Analytical applications of gas-phase ion chemistry enabled by mass spectrometry

Abstract

Analytical mass spectrometry (MS) has been employed to study a wide variety of analytes, including metabolites, lipids, pharmaceutical compounds, pesticides, petroleum, peptides, proteins, protein complexes, nucleic acids, and glycans. The field of gas-phase ion chemistry in mass spectrometry studies the impacts of the chemical behaviors and properties of ions produced from these samples, along with instrumental and methodological parameters of the MS experiment, in determining the appearance of the resulting mass spectra. This subfield includes the dynamic interactions of the ions with neutral molecules, electrons, photons, and other ions. These interactions are particularly useful in tandem mass spectrometry (MS/MS or MSⁿ) experiments, which provide high specificity and enable analyte structural characterization by fragmenting a compound of interest and then analyzing the product ions. These bimolecular interactions can also result in non-dissociative processes, leading to ion transformation, charge alteration, or the formation of ion/molecule and ion/ion complexes. Such reactions offer valuable insights into chemical behavior across various reaction environments by simulating those conditions within the mass spectrometer. This information can then be used to make novel inferences about the sample and promises to inform MS studies in areas such as metabolomics, lipidomics, drug pharmacology, exposomics, proteomics, glycomics, genomics, and environmental, supramolecular, and interstellar chemistry. In this review, we highlight novel applications of tandem mass spectrometry that have been published in the past ten years, focusing on reactions that take place in the gas phase in a reduced-pressure environment (i.e., ion/neutral atom/molecule, ion/electron, ion/photon, and ion/ion reactions), largely after the ionization step.

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1. Introduction

Gas-phase ion chemistry is important in determining the spectra produced from tandem mass spectrometry (MS/MS or MSⁿ) experiments. Gas-phase interactions between ions and neutral molecules, electrons, photons, or oppositely charged ions can cause chemical and physical changes to the analyte ion, such as altering the charge state and breaking bonds, which can produce structural information on the analyte ion. A wide variety of gas-phase reactions have been reported in recent years, leveraging a number of advantages, including proceeding faster than equivalent reactions in the condensed phase and enabling high reaction purities via precise computer-controlled timing and admission of the reactants. Herein, we briefly discuss the fundamental principles and mass spectrometry instrumentation of these gas-phase reactions before focusing on recent novel applications. More comprehensive reviews on reaction mechanisms can be found elsewhere.^1–5 This introduction of gas-phase reactions is not intended to be an exhaustive description of these fields. Instead, this prologue is meant to introduce these reactions and provide a context for understanding the recent applications of this technology to analytical challenges highlighted in later sections. Readers are directed to other review articles for more in-depth discussions of ion/neutral atom/molecule reactions,^6–8 ion/electron reactions,⁹ ion/photon reactions,^10–13 and ion/ion reactions,^14–17 as well as the specific gas-phase ion chemistry of lipids,¹⁸ glycans,¹⁹ and proteins.⁴ We aim to describe a wide range of gas-phase chemistry applications to demonstrate the versatility of the reactions and highlight the potential for broader adoption. The types of reactions discussed in this review are summarized in Fig. 1.

Fig. 1 Summary of mass spectrometry-based gas-phase chemistries discussed. (a) Ion/neutral atom/molecule reactions involve interactions of analyte ions with inert gas, ozone, radical, and deuterated molecules. (b) Ion/electron reactions involve radical-directed dissociation of analyte ions following electron capture or detachment. (c) Ion/photon reactions involve the deposition of internal energy through photoabsorption, followed by the dissociation of the analyte. (d) Ion/ion reactions involve interactions between oppositely charged ions, followed by particle transfer or covalent bond formation.

2. Applications of ion/neutral atom/molecule reactions

Ion/neutral atom/molecule reactions occur between analyte ions and inert or reactive neutral gas molecules.²⁰ In this review, we discuss the recent applications of four different dissociative or non-dissociative ion/neutral atom/molecule reactions, each reaction presenting unique gas-phase chemistries and requiring specific instrumentation: collision-induced dissociation (CID), ozone-induced dissociation (OzID), ion/neutral radical reactions, and hydrogen/deuterium exchange (HDX). CID is the most commonly used form of ion/neutral atom/molecule reaction in mass spectrometry and is available on most MS instrumentation, representing the most widely accessible dissociation method in MS/MS for structural identification of a variety of samples.²¹ CID generally involves collisions of accelerated ions with inert buffer gas atoms and molecules, such as He, Ar, or N₂, which converts the relative translational energy into the internal energy of the ions, resulting in fragmentation from vibrationally excited states of the electronic ground state.²² Another dissociation method, OzID, is increasingly utilized to induce oxidative dissociation using reactive neutral reagents (i.e., ozone). OzID is mainly used to locate carbon–carbon (C–C) double bonds in a wide variety of samples, including unsaturated fatty acids (FAs),²³ glycerolipids,²⁴ glycerophospholipids,^24,25 cholesteryl esters,²⁶ and sphingolipids,^27,28 as well as in amphiphilic small metabolites containing olefinic hydrocarbon chains²⁹ and isoprene chains.³⁰ Ion/neutral radical reactions also involve using reactive neutral reagents (e.g., radical molecules) to facilitate dissociative or non-dissociative reactions. These reactions, for example, can be used to study chemistry in the atmosphere and interstellar space, where naturally occurring free radicals are common due to processes such as ultraviolet (UV) photolysis.^31,32 HDX is a non-dissociative, deuterium labelling method that provides structural information for various analytes, including carbohydrates,^33–35 small molecules,^34,36 and proteins.^37–39 Ion/neutral atom/molecule reactions using a variety of other neutral reagents, including hydrocarbons,⁴⁰ non-radical reactive oxygen/nitrogen species,⁴¹ organic acids,⁴² and aryl radicals⁴³ are discussed elsewhere.

2.1. Collision-induced dissociation

The type of instrument determines the range of accessible energies and timescales of CID experiments, which in turn determines the types of fragment ions that can be generated. For example, quadrupole ion traps and Fourier transform ion cyclotron resonance (FT-ICR) traps are typically operated under multiple collision conditions at low energies ranging from <1 to 100 eV and timescales of tens of milliseconds to seconds.⁴⁴

[M + nH]ⁿ⁺ + N (gas) → [M + nH]ⁿ⁺* + N* → fragmentation (1)

The internal energies generated from these collisions progressively accumulate with each collision, eventually exceeding a dissociation threshold within the ion and resulting in fragmentation at the weakest bonds via the lowest energy paths.²² Conversely, sector and time-of-flight instruments are generally operated at keV collision energies under single collision conditions, resulting in less selective dissociation of analytes.²¹ CID employing keV collisions is less common in recent years and is thus beyond the scope of this discussion.⁴⁵ In this review, we instead briefly highlight applications of low-energy CID for covalent and noncovalent bond dissociation in the analysis of lipids, peptides, and proteins.

Low-energy CID of lipid ions generated by so-called soft-ionization methods (i.e., those that tend to produce intact ions with minimal fragmentation) such as electrospray ionization (ESI) and matrix-assisted laser desorption/ionization (MALDI) preferentially breaks the most labile bonds. For example, this process results in head group loss from protonated glycerophospholipids and ester bond cleavage in deprotonated glycerophospholipids, enabling the identification of lipid subclasses and fatty acyl chains, respectively.¹⁸ However, this approach generally does not provide information on intrachain modifications, such as the sites of C–C double bonds,^46,47 which necessitates changing the ion type. Chemical derivatization of C–C double bonds via Paternò–Büchi reaction,⁴⁸ epoxidation,⁴⁹ or oxidation⁵⁰ before ionization allows unsaturation site-specific cleavages upon CID and produces diagnostic fragment ions. Fragmentation of unsaturated fatty acyl chains via radical-directed dissociation can also aid in unambiguously identifying C–C double bonds. For example, lipid ions noncovalently modified with bicarbonate during ESI^46,51 or covalently derivatized with 2,2,6,6-tetramethylpiperidine-1-oxyl-benzyl (TEMPO) reagents before ionization⁵² generate radical ions upon CID, which undergo nonselective dissociation along fatty acid chains. TEMPO-assisted radical-directed dissociation of phosphatidylethanolamines (PEs) produces extensive FA chain fragment ions by radical formation via the homolysis of the labile bond between the benzylmethyl carbon and the piperidine oxygen in the TEMPO reagent during CID.⁵² The presence of doublet fragment ions differing by 2 m/z units following radical-directed dissociation suggests that the loss of two hydrogen atoms occurs at C–C double bonds during their cleavage, thus pinpointing the locations of these unsaturation sites. This method effectively differentiates phosphatidylethanolamines with various combinations of intrachain modifications from an E. coli lipid extract (Fig. 2).⁵² Overall, this demonstrates the importance of the ion type in determining the type and extent of fragmentation observed during CID.

Fig. 2 (a) The PE profile of E. coli via neutral loss scan of 141 Da in positive-ion mode. (b) Extracted-ion chromatogram of m/z 832 from MS² CID of [PE^TPN 34:2 − H]⁻ (m/z 988) and the corresponding spectra for the peaks eluted at (c) 30.7 and (d) 31.5 min, respectively. (e) Identified PEs from E. coli with relative ion abundances normalized to PE 33:1. The locations of C=C and cyclopropane in fatty acyls are indicated at the top of (e). TPN: 3-(2,2,6,6-tetramethylpiperidin-1-yloxymethyl)-picolinic acid 2,5-dioxopyrrolidin-1-yl ester. Adapted/reproduced from Journal of American Society for Mass Spectrometry, 2022, 33, 714–721 (ref. 52) with permission from the American Society for Mass Spectrometry, Copyright 2022.

Low-energy CID of peptide ions generated via ESI leads to amide bond fragmentation, producing b-/y-type fragment ions along the backbone.⁵³ Labile post-translational modifications (PTMs), such as sulfation and phosphorylation,^54,55 are generally not retained due to the slow-heating nature of multiple low-energy collisions. However, CID of peptide ions derivatized with TEMPO reagents leads to homolytic bond cleavage in the reagent and produces radical ions. Subsequent radical-directed dissociation results in the production of a˙-/b-/c-/x-/y-/z˙-type fragment ions while preserving PTMs.^54,55 Additionally, charge-remote fragmentation via radical-directed dissociation results in more extensive fragmentation across the peptide backbones^56–58 than traditional charge-directed cleavages via mobile protons in CID.^53,59 Charge-remote fragmentation is particularly useful in enhancing the sequence coverage of large native proteins and protein complexes with relatively low charge states and, thus, low CID efficiency.⁶⁰ For example, TEMPO-assisted radical-directed dissociation offers approximately 15% greater sequence coverage compared to traditional CID for bovine serum albumin (66 kDa), about 30% more sequence coverage for Concanavalin A tetramer (106 kDa), and a nearly 50% increase for β-lactoglobulin A dimer (36 kDa).⁶⁰ TEMPO-assisted radical-directed dissociation also enables 3D protein structural analysis by facilitating through-space hydrogen atom radical transfer from the radical site to distant residues in sequence that are close in space. For example, the loss of the TEMPO moiety from the N-terminal benzyl succinic acid conjugate during CID results in the formation of a benzyl radical (Fig. 3), which subsequently induces radical-directed dissociation at C-terminal residues (e.g., E64, S65, and L67) in denatured 5+ ubiquitin, indicating a spatial proximity between the N- and C-terminal regions.⁶¹ However, higher ubiquitin charge states do not show fragmentation in this region, suggesting a larger spatial distance between the two termini and a more unfolded conformation compared to 5+ ubiquitin.⁶¹ Overall, the use of free radical initiator-assisted radical-directed dissociation for analyzing lipids, peptides, and proteins highlights the versatility of CID and demonstrates that limitations to conventional CID can be addressed without necessitating modifications to the instrument. The TEMPO experiments also do not require MS³ in an ion trap and can effectively utilize beam-type CID at the MS² level,⁵⁶ allowing this approach to be implemented in various commercial instruments.

