One asset of mass spectrometry in protein science is that ESI and MALDI [11, 75] can introduce noncovalent complexes to the gas phase [12, 76, 77]. If one can assume that the gas-phase ion abundances (peak intensities) for the complex, apo protein, and ligand are directly related to their equilibrium concentrations in solution, the relative and absolute binding affinities can be deduced [78-81]. Extended methods are now available that also make use of the intensity of the complex and the protein at high ligand concentration to determine binding constants [78, 82-84].
Unfortunately, ESI is discriminatory and peak intensities especially when measuring a system at equilibrium may not be reliable [85, 86]. Electrostatic forces in complexes are strengthened in the solvent-less environment of the mass spectrometer, making electrostatically bound protein-ligand complexes more stable in the gas phase than in solution. Binding that is largely governed by hydrophobic interactions in solution, however, weakens in the vacuum of a mass spectrometer, and complexes bound by hydrophobic forces break apart to an unpredictable extent, leading to incorrect affinities [76, 87, 88]. One may correct for fragmentation of a noncovalent complex in the gas phase by using response factors that relate the mass spectrometer signal to the concentration of the complex in solution and ultimately give the correct stability of the complex. A recently announced method  cleverly uses only the signal intensity of the complex and follows it in a titration, much the same way as PLIMSTEX takes only the changing mass of the protein during a titration. Modeling the changing intensity as a function of added ligand gives the response. Although use of response factors may avoid some of the problems of direct measurements, the ionization process must still bring detectable amounts of protein-ligand complex into gas phase, and this remains problematic for weakly bound systems. Furthermore, for systems having a small Ka, the titration must be performed at high concentration of ligand and protein, regions where the response of ESI may be nonlinear [90-94].
An additional problem for all direct methods is that they cannot use high ionic strength and nonvolatile buffers, which are needed to simulate physiological conditions, because ESI does not work under these conditions. Thus, nonspecific ad-ducts may be produced, confusing the stoichiometry and affinity determinations.
Furthermore, if the affinity is to be measured in water, then ESI must be done with solutions that have high contents of water, which is difficult or impossible. Another problem is that different source configurations (e.g. normal vs nano ESI), desolvation conditions, and instruments may give different results in affinity determinations .
PLIMSTEX avoids these problems by following changes in H/D exchange by using the mass shifts accompanying exchange; the signal intensities for the complex are not required. As such, it takes advantage of the increasing ability of mass spectrometers to measure accurately m/z, a measurement that is not compromised by the discrimination in measuring signal intensities by ESI. The basis for PLIMSTEX is reactivity, similar to footprinting , but there is a strong analogy to titration monitoring by spectroscopic methods (e.g. absorbance or fluorescence). SUPREX, another method for measuring the free energies of binding from H/D exchange rates during unfolding (for some examples, see [20, 97]), also takes a single parameter from the mass spectrum (i.e. the m/z) and avoids the complications of relying on ESI signal intensities.
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