Deciphering protein long-chain S-acylation using mass spectrometry proteomics strategies

Abstract

Protein long-chain S-acylation, the reversible attachment of fatty acids such as palmitate to cysteine residues via thioester bonds, is a widespread post-translational modification that plays a crucial role in regulating protein localization, trafficking, and stability. Despite its prevalence and biological relevance, the study of long-chain S-acylation has long lagged behind that of other dynamic PTMs due to the hydrophobic nature and lability of the lipid modification, which complicate conventional proteomic workflows. Recent advances in mass spectrometry-based strategies have significantly expanded the toolbox for studying long-chain S-acylation, with improved workflows enabling more sensitive, site-specific, and quantitative analysis. This review summarizes key developments from the past decade across both direct and indirect mass spectrometry-based strategies, including acyl-biotin exchange, lipid metabolic labeling, and novel enrichment and fragmentation methods. We also highlight emerging challenges in distinguishing lipid-specific modifications, achieving robust quantification, and mitigating artifacts from in vitro systems, while outlining future directions to advance functional and therapeutic exploration of the S-acyl-(prote)ome.

Fig. 1. Overview of the S-acylation regulatory machinery. Long-chain S-acylation is regulated by 23 protein S-acyl transferases and seven validated acyl-protein thioesterases.

Fig. 2. Acyl-biotin exchange and acyl-RAC workflows for identifying S-acylated proteins and modification sites via mass spectrometry. (A) Overview of ABE and acyl-RAC common to both workflows: proteins are treated with a reducing agent to reduce disulfide bonds into free thiols, while leaving S-acylated cysteines unaffected. Free thiols are then capped using an alkylating reagent. Acyl-RAC workflow: step 1a. Following hydroxylamine treatment, free thiols are captured on thiopropyl sepharose resin. Step 1b. Captured proteins are released from the resin with a reducing agent. Proteins are digested and measured by LC-MS/MS. ABE workflow: step 2a. Thioester bonds are selectively cleaved with hydroxylamine (neutral pH), converting S-acylated cysteines into free thiols, which are then biotinylated (e.g. using HPDP-biotin). Step 2b. Biotinylated proteins are enriched via NeutrAvidin resin and selectively eluted with a reducing agent. Step 2c. Eluted proteins are digested, cleaned, and analyzed using LC-MS/MS. In site-specific workflows proteins are digested prior to enrichment producing a mixture of biotinylated, alkylated and unmodified peptides (ssABE) or a mixture of thiopropyl sepharose captured peptides (site-specific acyl-RAC). (B) Chemical structures of commonly used cysteine-capping (alkylating) reagents, and the most widely used biotinylating reagent, HPDP-biotin.

Fig. 3. Lipid metabolic labeling workflows to detect S-acylated proteins and sites. (A) Classical LML: step 1. Cells are treated with various clickable fatty acid analogues resulting in alkyne-tagged proteins. Step 2. Alkyne-tagged proteins are conjugated to a biotin-azide reagent. Step 3. Biotinylated proteins are enriched using NeutrAvidin resin. S-Acylated proteins/peptides are selectively eluted of the NeutrAvidin resin using hydroxylamine. Step 4. Proteins are digested using e.g. trypsin (if performing a siteID workflow, biotinylated proteins are digested prior to enrichment), subjected to peptide clean-up and subsequent LC-MS/MS measurement. (B) Schematic representation of the “bump-and-hole” strategy for selective ZDHHC substrate identification. Cells are labelled with clickable “bump”-Alk probes (bumps: cPr = cyclopropyl, Ac = acetyl, Bz = benzoyl). Following intracellular activation by acyl-CoA ligase, transfer of the “bump”-Alk probes to wild-type ZDHHCs is blocked by steric hindrance, whereas “hole”-engineered ZDHHCs accommodate the probes during auto-acylation. The modified ZDHHC enzyme subsequently transfers the “bump”-Alk probe to its substrates. After cell lysis, enrichment handles are installed selectively on substrates carrying the “bump”-Alk modification through CuAAC-mediated click chemistry, enabling downstream enrichment and analysis. Schematic representation was inspired by Ocasio et al. (C) Schematic of two typical pulse-chase experiments, showing a multi-S-acylated protein to highlight the need for site-specific analyses. Cells are first labelled with clickable fatty acid analogues (“pulse”), followed by a wash-out period with palmitic acid (“chase”). Step 1. The palmitic acid chase is compared to a vehicle chase to gain information on lipid turnover. Step 2. The palmitic acid chase is supplemented with a deacylase inhibitor or a vehicle. Sites with a high turnover rate will quickly replace their alkynyl-acyl group with palmitate, sites that have low turnover rate, or that are stabilized by acyl-protein thioesterase inhibition, will retain their alkynyl-acyl group during the chase.

Fig. 4. Current techniques in the direct detection of protein S-acylation with mass spectrometry. Peptide-centric (bottom-up) approaches: workflow 1. Isolated proteins are digested with protease(s) and separated (off-line or on-line) by liquid chromatography using a C₄, C₈ or C₁₈ stationary phase (off-line for MALDI-TOF-MS, on-line for ESI-MS/MS) and analyzed either by MALDI-TOF MS or ESI-MS/MS. Workflow 2. Complex mixtures are digested and S-acylated peptides are enriched using nGF-SPE followed by LC-MS/MS analysis. Lipidomics: workflow 3. Complex mixtures undergo multiple precipitations, after which the pellet and MeOH layer are treated with hydroxylamine to acquire fatty acid hydroxamates from S-acylated proteins and acyl-CoAs, respectively. Protein-centric approaches: workflow 4. Native/intact MS approaches analyze proteins without digestion or fragmentation. These approaches can distinguish between different S-acyl-proteoforms but cannot pinpoint individual S-acylation sites. Workflow 5. Top-down MS can identify distinct S-acylation sites after fragmentation of the protein backbone. Workflow 6. Native top-down MS ejects complexes from their native membrane environment using a CO₂ laser. IRMPD fragmentation allows for protein sequencing with the advantage that S-acylations are not lost during fragmentation.

Anneroos E. Nederstigt

Samiksha Sardana

Marc P. Baggelaar