Synchrotron X-ray footprinting as a method to visualize water in proteins

Abstract

The vast majority of biomolecular processes are controlled or facilitated by water interactions. In enzymes, regulatory proteins, membrane-bound receptors and ion-channels, water bound to functionally important residues creates hydrogen-bonding networks that underlie the mechanism of action of the macromolecule. High-resolution X-ray structures are often difficult to obtain with many of these classes of proteins because sample conditions, such as the necessity of detergents, often impede crystallization. Other biophysical techniques such as neutron scattering, nuclear magnetic resonance and Fourier transform infrared spectroscopy are useful for studying internal water, though each has its own advantages and drawbacks, and often a hybrid approach is required to address important biological problems associated with protein-water interactions. One major area requiring more investigation is the study of bound water molecules which reside in cavities and channels and which are often involved in both the structural and functional aspects of receptor, transporter and ion channel proteins. In recent years, significant progress has been made in synchrotron-based radiolytic labeling and mass spectroscopy techniques for both the identification of bound waters and for characterizing the role of water in protein conformational changes at a high degree of spatial and temporal resolution. Here the latest developments and future capabilities of this method for investigating water-protein interactions and its synergy with other synchrotron-based methods are discussed.

Keywords: bound water; hydroxyl radical labeling; mass spectrometry; protein conformation; protein modification.

Figure 1

Major reactions scenarios for X-ray radiolysis in dilute protein samples. (a) Schematic showing the position of bulk-, surface- and bound-water (cavity- and buried-water) (light blue) in a protein molecule (dark blue). (b) Radiolysis of water and the timescale of sequence of events. Reproduced from Gupta et al. (2014 ▸) and Liljenzin (2002 ▸). (c) The location of hydroxyl radicals (red) generated in situ from ionization or activation of water by X-ray irradiation. The OH radicals react with nearby side chains in close proximity, and yield covalent modifications on the protein side chains (yellow). Radiolysis of bulk water starts with the ionization of water on the time scale of 10⁻¹⁶ s. The key product, OH, diffuses (10⁻⁷ s) out in the bulk (red arrows) and undergoes reactions with other OH, buffer molecules and protein side chains (bimolecular reactions are indicated by black arrows, and approximate values of the rate constants for different reactions are shown). The rapid counterproductive reactions, such as OH—OH recombination, as well as various other reactions, scavenge OH and reduce the concentration of OH in the bulk. Thus a sufficient X-ray dose or high-flux-density beam is needed to maintain a steady concentration of OH that will lead to a detectable yield of side-chain modification on the protein in solution. In contrast, OH radicals formed from activation of a bound water can react faster with side chains in proximity because of the translational and rotational ordering of water and because fewer scavenging reactions by other OH or highly reactive buffer constituents are available. Radiolysis of water also produces electrons, which rapidly become solvated and react with O₂ to produce superoxide radicals. In general, the reactivity of side chains to solvated electrons is lower than to hydroxyl radicals (Xu & Chance, 2007 ▸). Peroxides and superoxides undergo slow reactions with protein side chains and are quenched as described in §2.1 and §2.3. Solvated electrons consume O₂, which is also required for side-chain labeling by OH radicals (Fig. 2 ▸); thus short irradiation time as well as high flux density are the key factors for success of the XF-MS experiment. Reproduced in part from Gupta et al. (2014 ▸).

Figure 2

Reaction schemes for modification of side chains by OH. The hydrocarbon side chain (of aliphatic non-polar and polar amino acids) predominantly undergoes hydrogen abstraction by OH to give carbon-centered radicals, which then react with molecular O₂ under aerobic conditions and subsequently form stable hydroxylated (+16 Da) or carbonylated (+14 Da) products. Aromatics and sulfur-containing side chains directly undergo hydroxylation by OH followed by reaction with O₂ under aerobic conditions to form stable hydroxylated products (+16 Da) and other oxygen adducts. Aromatics and sulfur-containing side chains can be labeled by ¹⁸O (red) from radiolysis of H₂ ¹⁸O water.