Fig. 3 (1–3): p-TEMPO–Bn–Sc–NHS conjugation to guanidinated ubiquitin and (4) FRIPS MS. The ubiquitin protein solutions were first denatured (1), and seven lysines were blocked by guanidination reactions (2), where orange circles denote guanidinated lysines. The only remaining primary amine site, N-terminus, was conjugated with p-TEMPO–Bn–Sc–NHS (3). The resulting guanidinated p-TEMPO–Bn–Sc–ubiquitin was then thermally activated, and a free radical is generated by TEMPO radical loss (4). The loss of the TEMPO radical and the generation of the benzyl radical are represented by the loss of a NHS group. Note that the denatured structure in the figure is not the actual one. Adapted/reproduced from Journal of the American Society for Mass Spectrometry, 2022, 33, 471–481 (ref. 61) with permission from the American Society for Mass Spectrometry, Copyright 2022.

While CID is primarily used for breaking covalent bonds, collisional activation is also increasingly employed to study the bonding dynamics of noncovalent interactions. For example, collision-induced unfolding (CIU) of higher-order protein structures entails the breakdown of noncovalent interactions within the protein, which is determined by measuring collision cross-sections at a range of collisional activation energies using ion mobility mass spectrometry (IM-MS).^62–64 CIU₅₀ denotes the acceleration voltage at which 50% of a protein conformer undergoes conformational transitions.⁶⁵ Determination of CIU₅₀ values for specific ligands and proteins can serve as measures of ligand affinity to binding sites⁶⁶ and protein conformational stability upon binding⁶⁷ in protein–ligand complexes. For example, the shift in CIU₅₀ of the pikromycin PKS module 6 (PikAIV) from 45.3 ± 0.0 V to 52.0 ± 0.8 V when bound to the ligand hexaketide indicates an increased conformational stability of the protein–ligand complex relative to other ligands.⁶⁸ Collisional activation can also provide information on the noncovalent interactions in supramolecular chemistry between a cage-like, synthetic host molecule (H) and its guest molecule (G) encapsulated in the cavity.^69,70 Similar to CIU of protein–ligand complexes, CID of H–G complexes obtains survival yield of the complex as a function of collisional activation time and measures the selectivity and affinity of the noncovalent bonds.^71–73 The gas-phase dissociation kinetics of H–G complexes allows for studying artificial enzyme and self-assembly models that mimic those operating in nature, while facilitating a better understanding of the intricate characteristics of the relevant noncovalent interactions in a simpler and more controlled environment.^69,70 This gas-phase approach also provides more accurate measurements of intrinsic binding energies than the conventional condensed-phase counterpart, as it eliminates interferences from solvation and counterions present in the solution.^69,70 Overall, the use of CID for the selective dissociation of noncovalent interactions in protein–ligand and H–G complexes highlights the tunability and versatility of collisional activation.

2.2. Ozone-induced dissociation

OzID involves introducing gaseous ozone into various parts of commercial MS instruments, such as ion sources,^74,75 ion traps,^76,77 or ion mobility cells.⁷⁸ External ozone generators produce ozone for injection into MS instruments via a gas inlet system, which requires relatively simple hardware modifications.^7,24 The OzID mechanism proceeds through two competitive oxidative cleavage pathways.⁷⁹ The first is the concerted cycloaddition of an ozone molecule to a C–C double bond to generate an unstable ozonide, which spontaneously dissociates into diagnostic Criegee and aldehyde ions 16 Da apart:⁸⁰

The second, non-classical DeMore mechanism is a biradical stepwise addition of ozone to the C–C double bond that results in a diagnostic ketone and epoxide ([M + 16]⁺) ions.⁸¹

Identifying these diagnostic fragment ions reveals the locations of C–C double bonds in lipids, providing insights into the relationship between unsaturation levels and various physiological conditions, such as cell death^82–84 and diseases.^85–88 In this review, we discuss the applications of OzID, highlighting uses in MALDI imaging mass spectrometry and shotgun lipidomics.

Imaging mass spectrometry allows for the spatial mapping of analytes sampled directly from tissue surfaces.^89,90 Due to the lack of orthogonal separation techniques (e.g., gas or liquid chromatography) compatible with pixel-by-pixel, millisecond timescale acquisition, it can be challenging to identify analyte isomers without MS/MS.⁹¹ OzID has been performed on unsaturated glycerophospholipids from biological tissues,²⁵ such as human prostate⁹² and mouse brain,⁹³ to reveal the spatial distributions of olefinic C–C double bond positional isomers. Sequential CID (MS²) and OzID (MS³) provides additional information on sn-positional isomers of glycerophospholipids by producing structurally specific fragment ions containing sn-1 and sn-2 fatty acyl chains.^{25,47,78,94,95} For example, CID/OzID of lipids sampled from rat brain tissue provides for the identification of four phosphatidylcholine (PC) 34:1 isomers, PC 16:0/18:1(n-7), PC 16:0/18:1(n-9), PC 18:1(n-7)/16:0, and PC 18:1(n-9)/16:0, with the varying fractional distributions across different subsections of the tissue.⁷⁸ The use of sequential CID/OzID experiments in imaging mass spectrometry highlights the versatility of OzID in enabling high-throughput lipid isomer identification with spatial context in tissues.

OzID can also be coupled with chromatographic approaches⁹⁶ or integrated into shotgun lipidomics²⁶ for data-independent analysis (DIA) of the lipidome extracted from various samples, including blood plasma and cell cultures. In this context, the goal is primarily to resolve C–C double bond isomers of fatty acids that are often obscured in sum composition assignments in traditional shotgun workflows. For example, the analysis of the human plasma lipidome using combined DIA and OzID reveals different abundances of double bond positions in different lipid classes. Triacylglycerols (TGs) containing 50–52 carbon atoms have a high abundance of n-9, n-7, and n-6 double bonds, while phosphatidylcholines with 34–38 carbon atoms show a relatively low abundance of these double bonds. Additionally, n-4 double bonds are exclusively found in sphingomyelins.²⁶ The use of OzID in both imaging mass spectrometry and shotgun lipidomics highlights the ability of OzID to accurately identify C–C double bond locations in alkenes, providing additional molecular specificity that can be useful for understanding diseases.

2.3. Ion/neutral radical reactions

Ion/neutral radical reactions often involve exothermic charge transfers via the movement of electrons from neutral radicals with a low ionization potential to cations with a high electron affinity (A⁺ + B˙ → A˙ + B⁺).^97–102 Various free radical molecules (e.g., allyl,¹⁰¹ organic peroxy (, R = organic functional group),¹⁰² hydrofluorocarbon,⁹⁷ methyl (˙CH₃), and nitric oxide (˙NO)) have been used for ion/neutral radical reactions. Free radical molecules can be generated in a laboratory through a variety of methods, including electrical discharges,^97,98 laser photolysis,¹⁰³ and pyrolysis,^99,101,104 though it is difficult to produce a sufficient amount with high purity due to their high reactivities.^100,101,105 The short lifespan of free radicals (i.e., a few microseconds¹⁰⁶) limits the efficiency of ion/neutral radical reactions, which can be improved by extending the reaction time with a continual supply of the radical reactants. For example, flow tube reactors with integrated ion and radical sources^{97,101,102,107} and quadrupole ion traps¹⁰⁸ can serve as reaction vessels with extended ion/neutral radical reaction times (i.e., a few seconds and a few minutes, respectively).

Ion/neutral radical reactions performed under atmospheric conditions in mass spectrometry experiments are commonly used in environmental science for detecting air pollutants arising from various sources.¹⁰⁹ Chemical ionization mass spectrometry is often used to indirectly measure the concentration of pollutants in the air.¹⁰⁹ Reagent gases, such as methane or ammonia, are ionized through electron ionization to produce cations, which can then react with neutral radicals via protonation of the analyte¹¹⁰ or adduct formation between the two reactants.¹¹¹ For example, chemical ionization mass spectrometry provides a laboratory setting that enables the mechanistic study of the atmospheric autoxidation of α-pinene emitted from coniferous trees by ionizing the alkyl radical intermediates through NH₄⁺ adduction. This terpenoid generates a series of peroxy radical products (e.g., O,O–C₁₀H₁₅(O₂)_xO₂ with x = 0, 1, 2, 3) via ozonolysis in a flow tube, which then interacts with the NH₄⁺ in a reaction chamber to form ammonium adducts that can be detected using MS.¹¹² The quantitative measurements of these radicals are critical to understanding the self- and cross-reactions of intermediates that lead to the formation of secondary organic aerosols in the atmosphere,¹¹² which can cause health issues in humans, including respiratory diseases.^113,114

These types of barrier-free and spontaneous ion/neutral radical reactions under atmospheric conditions^100,101,105 do not occur in an interstellar medium, which consists of very low matter density and temperatures (<200 K).¹¹⁵ In this extreme environment, the reactant ions exist in their ground state and require excitation to certain quantum levels through cosmic radiation in order for ion/neutral radical reactions to proceed.^116–118 This interstellar chemistry produces a variety of chemicals that are considered the building blocks of earthbound molecules.⁴² Reactions between simple particles abundant in an interstellar medium, such as calcium ions (Ca⁺)¹¹⁹ and nitric oxide radicals (˙NO),¹²⁰ can be studied in laboratory environments that mimic astronomical environments to understand the formation mechanisms of extraterrestrial chemicals. For example, sympathetic cooling of ˙NO radicals using laser-cooled Ca⁺ in an ion trap^{117,118,121,122} brings the temperature of the mixture to approximately 180 K. The reaction between Ca⁺ and ˙NO produces CaO⁺ (known as lime¹²³ when in neutral form) and N only when Ca⁺ is repumped from the ground state (S_1/2) to the excited state (P_1/2) through laser irradiation (Fig. 4).¹⁰⁵ This application demonstrates the impact of quantum effects on collisions between cold ions and neutral radicals, which could allow for the study of additional interstellar ion/neutral radical reactions under laboratory conditions.^105,124

Fig. 4 Comparison of reactant and product energies for the Ca⁺ + NO system. The exothermicity of the reaction depends on the quantum state of Ca⁺. Excited states of Ca⁺ can be populated by excitation on the two cooling laser transitions at 397 and 866 nm. To achieve cooling, the 397 nm laser is red-detuned from resonance by an amount Δ. Only Ca⁺ in the P_1/2 state is energetically allowed to react. The charge exchange product channel barrier is overcome by the thermal energy of neutral NO at 180 K. (Energies not to scale.) Adapted/reproduced from Physical Review A, 2018, 98, 032702 (ref. 105) with permission from the American Physical Society, Copyright 2018.

