The binding of one protein to another provokes a variety of biophysical changes that can then be used as a measure of the binding reaction. Optical spectroscopy, particularly fluorescence, is the most flexible technique, but surface plasmon resonance biosensors, microcalorimetry and mass spectroscopy have recently shown significant development.
Our understanding of the molecular basis of biological functions depends;
- On our ability to identify interacting partners
- Characterize how they interact in space and time
- Imaging tools have ability to gives spatial and temporal information
Fluorescence resonance energy transfer (FRET) is a common technique when observing the interactions of different proteins. Applied in vivo, FRET has been used to detect the location and interactions of genes and cellular structures including integrins and membrane proteins.
Förster (or Fluorescence) resonance energy transfer (FRET) is a physical process in which energy is transferred non radiatively from an excited fluorophore, serving as a donor, to another chromophore (acceptor). Among the techniques related to fluorescence microscopy, FRET is unique in providing signals sensitive to intra- and intermolecular distances in the 1-10 nm range. Because of its potency, FRET is increasingly used to visualize and quantify the dynamics of protein-protein interaction in living cells, with high spatio-temporal resolution. Here we describe the physical bases of FRET, detailing the principal methods applied: (1) measurement of signal intensity and (2) analysis of fluorescence lifetime (FLIM). Although several technical complications must be carefully considered, both methods can be applied fruitfully to specific fields. A different approach, based on TIRFM measurement of the FRET intensity of fluorescently labeled recombinant proteins, suggests that a direct interaction also occurs between integrins and the ether-a-go-go-related-gene 1 (hERG1) K+ channel.
Bimolecular fluorescence complementation (BiFC) is a new technique in observing the interactions of proteins. Combining with other new techniques, this method can be used to screen protein–protein interactions and their modulators, DERB.
Affinity electrophoresis is used for estimation of binding constants, as for instance in lectin affinity electrophoresis or characterization of molecules with specific features like glycan content or ligand binding.
Label transfer can be used for screening or confirmation of protein interactions and can provide information about the interface where the interaction takes place. Label transfer can also detect weak or transient interactions that are difficult to capture using other in vitro detection strategies. In a label transfer reaction, a known protein is tagged with a detectable label. The label is then passed to an interacting protein, which can then be identified by the presence of the label.
Tandem affinity purification (TAP) method allows high throughput identification of protein interactions. In contrast to yeast two-hybrid approach the accuracy of the method can be compared to those of small-scale experiments and the interactions are detected within the correct cellular environment as by co-immunoprecipitation. However, the TAP tag method requires two successive steps of protein purification and consequently it cannot readily detect transient protein–protein interactions.
SPINE (Strep Protein interaction experiment) uses a combination of reversible crosslinking with formaldehyde and an incorporation of an affinity tag to detect interaction partners in vivo.
Proximity ligation assay (PLA) in situ is an immunohistochemical method utilizing so called PLA probes for detection of proteins, protein interactions and modifications. Each PLA probe comes with a unique short DNA strand attached to it and bind either to species specific primary antibodies or consist of directly DNA-labeled primary antibodies. When the PLA probes are in close proximity, the DNA strands can interact through a subsequent addition of two other circle-forming DNA oligonucleotides. After joining the two added oligonucleotides by enzymatic ligation, they are amplified via rolling circle amplification using a polymerase. After the amplification reaction, several-hundredfold replication of the DNA circle has occurred and fluorophore or enzyme labeled complementary oligonucleotide probes highlight the product. The resulting high concentration of fluorescence or chromogenic signal in each single-molecule amplification product is easily visible as a distinct bright spot when viewed with either a fluorescence microscope or a standard bright field microscope.