A ubiquitous yet underappreciated protein post-translational modification, isoaspartic acid (isoAsp, isoD or β-Asp), generated via the deamidation of asparagine or isomerization of aspartic acid in proteins, plays a diverse and crucial role in ageing, as well as autoimmune, cancer, neurodegeneration and other diseases. In addition, formation of isoAsp is a major concern in protein pharmaceuticals, as it may lead to aggregation or activity loss. The scope and significance of isoAsp have, up to now, not been fully explored, as an unbiased screening of isoAsp at low abundance remains challenging. This difficulty is due to the subtle difference in the physicochemical properties between isoAsp and Asp, e.g., identical mass. In contrast, endoprotease Asp-N (EC 3.4.24.33) selectively cleaves aspartyl peptides but not the isoaspartyl counterparts. As a consequence, isoaspartyl peptides can be differentiated from those containing Asp and also enriched by Asp-N digestion. Subsequently, the existence and site of isoaspartate can be confirmed by electron transfer dissociation (ETD) mass spectrometry. As little as 0.5 % of isoAsp was detected in synthetic beta amyloid and cytochrome c peptides, even though both were initially assumed to be free of isoAsp. Taken together, our approach should expedite the unbiased discovery of isoAsp.
Arising from spontaneous aspartic acid (Asp) isomerization or asparagine (Asn) deamidation, isoaspartic acid (isoAsp, isoD, or beta-Asp) is a ubiquitous nonenzymatic modification of proteins and peptides. Because there is no mass difference between isoaspartyl and aspartyl species, sensitive and specific detection of isoAsp, particularly in complex samples, remains challenging. Here we report a novel assay for Asp isomerization by isotopic labeling with (18)O via a two-step process: the isoAsp peptide is first specifically methylated by protein isoaspartate methyltransferase (PIMT, EC 2.1.1.77) to the corresponding methyl ester, which is subsequently hydrolyzed in (18)O-water to regenerate isoAsp. The specific replacement of (16)O with (18)O at isoAsp leads to a mass shift of 2 Da, which can be automatically and unambiguously recognized using standard mass spectrometry, such as collision-induced dissociation (CID), and data analysis algorithms. Detection and site identification of several isoAsp peptides in a monoclonal antibody and the β-delta sleep-inducing peptide (DSIP) are demonstrated.
Cystine knots or nested disulfides are structurally difficult to characterize, despite current technological advances in peptide mapping with high-resolution liquid chromatography coupled with mass spectrometry (LC-MS). In the case of recombinant human arylsulfatase A (rhASA), there is one cystine knot at the C-terminal, a pair of nested disulfides at the middle, and two out of three unpaired cysteines in the N-terminal region. The statuses of these cysteines are critical structure attributes for rhASA function and stability that requires precise examination. We used a unique approach to determine the status and linkage of each cysteine in rhASA, which was comprised of multi-enzyme digestion strategies (from Lys-C, trypsin, Asp-N, pepsin, and PNGase F) and multi-fragmentation methods in mass spectrometry using electron transfer dissociation (ETD), collision induced dissociation (CID), and CID with MS3 (after ETD). In addition to generating desired lengths of enzymatic peptides for effective fragmentation, the digestion pH was optimized to minimize the disulfide scrambling. The disulfide linkages, including the cystine knot and a pair of nested cysteines, unpaired cysteines, and the posttranslational modification of a cysteine to formylglycine, were all determined. In the assignment, the disulfide linkages were Cys138 - Cys154, Cys143 - Cys150, Cys282 - Cys396, Cys470 - Cys482, Cys471 - Cys484, and Cys475 - Cys481. For the unpaired cysteines, Cys20 and Cys276 were free cysteines, and Cys51 was largely converted to formylglycine (> 70%). A successful methodology has been developed which can be routinely used to determine these difficult-to-resolve disulfide linkages, ensuring drug function and stability.
The formation of isoaspartyl residues (isoAsp or isoD) via either aspartyl isomerization or asparaginyl deamidation alters protein structure and potentially biological function. This is a spontaneous and non-enzymatic process, ubiquitous both in vivo and in non-biological systems, such as in protein pharmaceuticals. In almost all organisms, protein L-isoaspartate Omethyltransferase (PIMT, EC2.1.1.77) recognizes and initiates the conversion of isoAsp back to aspartic acid. Additionally, alternative proteolytic and excretion pathways to metabolize isoaspartyl-containing proteins have been proposed but not fully explored, largely due to the analytical challenges for detecting isoAsp. We report here the relative quantitation and site profiling of isoAsp in urinary proteins from wild type and PIMT-deficient mice, representing products from excretion pathways. First, using a biochemical approach, we found that the total isoaspartyl level of proteins in urine of PIMT-deficient male mice was elevated. Subsequently, the major isoaspartyl protein species in urine from these mice were identified as major urinary proteins (MUPs) by shotgun proteomics. To enhance the sensitivity of isoAsp detection, a targeted proteomic approach using electron transfer dissociation-selected reaction monitoring (ETD-SRM) was developed to investigate isoAsp sites in MUPs. Thirty-eight putative isoAsp modification sites in MUPs were investigated, with five derived from the deamidation of asparagine that were confirmed to contribute to the elevated isoAsp levels. Our findings lend experimental evidence for the hypothesized excretion pathway for isoAsp proteins. Additionally, the developed method opens up the possibility to explore processing mechanisms of isoaspartyl proteins at the molecular level, such as the fate of protein pharmaceuticals in circulation.
Formation of aspartyl succinimide (Asu) is a common post-translational modification (PTM) of protein pharmaceuticals under acidic conditions. We present a method to detect and quantitate succinimide in intact protein via hydrazine trapping and chemical derivatization. Succinimide, which is labile under typical analytical conditions, is first trapped with hydrazine to form stable hydrazide and can be directly analyzed by mass spectrometry. The resulting aspartyl hydrazide can be selectively derivatized by various tags, such as fluorescent rhodamine sulfonyl chloride that absorbs strongly in the visible region (570 nm). Our tagging strategy allows the labeled protein to be analyzed by orthogonal methods, including HPLC-UV, LC-MS, and SDS-PAGE coupled with fluorescence imaging. A unique advantage of our method is that variants containing succinimide, after derivatization, can be readily resolved via either affinity enrichment or chromatographic separation. This allows further investigation of individual factors in a complex protein mixture that affect succinimide formation. Some additional advantages imparted by fluorescence labeling include, the facile detection of the intact protein without proteolytic digestion to peptides; and high sensitivity, e.g. without optimization 0.41% succinimide was readily detected. As such, our method should be useful for rapid screening, optimization of formulation conditions and related processes relevant to protein pharmaceuticals.
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