2.4. Hydrogen/deuterium exchange

Gas-phase HDX-MS (gHDX-MS) involves the exchange of labile hydrogen atoms of amides, amines, carboxylic acids, hydroxyls, and thiols with deuterium via reactions with deuterated reagents.³³

[M − H^L + nH]ⁿ⁺ + RD → [M − D + nH]ⁿ⁺ + RH^L (H^L: labile hydrogen) (4)

The difference in gas-phase proton affinities between analytes and deuterated reagents determines the rate of HDX and the extent of deuterium incorporation.^125–127 gHDX-MS often requires relatively simple hardware modifications and can be achieved by flowing a carrier gas, such as N₂ or He, through deuterated solvents (e.g., D₂O, CD₃OD, CD₃COOD, ND₃) prior to introduction to the instrument via a gas inlet system. Deuterated vapors are then injected via leak valves into ion funnel regions,¹²⁸ electrodynamic ion traps,^129–131 FT-ICR traps,^132,133 ion mobility cells,³⁶ and ion guides,^35,134 where gHDX reactions occur. In this review, we highlight two gHDX applications: a protein structural biology approach and MALDI imaging mass spectrometry for lipid spatial mapping.

Accurately assessing gas-phase protein conformation is crucial for mechanistic understanding of the charge-state-dependent ESI processes¹³⁵ and protein gas-phase folding dynamics.^136–142 Incorporating deuterium into the labile hydrogen atoms of protein ions through gHDX provides insights into gas-phase surface accessibility and 3-D conformation.¹³¹ A recent study of ubiquitin ions generated via ESI from native and denaturing solutions shows the sensitivity of gHDX in detecting local conformational differences.¹⁴³ In this study, the 6+ charge state of ubiquitin generated from an aqueous solution and exposed to gaseous ND₃ shows a significant decrease in deuterium uptake in specific local secondary structures (e.g., β2 and β3 strands and α- and 3.5-helices) compared to the rest of the protein, indicating a tightly folded conformation. By contrast, alcohol-denatured 11+ ubiquitin exhibits extensive deuteration across the entire protein, indicating a large extent of unfolding.¹⁴³ These results demonstrate the utility of gHDX-MS in detecting gas-phase protein unfolding at varying charge states.

Accurately mapping lipid spatial distributions in MALDI imaging mass spectrometry requires differentiating isobaric compounds (i.e., compounds of the same nominal, but different fractional, m/z value), which is particularly challenging at low mass resolving power.⁹¹ However, replacing labile hydrogen atoms with deuterium after ionization can alter the chemical profile of isobaric lipid ions and provide structural information specific to different lipid classes. The MALDI plume, where active chemical reactions occur within the expanding gas cloud of laser-ablated ions and neutral molecules, provides an excellent environment for gHDX.¹⁴⁴ For example, an ion mapped in mouse brain tissue with a nominal mass of 838 Da is likely comprised of multiple isobaric lipids, including phosphatidylcholines, phosphatidylethanolamines, and glycolipids, each having a unique number of labile hydrogens (i.e., one, three, and seven, respectively). An online database search based on the isotopic distribution alone cannot differentiate these isobaric lipids.¹²⁸ However, the incorporation of seven deuterium atoms into this analyte via in-plume gHDX identifies the lipid as a potassium-cationized glycolipid, [GalCer 22:0 (OH)/d18:1 + K]⁺ (<5 ppm). The seven labile hydrogens correspond to three ceramide and four galactosyl hydroxyl groups, demonstrating improved analyte identification using gHDX in MALDI imaging mass spectrometry without the need for MS/MS.¹²⁸

3. Applications of ion/electron reactions

Ion/electron reactions occur between analyte ions and energetic electrons, forming radical product ions that undergo radical-directed dissociation.^9,145 The energy regime of electron irradiation and the mechanism of odd-electron radical formation determine the types of electron-based dissociation (ExD) methods, though the energy ranges discussed below are not clear-cut boundaries. Electron-capture dissociation (ECD) is widely used for the sequencing and analysis of noncovalent interactions and labile PTMs of multiply charged peptide and protein cations by capturing low-energy (<2 eV) electrons.^146,147 Irradiation with electron of energies 3 to 13 eV, a process known as hot ECD (HECD),^9,145–147 is an extension of ECD that obtains additional information on amino acid stereochemistry and enhances the dissociation efficiency.^146,148 However, the ECD of peptides with low charge states is generally less efficient due to the small electron capture cross-section, and singly charged cations cannot produce detectable fragment ions due to neutralization following the electron capture.¹⁴⁹ Singly or doubly charged peptide anions can also capture electrons with 4 to 7 eV and dissociate in a process known as negative-ion ECD (niECD).¹⁵⁰ Electron-induced dissociation (EID) is an alternative to ECD, which involves bombarding analyte cations with high-energy (>10 eV) electrons, which results in excitation instead of capture.^149,151,152 Multiple terms are used to describe ion activation in this energy regime, including electron-excitation dissociation (EED)¹⁵² and electron-ionization dissociation (EIoD).¹⁴⁹ The application of electrons with kinetic energies greater than 10 eV can be extended to the analyses of multiply charged analyte anions, referred to as electron detachment dissociation (EDD).^9,145

ExD instruments enable facile control of electron energies, facilitating a smooth transition between ExD regimes based on analytical requirements. All ExD experiments require sufficient spatial overlap between the analyte ions and energetic electrons, typically using one of three distinctive instrument configurations with magnetic fields to confine both reactants. In one setup, a focused electron beam generated from an external cathode irradiates ions trapped within the electric and magnetic fields of a Fourier-transform ion cyclotron resonance (FT-ICR) instrument.^153,154 In other instruments, a focused ion beam can be directed through a dense electron cloud confined within an electromagnetostatic (EMS) cell,^{4,9,145,155–157} or the ion and electron beams can flow through one another.¹⁵⁸

3.1. Electron-capture dissociation

The initial step in ECD of peptides and proteins involves the formation of charge-reduced radical cations following the capture of low-energy electrons. These radical cations undergo radical-directed dissociation and produce c-/z˙-type fragment ions through non-ergodic N-Cα backbone cleavages,^9,147 as well as b-/y-type fragment ions generated via vibrational excitation:¹⁵⁹

[M + nH]ⁿ⁺ + slow e⁻ → [M + nH]˙⁽ⁿ⁻¹⁾⁺* → fragmentation (5)

However, the thermodynamically labile nature of N–Cα bonds in the aminoketyl radical intermediates⁹ directly contributes to rapid dissociation (i.e., within microseconds)¹⁶⁰ and the preservation of labile noncovalent interactions^161,162 and PTMs^163,164 without necessarily invoking non-ergodicity.^165–167 Since the slow heating of CID does not maintain PTMs and labile bonds, ECD is commonly used as an alternative to maintain and localize these interactions and modifications to specific amino acid residues in the peptide sequence.¹⁶⁸ For example, ECD can locate hydrogen bonds within endocrine hormone islet amyloid polypeptide (hIAPP) oligomers through the detection of noncovalent complexes composed of hIAPP units and c-/z˙-type fragment ions from the neighboring unit.¹⁶⁹ Fragment ions [M + c_28/29]⁵⁺ and [M + z₃]³⁺ from 7+ dimers and [2M + c₃₄]⁵⁺ and [2M + z₅]⁴⁺ from 8+ trimers indicate the protection of C-termini from ECD, localizing the aggregate binding site at S28/S29 in one monomer and N35 in the next, resulting in staggered orientation of the oligomerization.¹⁶⁹ While a clear consensus on the mechanism of cellular aggregation has not been reached,¹⁷⁰ this result contributes to a better understanding of how these oligomers aggregate in amyloid diseases, and can potentially inform targeted drug development.¹⁶⁹ Another noncovalent interaction ECD can explore is a salt bridge network within a phosphopeptide.¹⁷¹ In this application, ECD of VVGARRS(pS)WRVVSSI (pS indicates phosphorylation at S8) reveals strong salt bridge formations between pS8 and R5 or R10 within the binding motif RRS(pS)WR.¹⁷¹ The 3D conformation modelled based on these salt bridges¹⁷¹ aligns well with crystal structures,¹⁷² demonstrating ECD as an effective tool to localize noncovalent binding and PTMs simultaneously.

Localization of noncovalent binding sites in protein multimers using ECD is similar to that in peptide oligomers. ECD of α-synuclein proteins, which undergo neurotoxic aggregation in neurodegenerative diseases, results in extensive backbone cleavages across the N-terminal halves of the α-synuclein dimers. However, the C-terminal halves remain intact, indicating the presence of noncovalent binding sites within the intact regions that block ECD from occurring.¹⁶² Comparable ECD profiles obtained from wildtype and mutant α-synuclein dimers suggest the presence of natural, non-toxic oligomers in the brain.¹⁶² Similar to peptide phosphorylation, ECD can also be used to localize sites of protein glycosylation, such as in glycoforms of α-synuclein with four modification sites (T72, T75, T81, and S87) that prevent toxic aggregation (Fig. 5).¹⁶⁴

Fig. 5 (a) Schematic, tertiary structure, and sequence of the α-synuclein monomer, showing the N-terminal amphipathic (blue), hydrophobic nonamyloid β-component of plaque (NAC, yellow), and acidic unstructured C-terminal (red) domains. (b) The structure and position of the single O-GlcNAc modification at T72 (green), T75 (blue), T81 (purple), and S87 (red). Adapted/reproduced from Analytical Chemistry, 2023, 95, 18039–18045 (ref. 164) with permission from the American Chemical Society, Copyright 2023.

HECD and niECD complement conventional ECD by enhancing structural information and broadening sample compatibility for low-charge-state anions, respectively. The higher electron energies delivered via HECD produce additional secondary d-/w-type fragment ions through endothermic cleavage of primary c-/z˙-type fragment ions,¹⁴⁶ while still retaining PTMs, such as glycosylation^148,173 and methylation.¹⁷⁴ These secondary fragment ions enable the differentiation of amino acid isomers in peptides and proteins that are not identified by traditional ECD.¹⁷⁵ For example, z˙-type fragment ions containing valine and norvaline lose ˙CH₂CH₃ (29 Da) and ˙CH₃ (15 Da), respectively, to generate diagnostic w-type counterparts. Accurately identifying these isomers is critical in biologics, as the bacteria used for recombinant protein production naturally synthesize norvaline but can mis-incorporate it into proteins instead of leucine, causing toxicity of the therapeutic proteins.¹⁷⁶ During niECD, peptide and protein anions with low charge states and zwitterionic structures¹⁷⁷ capture 4–7 eV electrons and form charge-increased radical anions that undergo radical-directed dissociation to produce c-/z˙-type fragment ions while preserving PTMs, similar to ECD and HECD.¹⁵⁰ niECD is particularly useful for PTM analyses of acidic peptides, such as sulfopeptides¹⁷⁸ and phosphopeptides,¹⁷⁹ due to their greater ionization efficiency in negative mode. Singly charged anions typically show greater electron capture efficiency than multiply charged counterparts due to decreased coulombic repulsion of the energetic electrons by the analyte anions,^177,178 making MALDI an effective ionization source for niECD. Indeed, MALDI-generated, singly deprotonated hirudin sulfopeptide (∼m/z 1500) and β-casein phosphopeptide (∼m/z 2000) subjected to niECD result in near complete sequence coverage and accurate localization of PTMs.¹⁷⁹ Furthermore, similar niECD profiles of synthetic peptides with different salt bridge formations, ionized through MALDI or ESI, suggest that conformationally comparable peptide structures can be generated regardless of the MALDI matrix pK_a and the ion source, providing valuable insights into the relative ionization mechanisms.¹⁷⁹

