Mass Spectrometry Approaches for Investigating Alpha-Synuclein Structural Changes in Neurodegenerative Disease

Alpha-synuclein (aSyn) is a protein closely linked to neurodegenerative diseases such as Parkinson’s disease and Lewy body dementia because it can form harmful aggregates in the brain. Chemical changes to the protein, known as post-translational modifications (PTMs), including phosphorylation and oxidation, are commonly found in affected patients and may influence how the protein folds, interacts with other cellular components, and forms toxic structures. However, previous studies have produced conflicting results about whether these modifications increase or decrease protein toxicity and aggregation, making it important to better understand their structural effects.

Researchers at Indiana University Indianapolis used advanced mass spectrometry techniques to examine how phosphorylation and oxidation alter the shape and behavior of alpha-synuclein. By combining cross-linking and covalent labeling methods, the research identifies changes in protein folding, residue interactions, and solvent exposure with high sensitivity and structural detail. These spectrometry-based approaches provide valuable insight into how modified forms of alpha-synuclein may contribute to neurological disease progression and help improve understanding of the molecular mechanisms underlying synuclein-related disorders. LCGC International spoke to Ian Webb, corresponding author of a paper that resulted from this research.¹

How would you use mass spectrometry to distinguish and quantify different alpha-synuclein proteoforms (such as unmodified, pS129, fully oxidized methionines) in a heterogeneous sample, and what challenges arise due to alphasynuclein (aSyn) being intrinsically disordered?

Intact mass spectrometry can determine the molecular weight of each proteoform present in a heterogeneous sample. In this study, the proteoforms differ slightly in mass: unmodified aSyn is 14,460 Da, pS129 is 14,540 Da, and the fully methionine‑oxidized form is 14,524 Da. Top‑down mass spectrometry (TDMS) is the gold‑standard technique for distinguishing aSyn proteoforms because it isolates and fragments intact proteins, enabling direct localization and identification of post‑translational modifications. Quantification of proteoforms in heterogeneous samples can be achieved by deconvoluting the intact mass spectrum (through deisotoping and decharging) and comparing the relative abundances of the resolved species. These proteoforms are difficult to separate by liquid chromatography because aSyn is intrinsically disordered, adopts many conformations, and the proteoforms have very similar molecular weights.

What mass spectrometry (MS) strategies would you use to confidently localize phosphorylation at S129 and oxidation at specific methionines (M1, M5, M116, M127), and how would you address issues such as site ambiguity or neutral loss?

TDMS localizes and identifies post‑translational modifications (PTMs) by isolating and fragmenting intact proteins, allowing the modification site to be identified from the resulting fragment ions. For example, using electron‑based dissociation (ExD, including electron-capture dissociation [ECD] and electron-transfer dissociation [ETD]), phosphorylation at S129 produces characteristic +80 Da mass shifts. N‑terminal c‑type fragments exhibit a +80 Da shift on fragments that extend beyond S129, while C‑terminal z‑type fragments show the +80 Da shift on fragments formed N‑terminal to S129, together pinpointing the phosphorylation site. Addressing neutral loss is critical for phosphorylation, as labile phosphate groups are often lost during harsher fragmentation methods such as collision-induced dissociation (CID) or higher-energy collisional dissociation (HCD). ExD provides gentler fragmentation that better preserves this modification and also minimizes common neutral losses (‑H₂O, ‑NH₃, and ‑CO₂).

Similarly, methionine oxidation results in a +16 Da mass increase per oxidized residue (M1, M5, M116, and M127), which is reflected in the TD fragment ions. Under ECD/ETD conditions for fully-oxidized methionine, c‑type fragments would show a +16 Da shift for residues 1–4, a +32 Da shift for residues 5–115, a +48 Da shift for residues 116–126, and a +64 Da shift for fragments extending to the C‑terminus, consistent with oxidation at all four methionine sites.

In the context of studying post-translational modification (PTM) crosstalk and combinatorial proteoforms of aSyn, what are the advantages and limitations of top-down MS compared to bottom-up or middle-down approaches?

An advantage of TDMS for studying PTM crosstalk and combinatorial aSyn proteoforms is its ability to distinguish proteoforms with different intact masses and, in favorable cases, localize PTM sites. TDMS performs best when proteoforms are mass‑distinct; however, it has limitations when multiple proteoforms share the same nominal mass. In such cases, TDMS can reliably identify only the first and last modified residues along the sequence, while identical PTMs located internally may be ambiguous.

For example, aSyn contains four methionine residues (M1, M5, M116, and M127), each of which can be singly oxidized, producing proteoforms that differ from the unmodified protein by only +16 Da. During TDMS, all singly oxidized species are co‑isolated, resulting in highly convoluted MS/MS spectra. Consequently, fragment ions may simultaneously suggest both oxidized and unoxidized states at terminal residues such as M1 and M127. Once terminal oxidation is assigned, it becomes difficult to determine whether oxidation at internal residues (M5 or M116) reflects distinct proteoforms or fragment contributions from terminally oxidized species.

Bottom‑up and middle‑down approaches can identify all possible oxidation sites but generally cannot resolve distinct intact proteoforms. For instance, enzymatic digestion can reveal that all methionine residues are oxidized somewhere within a mixture, yet it cannot determine whether individual intact proteins are singly, doubly, triply, or quadruply oxidized.

