Introduction

Protein-ligand interactions are the basis for numerous biological processes for which quantitative understanding is vital [1, 2]. To quantify binding, investigators have used circular dichroism [2], fluorescence and fluorescence polarization [3, 4], nuclear magnetic resonance [5], surface plasma resonance [6, 7], and isothermal titration calorimetry [8]. The macroscopic binding affinity, however, does not permit complete understanding of a protein-ligand system. The requirements for specific fluorescence labeling [3, 4], special sample preparation [5, 6], and large sample amounts [5] also limit these approaches.

Among the mass spectrometry (MS)-based approaches for quantitative protein-ligand interactions, one class, termed direct methods, utilize the spectrometer to measure concentrations at equilibrium [9,10,11,12]. Despite their convenience, there is always a question of whether the measured gas-phase concentrations represent those in solution. To avoid this, indirect methods using hydrogen/deuterium exchange (HDX) can also be used. Stability of unpurified proteins from rates of HDX (SUPREX) [13] yields a protein-ligand binding affinity by measuring the stability of proteins as bound and unbound.

Another method, protein-ligand interactions by MS, titration and HDX (PLIMSTEX) [14, 15], gives binding affinity in a titration-based experiment [16,17,18,19]. Titrating the protein at high-concentration (e.g., 100 times Kd) yields binding stoichiometry, whereas titration at ~ Kd gives the binding affinity without any modifications (e.g., tagging by a fluorophore) [14, 15]. Indirect methods also characterize binding-induced conformational changes at a regional level represented by peptides from digestion [20,21,22]; an example is apolipoprotein E3 and a small-molecule drug candidate where location and affinity were determined [19]. HDX is reversible and occurs at sec-to-min timescale [22], which may be competitive with off-rates of protein-ligand equilibrium, giving convergence in HDX for bound and unbound at long times. Further, D2O dilution decreases protein concentration, making tight binding challenging. Back exchange and D scrambling during fragmentation challenge the extension beyond the peptide level.

Fast photochemical oxidation of proteins (FPOP) labels proteins with hydroxyl radicals generated by hydrogen peroxide photolysis [23,24,25,26,27,28,29]. Hydroxyl radicals irreversibly label solvent-accessible areas [23, 24] in sub-milliseconds, faster than changes in protein conformation [24, 26, 27]. This ensures that FPOP labeling will not distort or compete with the binding equilibrium. Because the labeling is irreversible, experiments can be executed off-line, where concentrations are no longer determined by the MS detection limit. Moreover, there are no back exchange and scrambling, facilitating detection at the amino-acid level [26, 28, 29].

Here, we report a novel FPOP-based ligand titration method, protein-ligand interaction by Ligand Titration, Fast Photochemical Oxidation of Proteins and Mass Spectrometry (LITPOMS). This new approach allows measurement of binding stoichiometry and site-specific binding constants for any protein-ligand system that experience a change in solvent-accessible area upon binding. Such measurements also access the equilibrium composition indirectly and, importantly, provide the affinity in the liquid phase. LITPOMS overcomes disadvantages of PLIMSTEX, as stated above, providing another significant strategy to assess binding stoichiometries, affinities, and dynamics.

Results and Discussion

To demonstrate LITPOMS, we chose calmodulin-melittin as a model. Melittin (Mel) binds to calcium-bound calmodulin (holo-CaM) at a ratio of 1:1 with a Kd of 3 nM [30,31,32,33]. Crosslinking suggests that melittin binds at the N-terminus, C-terminus, and the central linker region of the holo-CaM [31, 32]. In the current study, we equilibrated holo-CaM with melittin overnight in Tris buffer (pH = 7.4). A series of aliquots were prepared at [Mel]:[Holo-CaM] of 0–3 to fulfill the titration process. Hydrogen peroxide and L-histidine were added followed by a pulsed 248 nm laser irradiation to footprint the complex. The sample was introduced by a syringe pump into a capillary tubing with a transparent window. L-histidine works as a hydroxyl radical scavenger to control the lifetime of the radicals. The footprinted sample was collected at the end of the capillary in a solution of L-methionine and catalase to quench any residual oxidants. Global-level responses were by direct measurement of the intact complex with a Bruker Ma** the topology and determination of a low-resolution three-dimensional structure of the calmodulin-melittin complex by chemical crosslinking and high-resolution FTICRMS: direct demonstration of multiple binding modes. Biochemistry. 43, 4703–4715 (2004)" href="/article/10.1007/s13361-018-2076-x#ref-CR32" id="ref-link-section-d250125864e796">32]. The coupling of FPOP with titration and MS permits characterization of protein-ligand binding stoichiometry, binding sites (peptide and amino-acid levels), and site-specific binding constants, even for a tight binding system like melittin:holo-CaM.

In conclusion, the high-concentration mode for determining stoichiometry and the low-concentration mode for locating binding sites and determining site-specific affinities make LITPOMS promising for characterizing protein-ligand binding for high picomole quantities of protein. Moreover, the irreversible labeling allows rigorous post-labeling digestion without erasing any labeling information (e.g., from back exchange). LITPOMS is also readily compatible with various buffers, pH, salts, lipid-based media (e.g., nano and pico disks), and binding affinities (ranging from nM to μM), making it generally applicable even to membrane proteins. The approach should not be affected by high off-rates for weaker binding systems as for HDX, but that remains to be established. Importantly, residue-level analysis becomes possible owing to the irreversible labeling that will be maintained for MS/MS, and more complete residue locations will be enabled by complementary footprinters (e.g., CF3∙ [34], carbenes [35]). Although we demonstrated the applicability to tight binding (nmolar) of 1:1 binding system, we plan to extend the method to other, more complex binding systems with stoichiometries greater than 1:1.