3.2. Electron-induced dissociation

A generally accepted dissociation mechanism for EID of analyte cations involves the formation of an oxidized radical cation [M + nH]˙⁽ⁿ⁺¹⁾⁺ from the precursor ion [M + nH]ⁿ⁺ via ionization (i.e., an increase in the total positive charge of the analyte from +1 to +2). A slow, low-energy electron is ejected during this bombardment process, which is then recaptured by the [M + nH]˙⁽ⁿ⁺¹⁾⁺ to form an electronically excited [M + nH]ⁿ⁺* ion type that then undergoes radical-directed dissociation.^{9,145,149,152}

[M + nH]ⁿ⁺ + fast e⁻ → [M + nH]˙⁽ⁿ⁺¹⁾⁺ + fast e⁻ + slow e⁻ → [M + nH]ⁿ⁺* → fragmentation (6)

Direct electronic excitation of the oxidized radical cation [M + nH]˙⁽ⁿ⁺¹⁾⁺* and subsequent dissociation has also been proposed.^149,180 EID has been used for the structural elucidation of peptides and proteins of varying charge states^{149,151,152,181,182} as well as singly-charge glycans,¹⁸³ lipids,^184–188 metabolites,^189,190 and other small molecules.^191–193 In this review, we limit the discussion of EID to the study of peptides, proteins, and lipids.

Side-chain loss upon EID of singly or doubly protonated peptides can be abundant for specific amino acid residues, which improves sequencing while preserving labile PTMs, such as glycosylation^181,182 and sulfation,¹⁹⁴ similar to HECD. However, unlike HECD,¹⁸² the dissociation efficiency for EID remains unaffected by charge states due to the electronic excitation that occurs through the interactions between analyte ions and high-energy electrons.¹⁴⁹ EID also produces more abundant internal fragment ions from large, multiply protonated protein ions compared to (H)ECD, which can enhance structural information.^195,196 For example, the efficient internal fragmentation of lysozyme, ribonuclease A, and α-lactalbumin through EID provides a three-fold increase in sequence coverage and localizes more disulfide bonds compared to ECD.¹⁹⁷ Overall, EID studies of peptides and proteins highlight the enhanced and charge-independent sequence coverage that can be obtained with this approach.

EID has also been incorporated into an imaging mass spectrometry workflow to explore the spatial distribution of singly charged glycerophospholipids generated directly from tissue via MALDI, providing information on lipid classes, FA chain lengths, and C–C double bond sites.^198,199 Similar to the sequential CID/OzID experiments discussed above, prior CID on glycerophospholipids generates dioxolane intermediates that provide sn-positional information upon subsequent EID.²⁰⁰ Unlike OzID, which produced specific and selective cleavage at double bonds, EID generates extensive cleavage across the entire length of fatty acyl chains, with “V”-shaped dips in allylic and vinyl fragment ion intensities around C–C double bonds. This sequential CID/EID method enables comprehensive structural characterization of singly protonated PC 34:1 from five different regions of rat brain tissue, which shows abundances of PC 16:0/18:1 relative to PC 18:1/16:0 ranging from 57.76 ± 2.23% in the granular layer to 75.77 ± 1.94% in the corpus callosum (Fig. 6).²⁰⁰ These results highlight the facile integration of EID into imaging mass spectrometry, enabling quantitative mapping of glycerophospholipids at sn-isomeric levels without prior separation.²⁰⁰

Fig. 6 (a) Representative CID/EID spectrum of m/z 782.56 (±1 Da isolation width) from the cortex of rat brain tissue. (b) H&E stain and ion image for m/z 782.5695 ± 0.01 allow for visualization of sodiated PC 34:1 (3.21 ppm) in the tissue. The image is normalized to TIC and acquired using a 75 µm spatial resolution. (c) Calibration curve for the relative quantification of the sn-isomers is constructed using the summed intensities of the three PC 16:0/18:1 diagnostic ions divided by the summed intensities of all diagnostic ions for both isomers (n = 10). (d) Relative percentages of the PC 16:0/18:1 and PC 18:1/16:1 sn-isomers vary in different regions of the rat brain tissue (n = 5). Adapted/reproduced from Analytical Chemistry, 2023, 95, 15707–15715 (ref. 200) with permission from the American Chemical Society, Copyright 2023.

3.3. Electron detachment dissociation

While fewer mechanistic studies have been made for EDD than other ExD methods, a generally accepted mechanism involves bombarding negative ions with high-energy electrons, resulting in subsequent electron detachment.²⁰¹ Charge-reduced radical ions [M − nH]˙⁽ⁿ⁻¹⁾⁻ generated via the electron detachment undergo radical-directed dissociation:²⁰¹

[M − nH]ⁿ⁻ + fast e⁻ → [M − nH]˙⁽ⁿ⁻¹⁾⁻* + 2e⁻ → fragmentation (7)

The charge reduction process requires multiply charged analyte anions to avoid neutralization. A variety of sample types, including glycans,^202,203 nucleotides,^204,205 and peptides,^206,207 can undergo EDD for structural determinations. Here, we discuss recent applications of EDD to differentiate peptide and glycan isomers.

EDD of multiply charged peptide anions leads to cleavages at N–Cα and Cα–C bonds, resulting in the production of a˙-/c-/x-/z˙-type fragment ions, similar to ECD.¹⁴⁵ In a recent study, EDD of [M − 3H]³⁻ of antibacterial 21-mer lasso peptides from E. coli (Microcin J25) produces diagnostic bi-peptidic fragment ions that signify the lasso topology, such as ²⁰⁷ c₈⁻ represents the intact N-terminal macrolactam ring, and × denotes covalent or noncovalent attachment with z˙- or y-type fragment ions dissociated from the C-terminal tail sterically entangled with the ring.²⁰⁷ By contrast, CID of the same topoisomeric anions predominantly results in side-chain loss from T15 residues and fails to provide structurally meaningful information, highlighting the sensitivity of EDD to the complex 3D topology of peptides.²⁰⁷

EDD also enables structural differentiation of glycosaminoglycans isomers, which is typically challenging due to the subtle variation in stereochemistry. For example, EDD of epimeric heparan sulfate tetrasaccharide anions possessing different stereochemistry at C5 of ring-3 produces diagnostic fragment ions that distinguish glucuronic acid (GlcA) from iduronic acid (IdoA) substitution.²⁰³ Cleavage at the glycosidic bond between ring-3 and ring-4 (fragment ion ), in combination with loss of CO₂ , indicates the presence of a GlcA epimer at the ring-3 position, which is lost with the substitution of IdoA for GlcA.²⁰³ The detailed glycosaminoglycans stereochemical information provided by EDD is critical to understanding their binding specificities toward on glycosaminoglycans-binding proteins.

4. Applications of ion/photon reactions

Ion/photon interactions occur when analyte ions absorb photon energies of specific wavelengths.² Here, we highlight ultraviolet photodissociation (UVPD)^11–13 and infrared multiphoton dissociation (IRMPD),²⁰⁸ in which nonselective or selective absorbance of photons of specific wavelengths by analyte ions can result in fragmentation once the ion internal energy exceeds a dissociation threshold.¹¹

[M + nH]ⁿ⁺ + nhν → [M + nH]ⁿ⁺* → fragmentation (8)

F₂ excimer lasers (157 nm, 7.9 eV per photon), ArF excimer lasers (193 nm, 6.4 eV per photon), and Nd:YAG lasers operated using the 5th (213 nm, 5.8 eV per photon), 4th (266 nm, 4.7 eV per photon), and 3rd (355 nm, 3.5 eV per photon) harmonics are the most commonly employed wavelengths for UVPD, while CO₂ lasers (10.6 µm, 0.12 eV per photon) are often used for IRMPD. However, various chromophores and diverse intrinsic electron densities around chromophores dictate the absorption selectivity and efficiencies, and thus the division based on wavelength alone is not clearcut.¹¹ These photodissociation experiments in mass spectrometry require sufficient spatial overlap between the analyte ions and laser beams for efficient dissociation, similar to the ExD experiments discussed above. This can allow for prevalent secondary (i.e., consecutive) dissociation of first-generation fragment ions, as primary fragment ions remain in the laser path and continue to undergo irradiation. Alternatively, some instruments can employ “protective” mass-selective resonant excitation of primary fragment ions to displace them from the laser path and minimize consecutive fragmentation.^209–211 RF-ion traps,^212,213 RF-ion guides,²¹⁴ and FT-ICR cells^215,216 are commonly used to achieve optimal photoabsorption cross-sections by fine-tuning the ion trajectories, while laser beams are introduced into instrument vacuum system through windows transmitting light at the specified wavelength.^11,21,208

4.1. Nonselective UV photodissociation

UVPD of peptide and protein ions at 157, 193, or 213 nm generally involves the selective absorption of single energetic photons via chromophores that are evenly distributed along the backbones, such as π orbitals of amido groups leading to π-to-π* excitation or nonbonding electrons of carbonyl oxygens resulting in n-to-π* excitation.^12,13 This high-energy deposition exceeds the energy of peptide bonds (i.e., 3–4 eV) and leads to nonselective dissociation directly from the excited π* electronic states, causing extensive backbone cleavage into a-/b-/c-/x˙-/y-/z˙-type fragment ions.^12,13,217 The non-ergodic cleavages at N–Cα and Cα–C bonds enable the study of preserved noncovalent interactions^218–220 and labile PTMs, including acetylation,²²¹ methylation,^221,222 glycosylation,²²³ phosphorylation,²²⁴ and sulfation.²²⁵ Our discussion centers on 193 nm UVPD, as it is the most widely adopted wavelength.

