While the research focus of the Sinz lab is on advancing the cross-linking mass spectrometry (XL-MS) approach for structural and functional proteomics applications, other techniques in structural mass spectrometry are also being increasingly used. These techniques comprise native mass spectrometry, hydrogen-deuterium exchange (HDX), ion mobility-mass spectrometry, and footprinting methods.

1. Protein Structure Analysis by Mass Spectrometry

Until a few years ago, and before the advent of system-wide approaches, analysis by MS-based methods usually required the purification of the protein or protein complex of interest. Notably, the degree to which the sample is purified dictates the quality of the data that are obtained during subsequent MS analysis. For obtaining large yields based on recombinant expression systems are usually applied. As host cells, both prokaryotic and eukaryotic systems can be exploited nowadays and expression vectors for many host cells are commercially available. Highest yields of recombinant protein (up to 80 % of total cellular protein, depending on protein and expression vector) can be retrieved from the Gram-negative bacterium Escherichia coli as host system. However, post-translational modifications and associated interacting protein partners are lost unless they are co-expressed. To meet the latter challenge, various biochemical approaches have been developed for the isolation of endogenous protein complexes directly from cells. For example, one of the subunits or components of the protein complex can be labeled via genetic engineering with a N- or C-terminal peptide tag (FLAG-, His-, GST-, TAP-tag). The high-affinity tag interacts with another component that allows the isolation of the entire protein complex by affinity chromatography under conditions that maintain the integrity of the protein complex ²³. In the last years, adoption of these approaches has caused a significant shift in the development of MS-based methods towards the study of highly complex protein mixtures, including system-wide studies in cellulo and in vivo ²⁴.

In general, MS measures the mass-to-charge ratio (m/z) of an ion. Analytes, such as peptides, proteins or medium- to low-molecular weight molecules, are ionized in a first step and then transferred into the gas phase. The ions are subsequently separated, depending on their m/z ratios, in the mass analyzer and ultimately reach the detector. One of the main ways to ionize peptides and proteins is electrospray ionization (ESI) ²⁵, which has evolved as the method of choice for protein analysis. A complementary soft ionization technique, employing matrix-assisted laser desorption/ionization (MALDI), can also be used, although its application has lost some of its importance for protein analysis during the last few years ^26,27,28,29. Hybrid instruments that are based on a combination of two or more mass analyzers allow performing tandem MS (MS/MS) experiments ³⁰. In this type of analysis, a specific m/z ratio is isolated from the rest of the ions and activated in a collision cell to induce dissociation of the molecule into smaller components. The resulting fragment ions are then interrogated in a second mass analyzer. Several types of mass analyzers, including time-of-flight (TOF), quadrupole, orbitrap, are currently available, as are various dissociation methods, e.g., collision-induced dissociation (CID), electron-transfer dissociation (ETD), electron-capture dissociation (ECD), surface-induced dissociation (SID), and ultraviolet-photodissociation (UV-PD) ^31,32. As a result, a number of configurations of mass spectrometers, each of which vary in their capabilities and limitations, can be designed for different purposes and scientific needs.

2. Native MS

Intact proteins and macromolecular complexes can be studied under non-denaturing conditions in the gas phase of a mass spectrometer through gentle ionization, such as ESI ³³. This technique has been termed “native MS” and can provide information on the stoichiometry of non-covalent partners including proteins and non-proteinaceous molecules ³⁴. Native MS provides information on the preserved overall architecture and the conformational changes of complexes ^35,36. When used in combination with high-resolution mass spectrometers, native MS has become a valuable technique in the structural proteomics toolbox ³⁷, with one major benefit being able to capturing all coexisting subpopulations in a single spectrum. This enables analysis of different conformational states within multimeric protein complexes that are prompted by ligand bonding and aids elucidation of allosteric mechanisms ³⁸.

Over the last decade, native MS has become a powerful tool for the study of challenging membrane protein complexes and their associated lipids that stabilize a complex ³⁹ or are required for its formation or regulation ^40,41. The first structural study with native MS of a membrane complex preserving bound ligands was performed by protecting the complex in micelles using N-dodecyl-ß-D-maltoside as detergent ⁴². Although new detergents, which are compatible with native MS, are continuously under development ⁴³, alternative approaches are currently being investigated to mimic the phospholipid bilayer of cell membranes, such as nanodiscs ⁴⁴ and more recently lipodisqs ⁴⁵. A significant recent advance has involved the analysis of membrane complexes ejected from lipid vesicles obtained from endogenous biological membranes ⁴⁶.

