Frequently Asked Questions

Other commonly used super-resolution techniques with epi-fluorescent microscopes are Stimulated Emission Depletion (STED) and Structured Illumination Microscopy (SIM). Lattice light-sheet microscopes achieve SIM-level resolution. The resolution of STED is 30 nm, and SIM usually cites a resolution of around 100 nm. MINFLUX is a super-resolution microscopy technique combining STED and single-molecule localization. It allows you to resolve structures as small as a molecule along all three dimensions but is very slow.

SMLM is a super-resolution fluorescence microscopy technique that breaks the optical diffraction limit of standard light microscopy. The diffraction limit is between 200 nm and 350 nm.

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The typical resolution cited for SMLM is 20 nm. However, SMLM can achieve higher resolutions of about 15 nm with SMLM methods such as DNA-PAINT.

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Single Molecule Localization Microscopy (SMLM) and other super-resolution techniques differ in several ways. For example, STED and SIM both use manipulation of excitation illumination. STED uses a second laser to deactivate fluorophores and reduce the illumination area in the specimen. It is a laser scanning method like a laser scanning confocal. SIM collects a series of images using patterned illumination to generate an image computationally.

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The principle of Single Molecule Localization Microscopy (SMLM) is to localize individual fluorescent molecules with high precision and reconstruct a super-resolution image from their coordinates. SMLM techniques control the switching of fluorophores between a bright (on) state and a dark (off) state, such that only a sparse subset of molecules emit light at any given time, and their point-spread functions (PSF) do not overlap. This approach allows the separation of overlapping signals from different fluorophores and the fitting of their point spread functions to determine their positions. Repeating this process over many frames allows the fitting algorithm to localize many molecules and use their coordinates to generate a high-resolution image.

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Photoactivated Light Microscopy (PALM) uses photoswitchable dyes or fluorescence proteins to achieve the on-off switching that enables all SMLM techniques. E.g., a fluorescent protein exists in a dark state and converts to a fluorescent state after exposure to 405 nm light. Now it emits photons when excited with the appropriate wavelength. The microscope collects the photons and uses them to localize the protein and the attached structure. The power of the 405 nm conversion laser is so low that only so few fluorophores are switched on that their point-spread functions (PSF) do not overlap. After localization, the fluorophores are bleached (switched off), and a new subset is activated. This process is repeated until the structures of interest are reconstructed. Often fluorescence proteins that change their emission wavelength after activation are used as an alternative.

PALM probes are typically fluorescent proteins. Examples of photoactivate probes used for PALM are paGFP, pamCherry, paTagRFP. Examples of photoconvertible probes used for PALM are mEos, Dendra, and mMaple.

PALM probes are commonly used in live cell experiments because they are endogenous. However, the length of an experiment is limited because the probes blink and then become photobleached. Since each fluorophore only blinks once, PALM probes are suitable for experiments that quantify the exact number of molecules.

Stochastic Optical Reconstruction Microscopy (STORM) and direct Stochastic Optical Reconstruction Microscopy (dSTORM) are two super-resolution microscopy techniques that rely on organic probes that photo-switch from a dark state to an off state and back to a dark state. STORM and dSTORM require buffers with a thiol for a reduction and typically an oxygen scavenger. STORM uses an activator probe in addition to the imaging probe, while dSTORM does not. A high laser power drives the imaging molecules into a dark state, from which they can emerge periodically to the singlet state and fluoresce before returning to the dark state, thus producing a blink. The duration of individual blinks is typically 5-20 ms. A fluorophore molecule used for STORM or dSTORM can go from on to off state several times before the molecule photo-bleaches.

When choosing probes for dSTORM, their photophysical properties, such as their blinking ability, must be considered. Alexa 647 is an excellent photo-switching probe that can cycle up to 16 times before photobleaching and activates well. Cy3B is a good second probe for two-color experiments with AF 647. For three-color experiments, AF 488 is a viable choice for labeling the most densely labeled structure being imaged.