The nonselective fragmentation channels and charge-independent dissociation efficiency make 193 nm UVPD particularly useful for low-charge-state peptides and proteins.²²⁶ By comparison, CID,²²⁷ higher-energy collisional dissociation (HCD, <100 eV),^226,228,229 ECD,¹⁴⁷ and 213 nm UVPD²³⁰ all exhibit strong charge state dependency in fragmentation efficiency, though 213 nm UVPD still provides greater sequence coverage than CID and HCD.²³¹ For example, HCD of ESI-generated 5+ ubiquitin (8.6 kDa), 8+ Staph nuclease (16.1 kDa), and 7+ calmodulin (16.6 kDa) shows limited sequence coverages of 84%, 53%, and 43%, respectively, due to the reduced proton mobility responsible for charge-directed fragmentation.²³² However, 193 nm UVPD achieves significant improvements in sequence coverage, up to 98%, 79%, and 66% for each protein, respectively.²³² HCD efficiency is even more limited with singly charged proteins commonly produced by MALDI,^226,233 as shown by 8% sequence coverage for the 1+ charge state of ubiquitin, which is significantly enhanced to 87% by using 193 nm UVPD.²³⁴ These results demonstrate that 193 nm UVPD provides better sequence coverage than conventional HCD at low charge states, and suggest that the MALDI applications, including MALDI imaging mass spectrometry, can potentially adopt UVPD for top-down protein identification of singly charged proteins directly from tissue.^233,234

Noncovalent interaction sites in peptide²³⁵ and protein complexes^236–238 can be explored using 193 nm UVPD, similar to topological mapping studies enabled by ECD. For example, paratopes on engineered biotherapeutics, such as in a complex of a Gnb nanobody and its green fluorescent protein (GFP) antigen, can be deduced from a significant suppression in UVPD fragmentation adjacent to the protein–protein interfaces (Fig. 7).²³⁹ Additionally, weighted average charges acquired from a-/x-type fragment ions²⁴⁰ aid in determining sites of inter-subunit salt-bridge formation at Gnb (R36)–GFP (E142) and Gnb (R58)–GFP (E172/D173), which is supported by crystal structures.²³⁹ This application highlights the ability of 193 nm UVPD to locate different types of noncovalent interactions and map protein quaternary structures.²³⁹

Fig. 7 Suppression and enhancement of backbone cleavage sites based on abundances of UVPD fragment ions induced for Gnb by antigen binding. Heat plots display significant differences (p < 0.05, n = 5) for the abundances of each UVPD fragment type between the free and bound states. Blue and red indicate suppression and enhancement of fragment abundances, respectively, for the nanobody upon complexation. Positions that display no significant change are shown in grey. Color maps highlighting interface residues and CDRs are also included. ΔUVPD values correspond to the fragment abundance per residue for the bound state minus the fragment abundance per residue for the free state. Adapted/reproduced from Chemical Science, 2022, 13, 6610–6618 (ref. 239) with permission from the Royal Society of Chemistry, Copyright 2022 The Authors.

4.2. Selective UV photodissociation

UV photodissociation at longer wavelengths, 266 and 355 nm,²⁴¹ is not as widely utilized in mass spectrometry. However, the high absorption selectivity of these wavelengths by certain chemical bonds provides a unique utility for the application of peptide and protein ions. For example, selective 266 nm UVPD involves modifying peptide ions prior to ionization by introducing carbon–iodine (C–I) bonds via derivatization with chloramine-T^242,243 or iodobenzoic acids.²⁴⁴ Irradiation of these C–I bonds with 266 nm light results in selective homolytic cleavage and generates stable free radical ions.^242,243 Subsequent CID of the radical cation initiates radical-directed dissociation sensitive to the stereochemistry of neighboring amino acid residues, including Asp, Ser, His, and Glu, via structure-dependent radical migration pathways.²⁴⁵ For example, L-aspartic acid and L-isoaspartic acid in the iodinated synthetic peptide GISEVRSDG are differentiated by assessing the relative abundances of a₈⁺ fragment ions and CO₂ loss.^244,245 Sequential 266 nm UVPD/CID can also provide information on protein topology.²⁴² Another example of selective UV photoabsorption utilizes a chromogenic diazirine-tagged peptide^246,247 for cross-linking mass spectrometry, where distance restraints imposed by cross-linkers provide low-resolution 3-D structural information for peptides and proteins.²⁴⁸ The diazirine group selectively absorbs a 355 nm photon and converts to a reactive carbene intermediate, which then forms a covalent bond with the immediately neighboring amine or carboxylic acid group, signified by the loss of the N=N moiety.^249,250 This zero-length chromogenic cross-linker does not introduce any physical distance between the covalently linked molecules, allowing it to probe protein–ligand or protein–protein interfaces via carbene insertion into X–H bonds (X = C, N, or O).^249,250

Iodinated lipid ions can also form radical cations by selectively absorbing 266 nm photons at C–I bonds, leading to iodine atom cleavage due to UV irradiation. Subsequent radical-directed dissociation facilitated by CID results in allylic cleavage that unambiguously identifies C–C double bond sites^251–255 Alternatively, absorption of 193 and 213 nm light by lipid ions with isolated, non-conjugated C–C double bonds occurs by the entire molecule acting as a chromophore, due to high-energy σ → σ* and n → π* transitions, similar to the universal absorption by peptide and protein backbones.^256–260 Following photoexcitation, the molecule redistributes energy internally and fragments preferentially at labile sites, especially near double bonds, producing diagnostic allylic and vinylic fragment ions that localize the unsaturation sites.^{253,256–262} Sequential HCD/UVPD can be used to identify sn-positional isomers of glycerophospholipid²⁶¹ and cardiolipin (CL) ions²⁶³ by generating dioxolane intermediates through thermal dissociation enabled by HCD, and then subsequent cross-ring cleavages via selective UVPD, similar to the CID/OzID approach discussed above.^261,263 For example, HCD/UVPD enables the identification of the tumor-specific and unusual prevalences of phosphatidic acid (PA) 18:2/18:1 (76.5%) over PA 18:1/18:2 (23.5%) and PC 18:1/18:2 (70.2%) over PC 18:2/18:1 (29.8%) in CL 72:6 in a total lipid extract from papillary thyroid carcinoma tissue.²⁶³ In another study, HCD/UVPD can be used to identify ester bond locations in isomeric palmitic acid esters of hydroxystearic acids by creating a new C–C double bond specifically at the ester site during the HCD process, further demonstrating selective UVPD for lipid identification.²⁶⁴

4.3. IR photodissociation

Protein molecules have IR-active functional groups that enable the broad absorption of 10.6 µm CO₂ laser light, including C–C, C–N, C–O, and P–O stretching and O–H bending vibrational modes.²⁶⁵ However, the degree of energy deposition is highly tunable due to the multi-step absorption process of low-energy photons, resulting in slow heating via intramolecular vibrational energy redistribution.²⁶⁶ This process contributes to the selective fragmentation of labile PTMs, amide bonds, and noncovalent bonds via low-critical-energy pathways.²⁰⁸ For example, step-wise selective dissociation of membrane protein–detergent complex ions is demonstrated using a low-power CO₂ laser beam (typically <10 W).^265,267 The detergents stabilize the hydrophobic proteins in solution prior to ionization, but should be removed in the gas phase prior to mass analysis of the intact proteins, as the bound detergents complicate spectral interpretation. A low amount of laser power and irradiation time liberates trimeric ammonia channel AmtB protein ions from the protein–detergent complex ions, while additional energy deposition dissociates trimers into monomer subunits (Fig. 8).²⁶⁵ Collisional activation methods can also remove detergents^268–270 though with more limited and less efficient energy deposition that leads to incomplete removal of detergents and compromised protein sequence coverage.²⁶⁵ IR irradiation allows for more precise and effective tuning of the energy deposition, decreasing the chemical interferences and enhancing protein sequence coverage.^265,267 For example, step-wise IRMPD successfully liberated a class A G protein-coupled receptor (GPCR), beta-1-adrenergic receptor (β1AR), from the detergent and achieved 28% sequence coverage, while an analogous experiment performed using HCD only achieved 17% sequence coverage.²⁶⁵

Fig. 8 The effect of modulating laser output power and irradiation time for AmtB solubilized in DDM detergent. At low laser output power and short irradiation times, the peaks corresponding to AmtB trimer appear at low abundance due to signal dilution across many adduct peaks. With increasing irradiation time, peaks corresponding to lipid-bound trimers are observed as the protein is liberated from DDM detergent micelles. These peaks dominate the mass spectrum ∼6000 m/z. At intermediate laser output powers from short to long irradiation times, the relative abundances of peaks corresponding to trimer and monomer can be modulated. Finally, at high laser output power and short irradiation times, peaks corresponding to monomer and trimer are approximately equal in abundance, while at long irradiation times, the peaks corresponding to trimer are absent as the protein complex has expectedly dissociated into monomeric subunits (peaks <5000 m/z). Adapted/reproduced from Angewandte Chemie International Edition, 2023, 62, e202305694 (ref. 265) with permission from Wiley-VCH GmbH, Copyright 2023 The Authors.

Lipids also have universal chromophores for nonselective absorption of IR laser light and can undergo selective dissociation via the lowest energy pathway, such as the loss of fatty acyl moieties from glycerophospholipid anions via fragmentation at labile ester bonds.^271,272 This feature can be used in MS/MS imaging mass spectrometry to study the spatial distribution of glycerophospholipids from tissue at the sn-positional isomer level.²⁷³ In this study, IRMPD of singly charged PC 34:1 anions produces varying abundances of fatty acyl moieties depending on the sn-positions, revealing differential isomer expression in different rat brain regions.²⁷³ MS/MS performed at every pixel in this experiment allowed mapping the relative distribution of PC 16:0/18:1 to PC 18:1/16:0 in regions such as the granular layer (64.5 ± 2.5%) and white matter (78.1 ± 2.2%).²⁷³ Though not discussed in detail in this review, blackbody infrared radiative dissociation (BIRD) is another photodissociation technique that leverages nonselective photoabsorption of a range of blackbody IR wavelengths emitted from heated ICR cell walls.¹⁰ Selective bond dissociation via BIRD is often used to study dissociation kinetics of noncovalent host–guest binding,^274–277 similar to CID-based approaches discussed above.

4.4. Action spectroscopy

Spectroscopic analysis of gas-phase ion chemistry in mass spectrometers is known as action spectroscopy,²⁷⁸ in which light absorption is measured indirectly (i.e., by monitoring photodissociation) due to the low ion concentrations that make traditional absorption spectroscopy impractical.^279–284 The photodissociation yield can be plotted as a function of wavelength, providing information on the thermodynamic threshold for the photodissociation and electronic transition dynamics of the precursor ion.^285–291 Light absorption by gas-phase trapped ions can also be indirectly measured through sensitive fluorescence detection,^292,293 facilitating advanced applications such as action – Förster resonance energy transfer (FRET) spectroscopy for protein tertiary structure analysis²⁹⁴ and quantum computing utilizing quantum state-specific fluorescent ions to convert superposition states into classical 0/1 outcomes.²⁹⁵ A detailed discussion of action spectroscopy fundamentals and applications can be found elsewhere,^{279,281,282,284,286–290,296} as it is beyond the scope of this review.