Figure 3

Schematic of the X-ray footprinting method using synchrotron X-ray radiolysis and mass spectrometry. (a) Overall method, and (b) case study for a protein that undergoes a conformational change from the closed to the open state. The protein is covalently modified after a series of X-ray irradiations of the order of microseconds followed by rapid quenching by methionine amide. Irradiated protein is digested with proteases to generate peptide fragments of known mass. Digested protein is analyzed by reverse-phase liquid chromatography coupled with electrospray mass spectrometry (LC-ESI-MS), in which peptides are separated in the total ion chromatogram (TIC) or mass chromatogram. For any peptide fragment the unmodified and modified m/z is extracted and visualized by selected ion chromatogram (SIC). High-resolution mass spectrometry is used to identify the unmodified and modified peptide fragments by their accurate m/z and isotope distribution. The site of modification is identified from the mass assignment of the y and b fragment ions from the tandem mass spectrometry (MS/MS) of the corresponding peptide. The extent of modification for the series of irradiation points are quantified from the SICs of unmodified and modified pepide fragments. The fraction of unmodified peptide versus exposure time (dose-response or DR-plot) provides site-specific modification rate constants (k s⁻¹). The rates are compared among different sample conditions, and their ratios, which are independent of intrinsic reactivity, account for the degree in solvent accessibility changes due to any conformational transition/interactions. The final results are mapped onto available structures or used as constraints for structural modeling.

Figure 4

Detection of the site of bound water interaction by temperature-dependent XF-MS. The colored surfaces on the X-ray crystal structures of cyt c (PDB 1HRC) (a) and ubiquitin (PDB 1UBQ) (b) indicate the side-chain residues that are consistently modified at room temperature and/or frozen conditions. Blue indicates the residues that show a 13- to 200-fold decrease in modification upon freezing; violet indicates a 3- to 10-fold decrease; and red indicates minimal to no change (<2-fold) in the modification rate when the sample is frozen as compared with room temperature. In cyt c the residue Y67, which also showed minimal to no change, is completely buried inside the heme cavity and not visible in these orientations. Reproduced from Gupta et al. (2012 ▸).

Figure 5

Determination of residence time of tightly bound water by XF-MS. (a) Rapid mixing combined with ¹⁸O-mediated hydroxyl radical labeling to monitor the time-course of exchange of water in cyt c. LC-ESI-MS is used to identify and isolate the modified peptides, targeted MS/MS is used to identify the sites of ¹⁸O labeling, and zoom scans are used to quantify the ratio of ¹⁸O- versus ¹⁶O-labeling at various mixing delays. (b) Zoom scans for singly protonated peptide 61–72 showing the decrease in the abundance of the 2m/z shifted ¹⁸O monoisotopic mass (arrow) that corresponds to the water exchange at M65 and Y67 with increase in the mixing delays. (c) Progress curves (circles and error bars) of water exchange for the ¹⁸O labeled side-chain residues. The solid line represents the fit to a single exponential function. Residues W59 and F36 have exchange that is complete at the first measurement, while the rates of exchange of C14, C17, F46, Y48, M65, Y67 and M80 are discretely measured. (d) Sites of ¹⁸O-modifications are visualized from the crystal structure 1HRC using PyMOL (DeLano Scientific). The ¹⁸O-labeled residues (light blue) in and around the heme (light pink) crevices, and the position of residue T78 (gray) and conserved waters (cyan spheres) HOH112, HOH139 are shown in two orientations of the cyt c molecule. Reproduced from Gupta et al. (2012 ▸).