How does cross-linking mass spectrometry function as a “molecular ruler” for intrinsically disordered proteins like aSyn, and what considerations are important when interpreting cross-link distance restraints for highly dynamic ensembles?

In this context, cross‑linkers are molecules that covalently connect one amino acid to another within the same protein and have a defined, measurable spacer length. This distance constraint allows cross‑linkers to function as molecular rulers that report on residue proximities within proteins. When applied to highly dynamic systems such as aSyn and other intrinsically disordered proteins (IDPs), it is important to recognize that observed cross‑links reflect transient contacts rather than stable structures. Because IDPs sample a wide range of conformations, the identified cross‑links represent snapshots of these rapidly interconverting ensembles.

What types of covalent labeling reagents would be appropriate for probing solvent accessibility and folding in aSyn and how would changes in labeling patterns indicate PTM-induced conformational shifts?

Covalent labeling reagents can be broadly classified as either specific or non‑specific. Non‑specific labels are particularly useful for probing protein solvent accessibility and folding because they can modify multiple residue types, providing broader sequence coverage and richer topological information. Differences in labeling patterns between proteoforms—for example, one proteoform showing covalent labeling at the C‑terminus while another does not—can indicate changes in solvent accessibility arising from PTM‑induced conformational differences.

How would you integrate data from cross-linking MS and covalent labeling MS to build a coherent model of how S129 phosphorylation or methionine oxidation alters the conformational ensemble of aSyn?

Cross‑linking mass spectrometry provides information on residue–residue interactions and distance constraints, whereas covalent labeling mass spectrometry reports on solvent accessibility and local microenvironment. Integrating these complementary datasets—regions involved in intra‑protein contacts and regions exposed to solvent—enables the construction of coherent models describing how specific PTMs reshape the conformational landscape of aSyn. For example, pS129‑aSyn exhibits more long‑range cross‑links than wild‑type aSyn across all cross‑linkers tested, suggesting a higher population of folded conformers. Consistently, diethyl pyrocarbonate (DEPC) labeling of pS129‑aSyn resulted in modification of serine, threonine, and tyrosine residues. Previous work from Richard Vachet’s group has shown that DEPC labeling of S, T, and Y residues requires a hydrophobic microenvironment. Together, these observations indicate that pS129‑aSyn adopts a more hydrophobic and folded conformational ensemble.

What additional information can ion mobility–mass spectrometry (IM-MS) provide about aSyn structural heterogeneity, and how would you interpret changes in collision cross sections upon phosphorylation or oxidation?

Ion mobility–mass spectrometry provides insight into the gas‑phase conformational ensembles that underlie the structural heterogeneity of α‑synuclein (aSyn). Ion mobility is an orthogonal separation technique to liquid chromatography, resolving ions in the gas phase based on their collision cross section (CCS) through differences in arrival time. Distinct aSyn conformer classes can be distinguished by the presence of multiple ion‑mobility peaks, corresponding to different shapes or conformations. A decrease in collisional cross section upon phosphorylation would indicate that the modified proteoform adopts more compact gas‑phase structures.

What are the key sample preparation challenges when analyzing aSyn by MS, and how would you minimize experimental bias?

Key sample preparation considerations for aSyn include thawing the protein slowly on ice, minimizing freeze–thaw cycles, avoiding bubble formation, and ensuring the absence of contaminants during purification. When analyzing intact aSyn by electrospray ionization mass spectrometry (ESI‑MS), it is critical to use the lowest possible spray voltage that still maintains stable electrospray and good signal‑to‑noise. Elevated voltages can significantly alter the charge‑state distribution and may induce aggregation. To minimize experimental bias, results are typically evaluated in comparison with prior studies to assess consistency across techniques and methodologies.

How can MS-based structural data be correlated with functional outcomes such as altered SNARE (soluble N-ethylmaleimide-sensitive factor attachment protein receptors) binding, vesicle trafficking, or aggregation propensity, given that MS experiments are typically performed in vitro?

MS‑based structural data can be correlated with functional outcomes like altered SNARE binding, vesicle trafficking, or aggregation propensity by integrating mass spectrometry measurements with biological activity assays. Native mass spectrometry has been particularly valuable for these studies because it preserves native‑like protein structures in the gas phase, despite being performed in vitro, enabling meaningful structural insights into protein behavior.

Given the conflicting reports on whether pS129 or methionine oxidation increases or decreases aSyn aggregation and toxicity, how can mass spectrometry uniquely contribute to resolving these discrepancies compared to nuclear magnetic resonance (NMR), circular dichroism (CD), or fluorescence-based techniques?

Mass spectrometry provides high‑resolution, site‑specific structural information that many other techniques cannot, while also distinguishing between coexisting proteoforms and directly identifying their oligomeric states (for example, monomers, dimers, or higher‑order assemblies). In contrast, NMR and CD spectroscopy have limited ability to differentiate between proteoforms. NMR additionally struggles with insoluble aggregates, while CD lacks the residue‑level resolution needed to pinpoint structural changes arising from PTMs such as S129 phosphorylation or methionine oxidation. Fluorescence‑based techniques typically rely on extrinsic labels to visualize aSyn, but these labels can perturb the native monomer structure and alter aggregation propensity, complicating interpretation of the results.

References

1. Dollar, A. N.; Webb, I. K. Cross-Linking and Covalent Labeling Mass Spectrometry Reveal Proteoform-Driven Conformational Changes in Alpha Synuclein. Anal Chem 2026.DOI: 10.1021/acs.analchem.5c05078