5. Applications of ion/ion reactions

Ion/ion reactions occur when analyte and reagent ions of opposing charge interact due to coulombic attraction, with reaction rates proportional to the square of the charges.¹⁴ The coulombic interactions facilitate a variety of changes in the conformational, electrochemical, and physical properties of analyte ions via the transfer of particles (i.e., electrons, protons, and metal ions) and covalent modifications.^297,298 Ion/ion reactions can be performed outside mass spectrometers in atmospheric environments, such as with ambient bipolar dual spray, where overlapping streams of positive and negative ions enable ion/ion interactions.²⁹⁹ However, in most ion/ion reactions under reduced pressure environments inside mass spectrometers, it is crucial to bring together ions with opposite charges close enough in space to form stable orbiting electrostatic complexes.²⁹⁷ Therefore, the critical step is securely and mutually trapping these ions in an electrodynamic trap.^298,300 Many types of electrodynamic traps can facilitate ion/ion reactions via simultaneous confinement of ions of opposite charges.^298,300 Electron transfer dissociation (ETD) is the most widely used ion/ion reaction, and has been commercialized on many instrument platforms.^301–303 A broad array of gas phase ion/ion reactions, including proton transfer charge reduction,^304,305 charge inversion,³⁰⁶ metal ion transfer,³⁰⁷ and covalent modification³⁰⁸ chemistries can be enabled on commercial instruments already equipped with ETD,^301–303 or these reactions can be enabled via instrument modification or construction.^{15,298,300,309}

5.1. Electron transfer dissociation

ETD was first developed to enable ECD-like peptide and protein sequencing in electrodynamic ion traps without requiring magnetic fields and thermal electrons.³⁰⁹ ETD involves transferring electrons from singly charged radical reagent anions to multiply charged protein cations during the ion/ion reaction, which electronically excites the analytes. The odd-electron radical protein cations produced undergo cleavages at N-Cα bonds to produce c-/z˙-type fragment ions while preserving labile PTMs, similar to ECD.^{9,17,145,310,311}

[M + nH]ⁿ⁺ + A˙⁻ → [M + nH]˙⁽ⁿ⁻¹⁾⁺ + A (9)

ETD is widely used for PTM analysis as an alternative to slow-heating collisional activation methods,³¹² similar to ECD. For example, ETD of U1 small nuclear ribonucleoprotein 70 kDa (U1-70K) enables the identification of 29 PTM sites from 55 peptides, while only 19 PTM sites from 13 peptides are identified via HCD.³¹² Identifying these PTM sites is crucial for understanding protein–protein interaction sites that can lead to aggregation in diseased individuals.³¹² However, ETD is often combined with HCD (i.e., EThcD)^313,314 to enhance sequence coverage by thermally dissociating ETD fragment ions that have undergone N–Cα bond cleavage, but that are still held together by noncovalent interactions.^315–319 For example, EThcD of trypsin-digested peptides from HeLa cell lysates shows abundant c-/z˙-type fragment ions for peptide sequencing and allows for the detection of roughly 1.7-fold more ADP-ribosylation PTM sites³²⁰ than conventional ETD.³²¹ ADP-ribosylation is a growing area of research related to cell signaling and DNA damage repair processes, and identifying this PTM enhances the understanding of associated enzyme activities.³²¹ Another benefit of EThcD compared to HCD or ETD alone is the ability to cleave disulfide bonds. For example, EThcD breaks disulfide bonds in the C-terminal cyclic structures of frog skin peptides, leading to enhanced sequencing of the cyclic region, which is typically challenging to sequence when the structures remain closed as with using ETD or HCD.^322,323 EThcD also facilitates differentiation of amino acid isomers, such as leucine and isoleucine, through different secondary w-type ions generated from the primary z˙-type fragment ions and provides near-complete sequencing of the amphibian peptides.^322,323

EDD-like reactions in electrodynamic ion traps can also be achieved via negative electron transfer dissociation (NETD), where singly charged reagent radical cations abstract single electrons from multiply charged analyte anions, producing charge-reduced, odd-electron radical anion analytes that undergo radical-directed dissociation.³²⁴

[M − nH]ⁿ⁻ + B˙⁺ → [M − nH]˙⁽ⁿ⁻¹⁾⁻ + B (10)

NETD can be used to analyze various sample types, such as peptides,^324–326 proteins,³²⁷ glycans,^328,329 and nucleotides.³³⁰ For example, NETD of glycosaminoglycans produces extensive glycosidic (B/Y and C/Z) and cross-ring (A/X) fragment ions, while preserving labile sulfate modifications. The intricate combinations of these fragment ions allow for the distinction of GlcNS3S (NS: 2-N-sulfation, 3S: 3-O-sulfation) and GlcNS3S6S (6S: 6-O-sulfation) in tetrasaccharide structural isomers, which differ only by sulfation on the glucosamine (GclN) moieties at the reducing ends.³³¹ Unambiguously identifying the 3-O- and 6-O-sulfation sites is critical for understanding the biological functions of glycosaminoglycans, as these modification sites play essential roles in the anticoagulant properties of heparan sulfate by binding to antithrombin, which inhibits enzymes responsible for blood clotting.^331–333

The transfer of electrons from radical reagent ions to analyte ions does not always result in dissociation.^334–336 Still, oxidation or charge reduction resulting from electron transfer can enable charge manipulation to deconvolute crowded spectra of multiply charged protein ions or to explore charge-dependent gas-phase protein conformational changes.^337,338 For example, the gas-phase structures of ubiquitin charge conformers, formed either directly from the ESI source or via gas-phase charge manipulation of ESI-generated ions, can be examined. Specifically, the alcohol-denatured 6+ charge state of ubiquitin is compared to 6+ ubiquitin produced via non-dissociative electron transfer (i.e., charge reduction) of alcohol-denatured 8+ ubiquitin (8+ → 6+) in an ion trap.³³⁵ Incremental collisional activation significantly alters the collision cross-sections of 6+ ions produced directly from the ESI source, but not those generated via electron transfer charge reduction (Fig. 9). This result indicates that charge-reduced ions (8+ → 6+) retain the conformational memory of the original 8+ ubiquitin, which differentiates them from the 6+ ions formed directly from the ESI process.³³⁵ Conformational changes, similar to gas-phase protein folding via non-dissociative electron transfer, can initiate directional motion in molecular switches. High charge states elongate the molecule into a rod shape due to Coulomb repulsion, and a folded globule forms as the charge reduces through stepwise electron transfer, showing the mechanical control of the switch via redox chemistry.³³⁹ Gaining insights into the reversible structural changes of molecular switches in response to an external stimulus aids in designing precise control mechanisms at the molecular level for applications such as molecular electronics.³³⁹

Fig. 9 Collision cross sections for 6+ ubiquitin formed by either (a and b) electrospray ionization, or (c and d) charge reduction of ESI-generated [M + 8H]⁸⁺ by using ETD. Time axes of arrival time distributions were converted to CCS as described in ref. 340. Data shown in panels (a) and (c) were acquired using a sampling cone voltage of 40 V and a trap DC bias of either 25 (blue), 35 (orange), or 45 V (red). In panels (b) and (d), the bias was kept constant at 25 V and a sampling cone voltage of 50 (blue), 60 (orange), or 100 V (red) was applied. Colored text in panels indicates (cone voltage)/(bias voltage) for each trace. Gaussian components used in deconvolution of the lowest-energy (blue) trace in each panel are shown at the bottom at half-intensity. Values in parentheses are optimized apparent PTR/ETnoD ratios for each of these components, as described in the text. Weighted average CCS values (used to generate Fig. 2) are indicated with green arrows in panels (c) and (d). Adapted/reproduced from Journal of the American Society for Mass Spectrometry, 2017, 28, 69–76 (ref. 335) with permission from the American Chemical Society, Copyright 2017.

5.2. Proton transfer charge reduction

Proton transfer charge reduction features proton transfer from multiply protonated analyte cations to reagent anions with high gas-phase basicity via a single collision between the two reactants.^341,342

[M + nH]ⁿ⁺ + A⁻ → [M + (n – 1)H]⁽ⁿ⁻¹⁾⁺ + AH (11)

Proton transfer charge reduction is commonly employed to reduce the charge states of multiply protonated precursor or fragment ions that are present in an m/z range with significant spectral overlap among various ions of different charge states.²⁹⁸ Proton transfer charge reduction decongests complicated mass spectra and can reduce signal dilution from distribution across multiple charge states for a single analyte, which results in a improved signal-to-noise ratio^343,344 and sequence coverage^345–347 in complex mixtures.³⁴¹ For example, proton transfer charge reduction streamlines complex spectra from top-down ETD analysis of the antibody–drug conjugate, trastuzumab deruxtecan (T-DXd), enhancing fragment ion detection and resulting in a ∼10% increase in sequence coverage compared to ETD alone.³⁴⁸ The separation of overlapping fragment ions via proton transfer charge reduction also enables a more confident determination of the drug conjugation sites of topoisomerase I inhibitor payload (DXd), such as Cys214 on the light chain and Cys223, Cys229, and Cys232 on the Fd subunit of the heavy chain.³⁴⁸ Proton transfer charge reduction of complex UVPD MS/MS spectra generated from large proteins (>30 kDa) with high charges (>30+) also shows a significant increase in sequence coverage.³⁴⁹ Overall, proton transfer charge reduction demonstrates utility in simplifying MS/MS spectra of multiply charged proteins that commonly produce a large number of fragment ions with overlapping m/z values.

In addition to sequencing, proton transfer charge reduction is utilized to explore the 3D structures of proteins in the gas phase. IM-MS of product ions resulting from sequential proton transfer charge reduction of multiply protonated denatured proteins of various sizes, such as ubiquitin,¹³⁸ cytochrome c,¹³⁹ lysozyme,¹³⁷ and serum albumin,¹³⁶ reveals collisional cross-sections indicative of a gradual gas-phase folding of the protein ions as the charge states decrease. However, the collisional cross-sections derived from the proton transfer charge reduction products of protein ions initially formed under native solution conditions show overall consistent values across varying charge states, indicating compact conformations with less charge dependence.^136–139 The collisional cross-sections of low-charge proton transfer charge reduction products of denatured proteins are generally larger than those of native counterparts, suggesting the proton transfer charge reduction product protein ions retain some of their initial structure.^136–139 This trend suggests that the formation of a noncovalent bonding network differs in vacuo from in solution during protein folding,¹⁴⁰ providing insights into protein folding dynamics.

Proton transfer charge reduction can also be applied to lipid anions, such as in the charge reduction of [CL − 2H]²⁻ dianions to [CL − H]⁻ monoanions using singly charged proton-donor cations.³⁵⁰ Proton transfer charge reduction improves the signal-to-noise ratio of the dianions by concentrating signals into monoanions, alleviating the challenge of identifying CL dianions in congested m/z ranges dominated by phospholipid monoanions in the complex lipidome.³⁵⁰ This application also demonstrates that proton transfer charge reduction can be effected in either ion polarity.

5.3. Charge inversion reactions

Transferring multiple protons between reactant ions with opposite polarity can result in charge inversion of analyte ions rather than reduction.

[M + H]⁺ + [N − nH]ⁿ⁻ → [M − H]⁻ + [N − (n − 2)H]⁽ⁿ⁻²⁾⁻ (12)

[M − H]⁻ + [R + nH]ⁿ⁺ → [M + H]⁺ + [R + (n − 2)H]⁽ⁿ⁻²⁾⁺ (13)

Ion/ion charge inversion reactions can be used to access new and complementary gas-phase dissociation chemistries by altering the analyte ion type prior to CID. For example, multiply deprotonated phosphopeptides are charge inverted to multiply protonated cations by a reaction with multiply protonated polyaminoamine dendrimers.³⁵¹ The ionization of the phosphopeptides in negative ion mode allows for their efficient formation without ionization suppression, and the multiply protonated ion/ion reaction product phosphopeptide allows for the use of ECD for the localization of phosphorylation sites.³⁵¹ Multiply protonated proteins generated in positive ion mode can also be charge inverted to deprotonated anions via reaction with multiply deprotonated hyaluronic acid.³⁵² CID of the protein anion then provides for complementary dissociative channels for PTM localization.^353,354

Lipids exhibit different ionization efficiencies in positive and negative ion modes. For example, phosphatidylcholines have higher efficiency in positive mode due to the permanent positive charge embedded in the choline moiety. However, low-energy CID of protonated phosphatidylcholines generally only allows for the characterization of the lipid head group. Conversely, singly protonated phosphatidylcholines can be charge-inverted via an ion/ion reaction with 1,4-phenylenedipropionic acid (PDPA) dianions. Subsequent CID on the resulting demethylated phosphatidylcholine anion provides fragmentation indicative of the fatty acyl chains in the lipid.^355,356 The relative fractions of PC 16:0/18:1 compared to PC 18:1/16:0 in different rat brain substructures can be mapped via imaging mass spectrometry combined with this PDPA ion/ion charge inversion reactions, and range from 50.1 ± 1.9% in the granular layer to 76.5 ± 2.1% in the corpus callosum.³⁰⁶ Gas-phase ion/ion reactions can also be combined with other derivatization approaches. For example, a four-membered cyclic ether, incorporating two carbon atoms into C–C double bonds, can be derivatized by a condensed-phase Paternò–Büchi reaction at the double bond sites prior to ionization. When combined with the PDPA charge inversion reaction, subsequent CID provides information on C–C double bond positions within the glycerophospholipids.³⁵⁷ Overall, gas-phase charge inversion reactions resolve analytical challenges arising from different ionization and MS/MS efficiencies by decoupling ion formation from the ion type ultimately subjected to MS/MS. These reactions enable the formation of new ion types in the gas phase, eliminating the need for condensed-phase analyte derivatization.