Despite the great potential of native MS for investigating intact proteins, sample preparation is often tedious as it requires purified protein(s), often involving multiple biochemical isolation steps prior to the last buffer-exchange into the volatile buffer needed for native MS ⁴⁷. Recently, however, a novel approach has been developed that alllows a more direct analysis of overexpressed proteins from crude extract with minimal sample handling ^47,48. This approach is still in its infancy and currently depends on the system used for protein production. However, it will be valuable for the study of proteins at the endogenous level in cells, also taking into account naturally occurring post-translational modifications.

3. Ion Mobility MS

Ion mobility (IM) spectroscopy measures the mobility of an ion in a buffer gas present in a drift tube under the influence of a weak electric field. The time required for an ion to pass through the tube depends on its collision frequency with the drift gas molecules, and thus correlates with its size and shape. This correlation allows derivation of the rotationally averaged collisional cross section (Ω, CCS) of the ion through a mathematical equation ^49,50. The CCS derived from the experiment is used to infer structural information, with the deduced information being dependent on the method used ⁵¹. Consequently, in order to obtain comparable measurements on different IM platforms, calibration and harmonization of data reporting are important ^49,52. IM-MS has served for analytical purposes for more than 50 years, conferring an additional ion separation dimension with gas-phase mobility separation in the IM and m/z determination in the MS ⁴⁹. During the past 15 years IM-MS has experienced an exponential increase in interest in several scientific fields, particularly, for “omics” studies ⁵³. The rapid spread of IM-MS began with the introduction of the first commercially available mass spectrometer coupled with IM separation ⁴⁹ and evolved with the subsequent development of different IM technologies, such as drift tube (DTIMS), traveling wave (TWIMS), trapped (TIMS), and field asymmetric ion mobility spectroscopy (FAIMS) ³⁷. These IM platforms differ with respect to gas flow direction, electrostatic and to the electrodynamic fields applied ⁵⁴. It should be noted that the FAIMS technique, which relies on ion separation field strength, does not allow a reliable CCS measurement ⁵⁵. The most successful coupling of IM is in combination with nano-ESI and native MS ⁵⁶, which exploits the combined separation of isomeric ions of different sizes and shapes of all intact coexisting complexes based on their distinct CCS and m/z ratios.

IM-MS of intact protein complexes with its possibility of inducing gas-phase dissociation of the multimeric complex provides important insights into stoichiometry, size, and architecture of protein assemblies (Figure 1) ^10,57,58,59.

A more recent approach shows that the resolution of protein isoforms with small differences in their structural arrangement can be rapidly achieved using their characteristic unfolding behavior and stability after collisional activation before the drift tube ⁶⁰. Increasing collisional energies are applied to the ion of interest and different CCS values of the activated ion are recorded and plotted as a function of the applied voltage, resulting in a specific collision-induced unfolding (CIU) fingerprint of the molecule ⁶¹.

Just recently, IM-MS has been coupled with XL-MS (Figure 2). This combination allows a deeper and more sensitive identification of cross-links ^62,63,64, especially in a highly complex environment, paving the way for another successful IM combination.

4. Hydrogen-deuterium exchange MS

Hydrogen-deuterium exchange combined with mass spectrometry (HDX-MS) presents the most popular protein footprinting method so far developed. HDX monitors the rates of exchanging hydrogen by deuterium atoms at the amide backbone of proteins from solvent D₂O in solution ^65,66. These exchanges reflect the solvent-accessible surface area (SASA) and are recorded by a mass spectrometer as a function of time. Thus, HDX-MS provides information on solvent accessibility, secondary and tertiary structures, conformational changes, protein states and binding interfaces ^67,68. HDX-MS is able to capture protein dynamics and allosteric regulation in solution of a protein or a protein complex, and as such is a local sensor for conformational changes at single amino acid resolution ⁶⁹ (Figure 3).

Although HDX-MS has been used for 30 years ⁷¹, the development of mass analyzers and data analysis software - which remains the bottleneck - have only recently rendered HDX-MS a popular tool among the structural proteomics methods72 enabling studies ranging from single proteins ⁷¹ up to viral particles ⁷³. Some drawbacks of HDX are its propensity for back-exchange reactions and D/H scrambling effects ⁶⁹. In order to quench deuterium exchange and reduce the back-exchange after labeling, the pH value has to be adjusted to ~2.5 and the temperature has to be lowered to ~0°C ⁷⁴. However, these conditions cause suboptimal efficiency of downstream steps, such as disulfide bonds reduction and protease digestion. The digestion of glycosylated proteins is especially impaired, and among those proteins, the most studied are antibodies and their interactions with ligands ⁷⁵. To overcome these weaknesses, novel approaches are being evaluated, implemented and integrated in order to be used online and in an automated way, e.g. electrochemical reduction76 and by use of different deglycosylation enzymes that are active at low pH ⁷⁷.
Meanwhile, the hydrogen-scrambling effect that is mainly instrument-dependent, has been addressed with improvements in mass spectrometer components and electron-based fragmentation methods ⁷⁸.
In the last years, use of HDX-MS has experienced widespread application in drug development ⁷⁹ where epitope mapping in antibodies ⁸⁰, conformational changes in important targets proteins, such as G-protein-coupled receptors in the presence of ligands ⁸¹, or biopharmaceutical comparability are critical aspects ⁸².

