention must be paid to the details of specimen preparation to reduce autofluorescence and minimize potential artifacts induced by aberrations resulting from inhomogeneities and immersion media.
Strategic Overview of PALM Imaging
PALM (as well as STORM and FPALM) is a single-molecule widefield technique where the raw data consist of an image stack typically containing thousands of individual frames, each featuring a subset of diffraction-limited fluorescence images of single photoswitchable molecules present in the specimen. The images of single molecules, appearing as bright point sources of varying intensity approximately 200 to 250 nanometers in width, are analyzed to determine their centers with high precision and the resulting information is employed to generate a high-resolution PALM image of the individual molecular coordinates. The resolution in PALM is thus dependent on localization precision, or how accurately the position of each single molecule can be determined from its diffraction-limited image. Perhaps more subtle, but equally important in determining the ultimate PALM resolution, is the molecular density of fluorescent probes in the specimen and how well they represent the target structure being analyzed.
The localization precision depends on achieving the maximum signal-to-noise ratio in each image, a characteristic that depends on maximizing the number of photons collected from each photoswitchable fluorescent probe while simultaneously minimizing the background fluorescence. Localization precision also depends on the total number of photons emitted by each molecule before it photobleaches, which is an intrinsic property of any particular molecule. In general, fluorescent proteins generate far fewer photons than do synthetic dyes, such as the carbocyanines (Cy3 and Cy5) or the ATTO dyes. In addition, the ability to take full advantage of the molecular density in each specimen is contingent on the contrast ratio of the probe, another intrinsic property that gauges the difference in fluorescence intensity between the on and off (or native and photoconverted) states of photoswitchable molecules. In many cases, the contrast ratio suffers from the fact that molecules supposedly in the "off" state often emit weak signals and contribute to the background noise. Therefore, the single most important factor to consider in designing an experiment using PALM imaging is the choice of fluorescent probe.
Although PALM and STORM were originally implemented using total internal reflection fluorescence microscopy, FPALM was originally reported using a widefield microscope that does not limit the focal plane to the proximity of the coverslip. However, all three techniques are compatible with both imaging modes and by applying typical widefield geometry, three-dimensional specimens can be imaged within a single focal plane of thickness approximately equal to the depth of field. This approach will work for sectioned tissues, bacteria, yeast, and other thin specimens, but suffers from significant background noise when attempting to image relatively thick tissue specimens using these single-molecule techniques where three-dimensional superresolution approaches are perhaps the best choice. PALM, STORM, and FPALM can be employed to gather a significant amount of single-molecule information about a specimen, including the number of photons emitted, intermittency, brightness, and the absolute number of fluorescent molecules. Additionally, the techniques can in principle determine emission spectra and fluorescence anisotropies of localized molecules.

In stark contrast to normal fluorescence imaging where specimens are initially very brightly fluorescent upon illumination, the single-molecule superresolution techniques are designed with probes that are in a dark or native emissive state (that is not imaged) before photoactivation. Therefore, virtually no fluorescence is emitted from a majority of the probes in PALM or STORM prior to being photoactivated or photoconverted by the activation laser. Once activated, the fluorescent probes are imaged with a longer wavelength readout laser (see Figure 1) matching their excitation properties. By carefully regulating the activation laser intensity, the rate at which molecules are activated can be controlled. One of the keys to PALM, STORM, and FPALM is the ability to turn off or disable the active molecules to avoid the number of emitters from growing so high that individual molecules are no longer distinguishable from one another. Removing active molecules from the field can be implemented through photobleaching or photoswitching to provide the critical balancing factor that limits the total number of visible molecules. In either case, this mechanism to control the number of active emitters is necessary to ensure that the distance between each active molecule and its nearest neighbor exceeds the lateral diffraction limit of 200 to 250 nanometers.
Once the density of fluorescent probes in a specimen is controlled to the point of allowing single-molecule imaging and localization, the only remaining requirement is a highly sensitive camera to capture images of the bright single emitters as they are activated and photobleached (or photoswitched to the off state). Photobleaching typically occurs spontaneously in the presence of the readout laser, but in cases where photoswitching is necessary, a second deactivation laser may be necessary. By creating a time-lapse video containing repeated cycles of photoactivation, readout, and photobleaching, the positions of a large number of molecules (ranging from several thousand to millions) can be determined. The uncertainty in the position of each molecule can also be determined by repetitive imaging of that molecule after it has been activated and before photobleaching. The resulting PALM image contains the measured positions of all the localized molecules displayed together.
PALM images can be rendered either by plotting the unweighted positions of localized molecules or by using weighted plots of the positions of the localized molecules rendered as spots having Gaussian profiles where the intensity is proportional to the number of photons detected (from each molecule) and the radius is equal to the calculated or experimentally determined localization-based resolution. Because the weighted plots take into account the intensity and positional uncertainty of each molecule, the resulting images appear to the observer as a more realistic representation of a fluorescence image with high resolution (see Figure 2). Typically, all molecules localized within a specific area are rendered simultaneously, but in live-cell imaging or similar time-lapse sequences, time-dependent images can be created using subsets of molecules localized during various time periods. A threshold that includes only molecules featuring a particular range of intensities, or above a minimum intensity, can also be applied to increase the localization precision. In contrast, plotting only the unweighted positions of localized molecules leads to a plot of markers on a static background and appears more like a blueprint.
Illustrated in Figure 2 are summed widefield total internal reflection microscopy (TIRFM) and PALM single-molecule images of optical highlighter fluorescent proteins fused to targeting peptides or proteins that were localized within fixed mammalian cells. The filamentous actin cytoskeletal network is presented in Figures 2(a) and 2(b) using a fusion of tandem dimer Eos fluorescent protein (a green-to-red photoconverter) with human beta-actin and expressed in Gray Fox lung fibroblast cells. Note the individual fibers that are clearly resolved in Figure 2(b). In Figures 2(c) and 2(d), a single mitochondrion (outlined with the white box in Figure 2(b)) labeled with dimeric Eos fused to the mitochondrial targeting sequence from subunit VIII of human cytochrome C oxidase is shown in widefield (Figure 2(c)) and PALM (Figure 2(d)). Finally, in Figures 2(e) and 2(f), the intermediate filament network in HeLa cells is highlighted with human cytokeratin fused to mEos2, a monomeric version of Eos fluorescent protein. The boxed area in Figure 2(e) is shown in PALM to demonstrate molecular locations in Figure 2(f). In all cases, photoconversion was conducted with a 405-nanometer diode laser and readout occurred using a 561-nanometer diode-pumped solid state laser.