5.4. Metal ion transfer reactions

A variety of metal ions (i.e., alkali,^{199,358–360} alkali earth,^361–363 and transition^363–365) can form adducts with analytes during ionization,

[M + nH]ⁿ⁺ + Met_xL_y⁻ → [M + nH + Met_xL_y]⁽ⁿ⁻¹⁾⁺ → [M + (n − y)H + xMet]⁽ⁿ⁻¹⁾⁺ + yHL (14)

Facilitating novel gas-phase dissociation chemistries upon MS/MS compared to common protonated and deprotonated ion types. Condensed-phase metal ion adduction can also be brought to the gas phase via ion/ion reactions, enabling control over the reactant ion types and their reaction kinetics.^366–368 This can be used to obtain enhanced structural information on peptide and lipid analytes.

Cation switching of peptide ions can be achieved by reacting [AuCl₂]⁻ anions with [M + 2H]²⁺ peptide cations to form the electrostatic complexes [M + 2H + AuCl₂]⁺, which lose 2HCl during subsequent low-energy CID and produce aurated peptide ions [M + Au]⁺.³⁶⁹ Subsequent CID of the mass-selected aurated peptide ion produces a protonated six-membered cyclic iminium ion via HAuNH₃ loss facilitated by a lysine residue. This gives in an [M − H − NH₃]⁺ ion that results in preferential cleavage N-terminal to the lysine residue.³⁶⁹ This gold cation-mediated selective dissociation enables site-specific opening of lysine-containing cyclic peptide ions and produces fragment ions with common N- and C-termini, resulting in more unambiguous sequencing than protonated counterparts that produce non-specific CID fragmentation.³⁷⁰ This method can aid in the development of drugs and pesticides using cyclic peptides^371–373 by accurately sequencing peptide structures.

Gas-phase charge inversion reactions can also facilitate metal ion transfer with lipids. For example, the ion/ion reaction between FAs ([FA − H]⁻) and magnesium tris-phenanthroline ([Mg(Phen)₃]²⁺) transfers Mg²⁺ to fatty acid ions via formation of an electrostatic complex [FA − H + MgPhen]⁺.³⁶⁸ Mg²⁺ restricts the positive charge to the carboxylic acid group on the fatty acid ion via strong non-covalent interactions, which unlocks charge-remote fragmentation of the hydrocarbon chain during CID of [FA − H + MgPhen]⁺ and facilitates unambiguous localization of C–C double bonds.^{307,374–376} By comparison, low-energy CID of the original [FA − H]⁻ results in uninformative loss of CO₂ from the carboxylic acid group as the primary dissociative channel.³⁶⁸ This charge-switching cationization method can also be applied to glycosphingolipids. The positive charge is restricted at the amide oxygen, enabling charge-remote fragmentation upon CID that allows localization of C–C double bond positions on fatty acid ions released from the glycosphingolipids.^377,378 In some cases, differences in coordination of the transferred metal allows for differentiation of highly similar compounds in formed ligand–metal–lipid complexes.^377,378 For example, sulfatides with either an α-glycosidic or β-glycosidic bond can be separated via a reaction with strontium tris-phenanthroline [Sr(Phen)₃]²⁺, as the diastereomers form complexes containing different numbers of phenanthroline ligands.³⁷⁹ This workflow can be implemented in shotgun lipidomics for differentiating multiple glycosphingolipid isomers in complex lipidomes without the need for chromatographic separation.^377,380,381 These ion/ion reactions have also been used to spatially map glycosphingolipid isomers during imaging mass spectrometry analyses of brain tissue.³⁷⁹

5.5. Covalent modifications

Numerous gas-phase ion/ion covalent modifications can be achieved using bioconjugation reagents, such as oxidation by periodate,³⁸² the formation of Schiff bases by aldehyde reagents,³⁸³ and the formation of amide^384,385 or anhydride³⁸⁶ bonds by ester reagents. The reagent ions are designed to have an electrostatically “sticky” functional group, such as sulf(on)ate and phosph(on)ate groups,³⁸⁷ to aid in forming stable, long-lived electrostatic complexes to allow sufficient time and proximity for covalent chemistry to occur.²⁹⁸ The exothermicity of the ion/ion reaction often drives the spontaneous formation of covalent bonds during the encounter,^308,388 signified by the neutral loss of specific byproducts as a result of the reaction or upon subsequent slow-heating activation, such as CID^{383,389–391} and IRMPD:³⁹²

[M + nH]ⁿ⁺ + [R − H]⁻ → [M + (n − 1)H + R]⁽ⁿ⁻¹⁾⁺ → [M + (n − 1)H + ▲]⁽ⁿ⁻¹⁾⁺ + (R − ▲) (▲: covalent adduct) (15)

Here, we highlight the gas-phase formation of amide bonds^367,392 and Schiff base^391,393 chemistry that accompanies metal ion transfer and charge inversion reactions, respectively.

The formation of amide bonds can be used for gas-phase cross-linking mass spectrometry via ion/ion reactions between multiply charged protein cations and singly charged ethylene glycol bis(sulfosuccinimidyl succinate) (sulfo-EGS) cross-linker anions. Mutual storage of these oppositely charged reactants leads to the formation of charge-reduced electrostatic complex cations [M + nH + (sulfo-EGS − H)]ⁿ⁻¹, followed by spontaneous covalent cross-linking at neutral basic residues, which is signified by the neutral loss of N-hydroxysulfosuccinimide (sulfo-NHS) upon activation.³⁸⁹ The distance restraints imposed by sulfo-EGS provide low-resolution protein conformational information, similar to the cross-linking mass spectrometry studies discussed earlier.^249,250 Substituting [sulfo-EGS − 2H + Na]⁻ for [sulfo-EGS − H]⁻ extends cross-linking sites to protonated basic residues by increasing the nucleophilicity of nitrogen atoms in ammonium and guanidinium groups via gas-phase sodium–proton exchange during the ion/ion reactions.³⁹² This enhanced cross-linking reactivity using the linker with sodium allows for differentiation between two gas-phase conformers of alcohol-denatured 8+ ubiquitin, which exhibit subtle conformational differences in the N-terminal region, by detecting additional modification sites compared to using the linker without sodium.³⁹²

A Schiff base can be formed in the gas phase during charge inversion ion/ion reactions between analyte ions containing an amine functional group and reagent ions containing an aldehyde functional group.^383,391,394 This process is a spontaneous covalent reaction, signified by neutral loss of water during CID of the ion/ion complex formed by the two reactants.^383,391,394 For example, mutual storage of a singly deprotonated phosphatidylserine (PS) anion and a doubly charged aliphatic reagent cation, N,N,N′,N′-tetramethyl-N,N′-bis(6-oxohexyl) hexane-1,6-diaminium (TMODA), leads to the formation of a charge-inverted [PS − H + TMODA]⁺ complex ion.³⁹³ Abstraction of a mobile proton from the carboxylic acid group in the serine headgroup is a critical step in the Schiff base condensation reaction, suggesting that phosphatidylserines can be distinguished from other lipid subclasses with different headgroup structures by this reaction.³⁹³ Indeed, isobaric PS 40:6 and SHexCer d38:1 are indistinguishable at nominal mass m/z 834 at low mass resolving power, but are readily differentiated via the Schiff base ion/ion reaction by the resulting 18 Da difference in the [PS 40:6 − H + TMODA − H₂O]⁺ and [SHexCer d38:1 − H + TMODA]⁺ product ions. The use of this ion/ion reaction in an imaging mass spectrometry experiment enables differentiation of these lipids in rat brain tissue, with the phosphatidylserine found localized to the cerebral cortex and the SHexCer localized to the corpus callosum and hippocampus (Fig. 10).³⁹³

Fig. 10 (a) A full scan imaging mass spectrometry experiment enables the (b) observation of a single peak at m/z 834.523 and (c) visualization of the H&E staining image and its corresponding spatial distribution. (d) An ion/ion reaction of the ion population at nominal m/z 834 using the TMODA reagent produces (e) a charge inverted product ion complex containing TMODA and SHexCer d38:1 as well as spontaneous water loss signifying Schiff base formation for PS 40:6, which are readily separated and (f) produce distinct spatial localizations where PS 40:6 are localized to the cerebral cortex and SHexCer d38:1 localized to the corpus collosum and hippocampus in the brain. (g) A CASI experiment improves the mass resolving power in the absence of the ion/ion reaction and (h) allows for the separation of SHexCer d38:1 and PS 40:6 using high mass resolving power and (i) validation of the distinct lipid spatial distributions. The analysis times of the ion/ion reaction (565 pixels) and CASI (558 pixels) imaging mass spectrometry experiments were roughly 45 min and 15 min, respectively. Adapted/reproduced from Analytical and Bioanalytical Chemistry, 2023, 415, 4319–4331 (ref. 393) with permission from Springer Nature, Copyright 2023.

The gas-phase ion/ion covalent modifications described here exhibit high reaction efficiency due to the coulombic attraction between oppositely charged ions and the high selectivity of analyte and reagent ions achieved through mass-selective ion isolation prior to the mutual storage for reaction. These features make ion/ion reactions an attractive alternative to their condensed-phase counterparts, which are often slower due to the necessary diffusion of reactants in solvent and can result in low purity because of incomplete and side reactions.²⁹⁸

6. Concluding remarks

The gas-phase reactions in mass spectrometry discussed in this review offer a multitude of approaches to address analytical challenges in a wide variety of sample types, enabling elegant experiments that reveal analyte identity and structure. Gas-phase reactions are generally faster and easier to control than in solution due to the hydrophobic vacuum environment, where analytes do not need to overcome energy barriers involving diffusion, competition between multiple reactions due to the presence of impurities, or difficulty in manipulating equilibrium. The utility of gas-phase chemistries in mass spectrometry, which offer superb control over reactant purity, ion concentration, and reaction kinetics by fine-tuning electronic parameters controlling ion sources, ion optics, and ion traps, is increasingly appreciated by researchers, leading to their widespread adoption in diverse fields of research. However, each technique discussed here also poses unique analytical challenges, some of which can be mitigated by a current trend of combining various complementary methods (e.g., HDX/ECD, CID/EID, CID/UVPD, EThcD) for more comprehensive information on the analytes. Some of these techniques are highly specialized with niche applications, while others hold promise for broad adoption. Continuous efforts to democratize novel approaches are essential for integrating different gas-phase methods, as this process may incur significant costs due to the potential requirement for expert knowledge, hardware and/or software modifications, or the acquisition of a brand-new instrument. An instrument that is flexible enough to perform most of the techniques highlighted here but that is still user-friendly and cost-effective would be ideal.