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PAINT stands for Points Accumulation for Imaging in Nanoscale Topography. It is an imaging technique that relies on the temporary binding of a ligand with a fluorophore to produce a localization event or blink. When the PAINT ligand is not bound to the substrate, it moves around in the sample solution and appears as fluorescent background. However, when it attaches to the substrate, the ligand with the fluorophore becomes immobilized and can be localized.

One of the most common forms of PAINT is DNA-PAINT. In this technique, a small oligonucleotide with a fluorophore attached (called an imaging strand) is complementary to another oligonucleotide (called a docking strand) attached to the target being imaged. The imaging strands bind temporarily to the docking strands and are localized. Binding times range from 100 to 300 milliseconds and depend on the oligonucleotide length. Docking strands can be attached to antibodies, nanobodies, directly to a target protein, or part of a larger oligonucleotide sequence attached to DNA or RNA.

Using multiplexing with fluidics, DNA_PAINT can label more targets than can be separated chromatically with multiple fluorophores. Additionally, the localization precision of DNA-PAINT can be better than that seen with STORM or PALM because DNA-PAINT has higher photon counts.

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Single-Molecule Localization Microscopy (SMLM) can use standard fluorescent labeling methods. Immunofluorescence labeling methods are currently the most common labeling method for SMLM. Another option is to use fluorophores conjugated to nanobodies to label targets. This method has the advantage of having a smaller linkage error, which refers to the distance between the fluorophore and the labeled target molecule.

Genetically fused self-labeling protein tags, such as SNAP and Halo tags, can be used for Single-Molecule Localization Microscopy (SMLM) and offer the advantage of reduced linkage errors. Photoactivated Localization Microscopy (PALM) typically uses genetically encoded fluorescent proteins. However, for SMLM, a photoactivatable or photoconvertible fluorescent protein is required. Standard Green Fluorescent Protein (GFP) is unsuitable for SMLM but can be labeled with nanobodies.

It is important to note that for SMLM studies, samples must be mounted on coverslips rather than on covered slides because samples must be exposed to buffers during image acquisition.

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Single-Molecule Localization Microscopy (SMLM) data analysis fundamentally differs from other fluorescence microscopy techniques because it is based on localized, single-dye molecules rather than pixels representing the summed intensity of multiple dye molecules. SMLM data analysis uses the coordinates of each localization. The precision of a localization is 10-20 nm compared to the diffraction limit of a standard microscope of 200 – 300 nm. In SMLM, the analytical algorithms used for data analysis are statistical methods to analyze the density and distribution of molecules, clusters, and colocalization at the molecular level.

In contrast, other fluorescence microscopy techniques, such as wide-field, confocal, STED, and SIM, are pixel-based and rely on standard image analysis algorithms. While STED and SIM can provide a resolution of 30-100 nm in measuring the distance between objects, they cannot directly measure the density of objects, such as clusters or colocalization at the molecular level, like SMLM can.

Acquiring and analyzing 3D data with Single-Molecule Localization Microscopy (SMLM) is possible. While early implementations of SMLM were only in 2D, and some researchers still use 2D SMLM on home-built systems, currently available commercial systems provide 3D localization for SMLM. Most systems use astigmatism, a point-spread function (PSF) engineering method, to localize particles in 3D.

The Vutara VXL from Bruker is a unique Single-Molecule Localization Microscopy (SMLM) system that uses a biplane method to acquire 3D data. In this method, the emission is split into two paths with a 600 nm difference in path length. By calibrating the system using experimental point spread functions (PSFs), the biplane method can collect localizations over a one-um range for a single sample plane. It can also collect 3D data over a larger range by performing Z series. This approach allows for deeper imaging, imaging adherent cells and tissue slices of 50 um and thicker, and even model organisms.

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Single-Molecule Localization Microscopy (SMLM) can be used to analyze various samples. For example, the Vutara VXL from Bruker allows for imaging adherent cells, tissue slices 50 um and thicker, and even model organisms such as C. elegans and Drosophila. Samples can be fixed, or, with appropriate labeling methods, live cell imaging can be performed.

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Electron microscopy (EM) can provide a resolution of up to 1 nm or better, about 20 times better than the resolution of Single-Molecule Localization Microscopy (SMLM). However, SMLM allows the labeling, imaging, and identification of specific targets.

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