FRAP is a method to measure the mobility of molecules in living specimen; in addition FRAP can be used to measure the connectivity of compartments and binding properties. In a nutshell the equilibrium of fluorescent molecules is perturbed by bleaching a region of interest with a strong pulse of light. Subsequently the recovery of fluorescence in the bleached region is monitored over time. FRAP can be performed at any current commercially available point-scanning confocal microscope; wide-field or confocal spinning disk systems can be upgraded with special in-coupling of a bleaching laser for example with scanner to perform FRAP experiments.
FRET is a phenomenon of non-radiative energy transfer, which can occur between two fluorophores (donor and acceptor) in very close proximity, usually less than 10 nm. As this distance is in the scale of biological molecules it is often exploited to probe for molecular interactions or in optical sensors (with donor and acceptor on one molecule) measuring physiological values in the cell.
Preconditions for FRET:
A set of images of donor and acceptor channels is acquired before and after bleaching a region of interest in the acceptor channel. Donor molecules which interacted with acceptor molecules before bleaching, cannot transfer energy after acceptor photobleaching and emit more fluorescence instead leading to a brighter donor image in the region of bleached acceptor. Usually used with fixed samples, since differences between pre- and post-bleach images due to movement lead to artifacts.
Donor molecules, which can return to ground state by transferring energy onto the acceptor, bleach slower (less chances of excited states to chemically react). This can be exploited to qualitatively detect FRET. Unfortunately measurements are often influenced by acceptor bleaching, this is why we do not recommend this method.
The emission of acceptor molecules excited by energy transfer from donor molecules is called sensitized emission. For the majority of FRET pairs this sensitized emission signal is overlaid by cross-talk of donor emission and cross-excitation of acceptor molecules. These contributions can be determined by donor-only and acceptor-only samples using the same imaging conditions. Three channel datasets are needed for the measurement of sensitized emission: donor excitation + emission (‘donor’,ch1), donor excitation + acceptor emission (‘FRET’,ch2) and acceptor excitation + emission (‘Acceptor’, ch3).
FRET readout of optical probes containing both donor and acceptor on a single molecule (e.g. cameleons) can be measured by ratiometric imaging. With a fixed ratio between donor and acceptor the fluorescence emission, the FRET changes can be measured by the ratio between donor channel and ‘FRET’ channel (donor excitation + acceptor emission).
FCS is a method to measure diffusion dynamics and interaction of fluorescent particles in liquid environment. It is based on the measurement of autocorrelation of the fluorescence signal from molecules moving through the focal volume of confocal microscope. The autocorrelation curve calculated by the software gives information on apparent diffusion and concentration of molecules. If the size of the focal volume is calibrated, the diffusion coefficient of the investigated molecules can be accurately calculated. If two types of particles are marked with different fluorophores, one can measure cross-correlation between the signals (fluorescence cross-correlation spectroscopy, FCCS). If particles move independently there will be no cross-correlation, provided that there is no bleed through between fluorescence channels. If particles interact, which implies that they move together, it will create a significant cross-correlation between the signals. Thus the degree of cross-correlation can be an indication for both interaction and the percentage of interacting species. Method can be used for in vitro and in vivo measurements.
Focused intense laser light can be used to selectively alter or destroy volumes in biological specimen down to the size of the diffraction limited focus spot. Pulsed lasers are needed to achieve sufficient energy levels to form plasma localized to the focus volume. Regions of interest can be treated by moving the laser spot by a scanner, enabling different types of experiment from line cuts (e.g. cutting actin stress fibres in cells or even cutting a cell into two pieces) to ablating clusters of cells by scaning a larger region.
A number of fluorescent proteins and dyes are available, which can be activated (switched from a dark state to a fluorescent state; e.g. paGFP) or converted (e.g. from green to red fluorescence like Kaede or EOS2) by irradiation with low doses of UV/blue light. Using a scanner, it is possible to activate/ convert selected regions of interest. This can be used to mark cells or to measure molecule mobility similar to FRAP experiments.
Similar equipment can be used for photo-uncaging experiments: UV-light can cleave off so called cages from specifically synthesized molecules. The cage can either mask a bio-active side-chain of the molecule, rendering it biologically inactive, or quench fluorescence until uncaging. Alternative names of this method are photo-activation or photo-stimulation.
Availability in the ALMF:
Localization Microscopy is a super-resolution technique based on single molecule detection. The resolution can be improved approx. 10 fold compared to diffraction limited widefield microscopy resulting in a resolution of ~20nm laterally and ~50 nm axially. Localization Microscopy can be performed in TIRF and Epifluorescence and is therefore well suited for a wide range of (chemically fixed) samples. Photoactiveted Localization microscopy, direct Stochastical Optical Reconstruction Microscopy (dSTORM) and GSDIM (ground state depletion microscopy followed by individual molecular return) are all techniques based on the principle of single molecule detection. Whereas PALM is typically performed with photoactivatable or photo-convertible fluorescent proteins, dSTORM / GSDIM is using organic dyes, which are temporally switched off to achieve single molecule detection. GSDIM / dSTORM is often used in combination with immunofluorescence (IF) staining. Although many standard organic dyes (e.g. Alexa, Atto and Cy-Dyes) can be used, special buffers / embedding media are required for embedding / mounting.
STED is a scanning confocal microscopy based super-resolution technique. The resolution is improved more than 5 fold (<50 nm in x, and y) by shrinking the confocal volume using stimulated emission depletion of excited fluorophores in the outer part of the confocal spot. A wide range of organic dyes emitting from green to red (including common dyes like Alexa 488 and Alexa 568) provides multi-colour capability for super-resolution co-localization studies using fixed specimen. The compatibility with fluorescent protein like eYFP, eGFP or mStrawberry additionally allows the imaging of living cells. The system is equipped gated detection which provides enhanced resolution and better suitability for life cell imaging compared to standard CW-STED. In embedding media (nearly) matching the refractive index of immersion oil, STED-3D is available as an option reducing the resolution in x,y and z below 130 nm.
TIRF microscopy allows the selective illumination of a very thin optical slice (~70nm to ~250nm thick) at the cover-slip surface. This is achieved by shining the excitation laser at a high angle onto the coverslip-sample interface. This leads to the total reflection of the excitation laser and only a thin layer directly at the coverslip surface (the so-called “evanescence wave”) contains light to excite fluorophores. The advantage of this method is that no fluorophores in the bulk solution will be excited leading to a very high signal to background ratio. Typical applications of TIRF microscopy include: single molecule detection in in vitro experiments; studies related to the basal plasma membrane, such as endo- or exocytosis, focal adhesions as well as membrane translocation of signal molecules.