5. Chemical Footprinting

Different covalent protein labeling methods such as HDX (see above), covalent labeling, covalent protein painting (CPP), protein painting, hydroxyl radical footprinting (HRFP), and fast photochemical oxidation of proteins (FPOP) are united under the umbrella of MS-based chemical footprinting approaches70. Chemical footprinting relies on covalent labeling of a protein or protein complex in solution to cause a mass shift by the label, which is then detected by MS. The chemical modifications allow probing the structure of proteins, conformational changes, and binding sites in a complex ^70,83,84.

Covalent Labeling relies on a covalent chemical reaction targetting side chains of specific amino acids, such as lysines (containing amine groups) or cysteines (containing thiol groups) ^70,85. The labeling reagents are chemical compounds that, in contrast to free radicals, create mass tags on the molecules, which are identified in the subsequent MS analysis ⁷⁰.
Covalent labeling combined with MS has a lot in common with XL-MS and the two techniques complement each other perfectly: XL-MS delivers distance constraints between two amino acids, simultaneously giving information on the orientation of interacting proteins (see below), while covalent labeling indicates the protected areas of the proteins involved in the interaction ^85,86.
Covalent labeling has proven highly useful for elucidating the structures of membrane proteins and their interactions87, as well as determining protein-ligand affinities ⁸⁸. Moreover, it has also been applied to determine the energetic barrier of a structural change that initiates protein fibril formation ⁸⁹.

Protein Painting employs small-molecules that bind non-covalently to the surface of proteins and therefore block tryptic cleavage sites. However, a limitation of this method is that only peptides involved in the binding interface to another protein, or a small ligand, will be protected from being labeled and will appear as tryptic peptides in subsequent MS analysis ⁹⁰. The major advantage of covalent protein painting (CPP) is that it directly determines the specific functional groups in proteins, i.e., lysines or cysteines, which are available for a covalent modification on the surface of a protein. CPP is therefore synonymous with covalent labeling. Recently, CPP has been employed in a quantitative approach of dimethylating lysine residues in proteins with “light” (non-deuterated) and “heavy” (deuterated) reagent to detect structural changes of proteins in human postmortem brain samples of patients with neurodegenerative diseases ⁹¹.

Hydroxyl Radical Protein Footprinting (HRPF) is an irreversible, non-specific oxidative reaction that relies on hydroxyl radicals. The solvent-exposed side chains of the protein complex react on the order of milliseconds with the radicals providing direct information on the topology of the protein ⁷⁰. The modification the amino acid side chain undergoes differs for the individual amino acids ⁹². There are several methods that can be employed to produce hydroxyl radicals, with the most commonly used for protein footprinting being Fenton and Fenton-like chemistry ⁹³, synchrotron water radiolysis ^94,93, and laser photolysis of hydrogen peroxide ⁹⁵. Recently, two novel benchtop approaches have been proposed: Plasma-induced modification of biomolecules (PLIMB) ⁹⁶; and the Flash OXidation (FOX) Protein Footprinting System ⁹⁷, which will broaden the array of HRPF tools available for protein structural studies.

Fast Photochemical Oxidation of Proteins (FPOP) is a rapid HRPF method, taking advantage of covalent protein labeling, which occurs on the microsecond timescale. Radicals are produced from H₂O₂ through photolysis induced by a laser at 248 nm in a microflow system ^95,98. FPOP has been applied in a series of investigations, for example, to study protein folding kinetics in the microsecond range ⁹⁹, epitope mapping ¹⁰⁰, refolding processes of viral fusion proteins ¹⁰¹, and characterizing membrane proteins in their native environment ¹⁰². This broad range of applications makes FPOP a promising and exciting tool for structural proteomics studies ^83,84.
Clearly, one of the most innovative applications of FPOP is in vivo footprinting, where accessible protein regions are labeled directly inside cells without disrupting cellular structures ¹⁰³. This method has been successfully applied for intact cells and even for living organisms, such as Caenorhabditis elegans ¹⁰⁴ (Figure 4). In particular, in vivo FPOP has benefitted from a combination with microflow systems98 or static platforms ¹⁰⁵.