Proteomics/Protein - Protein Interactions/Experimental
Page Edited and Updated by: Dan Surdyk
Determining Binding Experimentally - In Vivo
Fluorescence Resonance Energy Transfer uses fluorescent energy transfer to visualize protein interactions. The procedure is simple. Fluorophores are fused to the proteins of interest and then bombarded with light at the excitation wavelength. The first fluorophore transfers some of the energy it absorbed from the light source to the second fluorohore, which in turn emits some of its energy into the environment where, with the use of a fluorescent microscope, it is visible.
Bioluminescence Resonance Energy Transfer is a biodetection system that utilizes energy transfer between a donor and receptor to visualize binding. When the two molecules are in close proximity, a green fluorescent protein on the receptor absorbs the energy from luciferase attached to the donor. As a result, the GFP emits light that appears green to the naked eye.
The yeast two-hybrid system is an in vivo system that utilizes the binding domain (BD) and activation domain (AD) of genes (used in transcription) to determine protein interactions. Proteins are fused to both the BD and the AD of a reporter gene, one to each. When the protein attached to BD interacts with the protein fused to the AD, transcription is initiated, and the reporter gene is transcribed. The effect is quantifiable by measuring the levels of reporter gene expressed.
Determining Binding Experimentally - In Vitro
Thanks to the success of DNA microarray technology, other areas of science are developing arrays or “chips” to test for multiple things at once. One of these areas is protein interaction studies, where array technology is currently expensive and complicated, but looks like it will soon be a realistic option. Protein interaction arrays have the advantage of allowing a scientist to test for numerous interactions quickly and conveniently. Read more about arrays in Chapter 8: Protein Chips.
To perform TAP, a “TAP tag” must be attached to the protein of interest. This is done by genetic recombination of the protein with the tag, usually using plasmids. The expressed fusion protein then contains the protein of interest covalently bound to the tag. The tag consists of two IgG binding domains from Protein A from Staphylococcus aureus and a calmodulin binding protein (CBP), with a TEV protease cleavage site between those two sections. The CBP is closest to the protein, and the IgG binding domains are farthest away. When the protein is extracted from the cells, any interacting proteins will come with it, giving you complexes of protein attached to the tag rather than just your one protein. The sample is applied to a column of IgG beads, so that the complexes of interest bind to them due to protein A, and any other proteins pass through. The bound proteins are released by cleavage with the TEV protease, and the resulting proteins passed through a column with calmodulin beads. The CBP causes them to bind these beads, and the protease and any other contaminants pass through. This results in a very pure sample of the protein of interest and its interaction partners. The resulting proteins can be identified by SDS-PAGE or mass spectrometry.
Unlike many interaction detection methods, solid-phase detection does not involve labeling of proteins. In these techniques, one protein is attached to the surface of a transducer. The other protein is contained in a solution that can be applied to the surface. This allows binding kinetics of a reaction to be studied in real time. Some of the common approaches use surface plasmon resonance or reflectance interference to detect and image the interactions. Just like the protein array trend, this area is moving more toward parallel analysis of several interactions. This can be done with a microfludic system, where proteins in solution are passed through a tiny channel that runs over a series of surfaces that each have a different type of protein attached to them.
Some of the newest and most exciting developments in studying protein interactions involve single molecules. These allow experiments to be carried out with low concentrations, and with well-defined samples so there won’t be other interactions confounding the results.
One method used is force spectroscopy. One member of a pair of interacting proteins is attached to the tip of an atomic force microscope, which can measure very small forces. The other protein is attached to a solid surface. The tip is brought down to the surface so that the proteins interact, and then pulled away. The amount of force needed to pull the tip away from the surface is the force of the interaction. This method was used effectively by Nevo et al. to determine whether the binding of an effector protein to a receptor caused a conformational change.
In addition, fluorescence can be used to detect single interacting pairs of proteins. Two proteins can be marked with different fluorescent labels, and confocal microscopy used to scan a very thin section of a small volume of sample and detect the fluorescence. When it detects both colors, it knows that it must have found an interacting pair, because they have to be very close to be in the same thin section.
- FRET: http://biowww.net/browse-44.html
- BRET (the BMG Lab site): http://www.bmglabtech.com/
- Yeast two-hybrid: http://www.uib.no/aasland/two-hybrid.html
References (Open Accss)
 Nevo, R., Brumfeld, V., Elbaum, M., Hinterdorfer, P., Reich Z. 2004. Direct discrimination between models of protein activation by single-molecule force measurements. Biophys J, 87:2630-2634.
 Piehler, J. 2005. New methodologies for measuring protein interactions in vivo and in vitro. Current Opinions in Structural Biology, 15:4-14.
 Puig, O., Caspary, F., Rigaut, G., Rutz, B., Bouveret, E., Bragado-Nilsson, E., Wilm, M., Se´raphin, B. 2001. The Tandem Affinity Purification (TAP) Method: A General Procedure of Protein Complex Purification. Methods, 24:218-229.