Despite the abundance of novel methods, there is still substantial room for future development and potential new applications through innovation in experimental and instrumental designs. Improvements to reaction and dissociation efficiencies will enable improvements in the limits of detection and sensitivity. Insights from adjacent chemical fields (e.g., organic and inorganic chemistry) can inspire the development and application of novel analytical mass spectrometry techniques, designed to elucidate challenging chemical structures (e.g., stereoisomers) and enhance our understanding of a broader range of analytes across various -omics disciplines. Cross-functional collaborations can lead to technological leaps in different fields, such as the development of quantum computers that use ion traps to confine Ca⁺ ions and wavelength-specific laser light to precisely manipulate the quantum states of trapped ions, which is essential for quantum gating.^395,396 Ion traps provide exceptional control over ion purity, long-term trapping stability, and sustained quantum coherence in well-isolated systems, contributing to their widespread use in quantum computing.^295,397 However, rapidly producing and accumulating multifaceted data on a large scale is inevitable considering the current and projected trends in multidisciplinary and multimethod research and automation. This trend requires the construction of powerful computational and software infrastructure to process and utilize big data more effectively and efficiently. Overall, we aim for this exploration of innovative applications and potential future directions in gas-phase ion chemistry to inspire creative solutions for analytical mass spectrometry challenges and to broaden the scope of gas-phase ion chemistry.

Conflicts of interest

There are no conflicts to declare.

Data availability

No primary research results, software or code have been included and no new data were generated or analysed as part of this review.

Acknowledgements

This work was supported by the National Institutes of Health (NIH) under Award R01 GM138660 (National Institute of General Medical Sciences (NIGMS)). WYK was supported by a Graduate Student Funding Award (GSFA) Fellowship from the University of Florida.

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366. K. A. Newton and S. A. McLuckey, Gas-phase peptide/protein cationizing agent switching via ion/ion reactions, J. Am. Chem. Soc., 2003, 125(41), 12404–12405.
367. B. M. Prentice, W. M. McGee, J. R. Stutzman and S. A. McLuckey, Strategies for the gas phase modification of cationized arginine via ion/ion reactions, Int. J. Mass Spectrom., 2013, 354–355, 211–218.
368. C. E. Randolph, D. J. Foreman, S. K. Betancourt, S. J. Blanksby and S. A. McLuckey, Gas-phase ion/ion reactions involving tris-phenanthroline alkaline earth metal complexes as charge inversion reagents for the identification of fatty acids, Anal. Chem., 2018, 90(21), 12861–12869.
369. D. J. Foreman, S. K. Betancourt, A. L. Pilo and S. A. McLuckey, Novel peptide ion chemistry associated with gold (I) cationization: Preferential cleavage at lysine residues, Int. J. Mass Spectrom., 2018, 427, 114–122.
370. D. J. Foreman, J. T. Lawler, M. L. Niedrauer, M. A. Hostetler and S. A. McLuckey, Gold(I) Cationization Promotes Ring Opening in Lysine-Containing Cyclic Peptides, J. Am. Soc. Mass Spectrom., 2019, 30(10), 1914–1922.
371. C. Lu, S. Nelson, G. Coy, C. Neumann, E. I. Parkinson and C. A. Rice, Cyclic Peptide Natural Product Inspired Inhibitors of the Free-Living Amoeba Balamuthia mandrillaris, J. Nat. Prod., 2025, 88(2), 274–281.
372. N. Chen, R. Huang, H. Lan, J. Jokela, H. Wei, S. Zhi and L. Liu, Cyclic peptides from cyanobacteria: Structural insights, biological functions, and mechanisms of cyclization, Algal Res., 2025, 86, 103925.
373. S. Zhang, S. Fan, H. He, J. Zhu, L. Murray, G. Liang, S. Ran, Y. Z. Zhu, M. J. Cryle, H.-Y. He and Y. Zhang, Cyclic natural product oligomers: diversity and (bio)synthesis of macrocycles, Chem. Soc. Rev., 2025, 54(1), 396–464.
374. C. E. Randolph, D. J. Foreman, S. J. Blanksby and S. A. McLuckey, Generating Fatty Acid Profiles in the Gas Phase: Fatty Acid Identification and Relative Quantitation Using Ion/Ion Charge Inversion Chemistry, Anal. Chem., 2019, 91(14), 9032–9040.
375. C. E. Randolph, D. L. Marshall, S. J. Blanksby and S. A. McLuckey, Charge-switch derivatization of fatty acid esters of hydroxy fatty acids via gas-phase ion/ion reactions, Anal. Chim. Acta, 2020, 1129, 31–39.
376. C. E. Randolph, D. S. M. Shenault, S. J. Blanksby and S. A. McLuckey, Structural Elucidation of Ether Glycerophospholipids Using Gas-Phase Ion/Ion Charge Inversion Chemistry, J. Am. Soc. Mass Spectrom., 2020, 31(5), 1093–1103.
377. H.-C. Chao and S. A. McLuckey, In-Depth Structural Characterization and Quantification of Cerebrosides and Glycosphingosines with Gas-Phase Ion Chemistry, Anal. Chem., 2021, 93(19), 7332–7340.
378. H.-C. Chao and S. A. McLuckey, Differentiation and Quantification of Diastereomeric Pairs of Glycosphingolipids Using Gas-Phase Ion Chemistry, Anal. Chem., 2020, 92(19), 13387–13395.
379. J. R. Bonney, A. E. Stratton, Y. Guo, C. B. Eades and B. M. Prentice, Imaging Mass Spectrometry of Sulfatide Isomers from Rat Brain Tissue Using Gas-Phase Charge Inversion Ion/Ion Reactions, J. Am. Soc. Mass Spectrom., 2025, 36(1), 119–126.
380. R. C. Barrientos and Q. Zhang, Recent advances in the mass spectrometric analysis of glycosphingolipidome – A review, Anal. Chim. Acta, 2020, 1132, 134–155.
381. K. Hořejší, D. Kolářová, R. Jirásko and M. Holčapek, Recent advances, challenges, and future directions in the mass spectrometry analysis of glycosphingolipids in biological samples, TrAC, Trends Anal. Chem., 2024, 178, 117827.
382. A. L. Pilo, F. Zhao and S. A. McLuckey, Gas-Phase Oxidation via Ion/Ion Reactions: Pathways and Applications, J. Am. Soc. Mass Spectrom., 2017, 28(6), 991–1004.
383. H. Han and S. A. McLuckey, Selective covalent bond formation in polypeptide ions via gas-phase ion/ion reaction chemistry, J. Am. Chem. Soc., 2009, 131(36), 12884–12885.
384. M. Mentinova and S. A. McLuckey, Covalent modification of gaseous peptide ions with N-hydroxysuccinimide ester reagent ions, J. Am. Chem. Soc., 2010, 132(51), 18248–18257.
385. J. Bu, Z. Peng, F. Zhao and S. A. McLuckey, Enhanced Reactivity in Nucleophilic Acyl Substitution Ion/Ion Reactions Using Triazole-Ester Reagents, J. Am. Soc. Mass Spectrom., 2017, 28(7), 1254–1261.
386. A. M. Pitts-McCoy, C. P. Harrilal and S. A. McLuckey, Gas-Phase Ion/Ion Chemistry as a Probe for the Presence of Carboxylate Groups in Polypeptide Cations, J. Am. Soc. Mass Spectrom., 2019, 30(2), 329–338.
387. M. Shih and S. A. McLuckey, Ion/ion charge inversion/attachment in conjunction with dipolar DC collisional activation as a selective screen for sulfo- and phosphopeptides, Int. J. Mass Spectrom., 2019, 444, 116181.
388. J. Bu, C. M. Fisher, J. D. Gilbert, B. M. Prentice and S. A. McLuckey, Selective Covalent Chemistry via Gas-Phase Ion/ion Reactions: An Exploration of the Energy Surfaces Associated with N-Hydroxysuccinimide Ester Reagents and Primary Amines and Guanidine Groups, J. Am. Soc. Mass Spectrom., 2016, 27(6), 1089–1098.
389. I. K. Webb, M. Mentinova, W. M. McGee and S. A. McLuckey, Gas-Phase Intramolecular Protein Crosslinking via Ion/Ion Reactions: Ubiquitin and a Homobifunctional sulfo-NHS Ester, J. Am. Soc. Mass Spectrom., 2013, 24(5), 733–743.
390. Y. Xia, B. A. Thomson and S. A. McLuckey, Bidirectional Ion Transfer between Quadrupole Arrays: MSn Ion/Ion Reaction Experiments on a Quadrupole/Time-of-Flight Tandem Mass Spectrometer, Anal. Chem., 2007, 79(21), 8199–8206.
391. N. Wang, A. L. Pilo, F. Zhao, J. Bu and S. A. McLuckey, Gas-phase rearrangement reaction of Schiff-base-modified peptide ions, Rapid Commun. Mass Spectrom., 2018, 32(24), 2166–2173.
392. W.-Y. Kang, A. Mondal, J. R. Bonney, A. Perez and B. M. Prentice, Structural Elucidation of Ubiquitin via Gas-Phase Ion/Ion Cross-Linking Reactions Using Sodium-Cationized Reagents Coupled with Infrared Multiphoton Dissociation, Anal. Chem., 2024, 96(21), 8518–8527.
393. X. Diao, N. R. Ellin and B. M. Prentice, Selective Schiff base formation via gas-phase ion/ion reactions to enable differentiation of isobaric lipids in imaging mass spectrometry, Anal. Bioanal. Chem., 2023, 415(18), 4319–4331.
394. J. R. Stutzman, K. M. Hassell and S. A. McLuckey, Dissociation behavior of tryptic and intramolecular disulfide-linked peptide ions modified in the gas phase via ion/ion reactions, Int. J. Mass Spectrom., 2012, 312, 195–200.
395. C. J. Ballance, T. P. Harty, N. M. Linke, M. A. Sepiol and D. M. Lucas, High-Fidelity Quantum Logic Gates Using Trapped-Ion Hyperfine Qubits, Phys. Rev. Lett., 2016, 117(6), 060504.
396. M. F. Brandl, M. W. van Mourik, L. Postler, A. Nolf, K. Lakhmanskiy, R. R. Paiva, S. Möller, N. Daniilidis, H. Häffner, V. Kaushal, T. Ruster, C. Warschburger, H. Kaufmann, U. G. Poschinger, F. Schmidt-Kaler, P. Schindler, T. Monz and R. Blatt, Cryogenic setup for trapped ion quantum computing, Rev. Sci. Instrum., 2016, 87(11), 113103.
397. C. D. Bruzewicz, J. Chiaverini, R. McConnell and J. M. Sage, Trapped-ion quantum computing: Progress and challenges, Appl. Phys. Rev., 2019, 6(2), 021314.

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