Technologies

The ‘classic’ fluorescence microscopy: illumination and detection occur via the same objective, the whole illuminated field of view is imaged onto a camera (typically sCMOS). Out-of-focus ‘blur’ from above and below the focal plane is not removed (no optical sectioning). However, this can be done subsequently by computational deconvolution. Overall, widefield microscopy stands out by speed, sensitivity and simplicity, making it an optimal choice for live imaging and high-content screening of relatively thin samples (e.g. mammalian cells in culture, thin tissue sections).

The workhorse for cell and developmental biology thanks to its optical sectioning capability (confocality) and versatility allowing for imaging of live and fixed samples labeled in multiple colors at an excellent XYZ resolution (down to ~140 nm XY and 800 nm in Z). The technology is based on a focused excitation laser beam that scans the sample, resulting in emitted photons that are being detected by point detectors. Optical sectioning is achieved by removing out of focus light using a pinhole. Since the sample is scanned point-by-point, this technology tends to be slow and especially limiting for large samples. However, signal-permitting, fast “resonant” scanners are available that enable rapid confocal recording of fixed or live samples with lower phototoxicity associated with the latter. Additionally, a confocal microscope may be used to acquire sample information from laser light that it reflects, thus circumventing the label requirement. Likewise, interference reflection signal may be recorded to study cell-surface adhesion in fixed or live samples.

The spinning disk confocal combines the speed of a widefield illumination with optical sectioning of a confocal microscope. Illumination and detection light passes through a fast-rotating disc with many pinholes, effectively parallelizing the single pinhole of an LSM. In combination with recent high quantum efficiency sCMOS cameras, this method allows exceptionally fast imaging of specimens and lower phototoxicity during live imaging. However, it is slightly less flexible and versatile than a confocal LSM. The speed of spinning disk systems gives a particular advantage over LSMs when looking at very fast processes in live cells, or when scanning large fields or large volumes of fixed samples, entire slides, for example. The combination with other techniques, such as fluorescence recovery after photobleaching (FRAP) is possible.

In TIRF microscopy, the sample is illuminated at the critical angle, leading to fluorophore excitation within the evanescent field, which extends only up to a few nm above the coverglass (~ 100 nm). This constellation results in an inherent, extreme optical sectioning, as well as a very good signal to noise ratio as nearly no fluorophores are excited away from the evanescent field. Illumination just below the critical angle leads to a highly inclined and laminated optical sheet (HiLO) which excites a thin, inclined plane within cells,  thicker than TIRF, but allowing imaging slightly deeper into the sample. The detection using a camera makes TIRF and HiLO a very fast and sensitive method used for selective plane illumination microscopy (SMLM) and for single molecule tracking.

Multiphoton microscopy is an LSM utilizing a high-power, pulsed, near-infrared laser. The most frequently used application of this method – two-photon microscopy – relies on the simultaneous absorption of two photons by fluorescent dyes. The likelihood of this event taking place decreases rapidly with an increase in distance from the focal point. When applied in LSM, fluorescent dyes are exited in and light is emitted from a well-defined optical section. Moreover, imaging of biological tissues up to 1 mm is possible thanks to reduced scattering by the utilized infrared excitation light. Therefore, two-photon LSM is the method of choice when acquiring optical sections from turbid, scattering, thick samples and is often used to acquire thick tissue sections, organoids, spheroids or utilized during intravital imaging. However, depending on the sample, the energy of two excitation photons may be combined to create a photon with twice the energy in a process referred to as second-harmonic generation (SHG). Collagen may be visualized via SHG in a label-free fashion. A further case involves the interaction of three photons within the focal point, which is utilized in third harmonic generation (THG) microscopy. It may be used to visualize myelin label-free.

The sample is illuminated with a thin light sheet typically generated by additional low NA illumination objective(s) positioned perpendicular to the detection objective, which projects the emitted light onto a camera. As opposed to an LSM or a spinning disk confocal, in the SPIM geometry only the region from which fluorescence is detected is illuminated, leading to a very low photo-toxicity and bleaching. In combination with optical clearing and an sCMOS camera, SPIM is the technique of choice for fast and gentle volumetric imaging of large specimens or whole organs.

Following excitation, a fluorophore emits photons within a specific time, the so-called fluorescence lifetime (in the range of nanoseconds for typical fluorophores). In fluorescence lifetime imaging (typically on an LSM microscope), photon arrival times after multiple excitation pulses are recorded at each pixel for the entire field of view. The lifetime may be highly specific and constant for some fluorophores, while others’ lifetime may correlate to variations in their molecular environment. Fluorescence lifetime can be used to separate the signal of fluorophores that are otherwise spectrally indistinguishable, to perform quantitative FRET (fluorescence resonance energy transfer) experiments for the study of molecular interaction and structure and to collect signal independent of background-scattering and autofluorescence. There is an ever-growing list of FRET/FLIM biosensors may be utilized toward the study of cellular signaling networks (e.g. https://doi.org/10.1021/acs.chemrev.8b00333).

Super-resolution denotes all methods capable of a resolution beyond the diffraction limit of light as defined by Abbe’s law law. Super-resolution can be achieved by shaping the illumination pattern (e.g. STED), temporally separating single emitting fluorophores (SMLM) and re-assignment of pixel intensities based on the diffraction pattern (e.g. AiryScan).
 

The position of a single emitting fluorophore can be determined with a precision that goes far beyond the resolution limit. However, in a typical sample nearly all fluorophores emit simultaneously making it impossible to determine the position of a single emitter. SMLM uses the fact that some fluorophores ‘blink’, i.e. can emit in a stochastic way (depending on the mechanism referred to either as PALM, STORM, GSD or PAINT). This temporarily separates emission events. Thus, the recording of many blinking events from the respective fluorophores over time and determining their position with an ever-evolving accuracy allows for the reconstruction of a super-resolved image. This technique requires excellent staining, thin samples, and several post-processing steps resulting in a resolution of ~ 10 – 20 nm in the XY plane.

STED is an LSM super-resolution method that uses a second, so-called depletion beam that concentrically surrounds the excitation beam in a doughnut shape. The depletion beam ‘switches off’ fluorophores effectively reducing the region of emission to the center of the excitation beam. Thereby, STED provides a super-resolved image with a typical resolution > 50 nm in XY and > 80 nm in Z direction (the latter requires 3D STED). Compared to SMLM, STED is quick and rather easy to use, and is compatible with live as well as relatively thick samples.

The mass photometer is an instrument that has become commercially available only recently. It uses interference reflection microscopy and interferometric scattering microscopy to measure the scatter of single molecules. Since the scatter signal is directly proportional to molecular mass, this technique is capable of measuring the mass of single molecules and macromolecular complexes in solution in their native state and without the requirement of any label. The technique is easy to use, fast and requires only a minimal amount of material (ca. 20 μl at 5 – 10 nM).

In a flow cytometer, cells are carried by a liquid stream (sheath fluid) and pass in a single file through laser excitation beam(s). Thereby, the system is able to very quickly measure the fluorescence intensity and scattering properties of thousands of cells. In the cell sorter, the detected signals are used to separate cell populations. The cell sorter generates liquid droplets containing single cells that get electrically charged and are then deflected to a specific container according to the measured fluorescence or scatter parameters. FACS is used to characterize cell populations, enrich populations of a specific type, and obtain single cell clones, e.g. for the generation of transgenic endogenous cell lines.

A plate reader can be considered as a single pixel microscope, where a measurement is performed per well of a multi-well plate. It is ideal to rapidly screen a plate and measure absorption, fluorescence, fluorescence anisotropy and other parameters.