Figure 6

XF-MS probes bound water mediated signal transfer pathway in bovine rhodopsin. (a) DR-plots of dark (black) versus meta II (red) for the peptide p333–348 (at the solvent exposed cytoplasmic side) and p160–164 (at the TM region) for modified residues as indicated. (b) Pictorial summary of relative solvent accessibility changes for the photoactivation of dark to meta II state. Residues with rate constants >0.1 s⁻¹ are shown as sticks. The color coding represents the ratio of rate constants between meta II and rhodopsin. Conserved transmembrane waters are shown as cyan spheres. The changes in rates of modification reflect local structural changes inside the TM domain upon formation of meta II. The results demonstrate disruption and reorganization of multiple close-packing interactions, mediated by both side chains and bound waters. The information is transmitted from the chromophore (ligand-binding site) to the cytoplasmic surface for G-protein activation. Results from Angel, Gupta et al. (2009 ▸).

Figure 7

XF-MS identifies conformation changes during gating of a K⁺ ion channel, KirBac3.1. (a) Cross-sectional view showing multiple surface-exposed and buried cavities in close KirBac3.1 (PDB-1XL4). TM1 (purple) and TM2 (green) denote transmembrane helix-1 (outer) and transmembrane helix-2 (inner), respectively. Pore helix and side helix are colored blue and red, respectively. (b) Solvent accessibility changes from the closed to the open conformation in KirBac3.1 are visualized on the structure of closed KirBac3.1, where the subunits are represented by different colors. The modified residues are shown by sticks. Color coding indicates the changes in the modification rates or solvent accessibilities upon transition from the closed to the open state. Residues that undergo increased interactions with water due to changes in the structure of the channel in the open state show dramatic increase in labeling efficiency. The results support the proposed existence of three potential gates within the channel. Results from Gupta et al. (2010 ▸).

Figure 8

XF-MS probes proton-coupled Zn²⁺ transfer mechanism in Zn transporter YiiP. (a) Bar plot showing radiometric water accessibility changes in response to Zn²⁺ binding measured by ratio (R) of OH labeling rates for residues with increase (blue), decrease (red) and no change (grey) in accessibility after rapid Zn²⁺ exposure. (b) Time courses of water accessibility changes for the indicated residues. Solid lines represent single exponential fits, which provide rate constants of conformational changes associated with Zn²⁺ binding and translocation. (c) X-ray footprinting reveals a hydrophobic gate at residue L152, which controls the opening of the inner cavity water pathway for zinc-proton exchange in the YiiP transporter. The cross-sectional view shows the position of TM helices, which separate two cavities at the intra-cellular (IC) and extra-cellular (EC) sides. Residues are color coded as in (a). XF-MS results suggest the protein conformational change alternates the membrane-facing on–off mode of zinc coordination (in D49) and protonation–deprotonation (H153) of the transport site in a coordinated fashion. (d) The red arrow indicates the proposed water pathway, which connects EC with IC after excluding the residue L152 from the surface drawing of the TM helices. Results from Gupta et al. (2014 ▸).

Figure 9

XF-MS probes bound water mediated signal transfer pathway in OCP. (a) Solvent accessibility changes from dark adapted-OCP^O to illuminated-OCP^R are visualized on the structure of OCP^O (3MG1). The modified residues are represented by sticks and the carotenoid is shown in pink. The color coding represents the ratio of rate constants between these two states. (c) The proposed signal propagation pathway from the carotenoid through the water side-chain hydrogen-bonding network to the protein surface that facilitates carotenoid shift, dissociation of NTD-CTD and detachment of the N-terminal helix from CTD. Conserved waters are shown in spheres and color codes indicate their depth from the surface of OCP. The modified residues are shown by green sticks. The results demonstrate disruption and reorganization of multiple close-packing interactions, mediated by both side chains and bound waters. (d) Ab initio bead reconstructions (gray volume) based on the SAXS results are shown for OCP^O and OCP^R. The subunit of OCP^O from the crystal structure is docked into the volume envelope with the far N-terminal helix (red), NTD (purple) and CTD (green). The SAXS results show dissociation of NTD and CTD. (d) Schematic of the photoactivation of OCP^O showing regions with the largest conformational rearrangement associated with changes in the hydrogen-bonding network and water rearrangements. Reproduced from Gupta et al. (2016 ▸).