Abstract
| I. | Introduction | 184 |
| II. | Mass Spectrometry and Disulfide Bond Determination | 185 |
| | A. Peptide Mass Analysis | 185 |
| | B. Peptide Mass Analysis for Determination of the Disulfides of Newcastle Disease Virus (NDV) Hemagglutinin‐Neuraminidase (HN) | 188 |
| | C. Tandem Mass Spectrometry (MS/MS) | 189 |
| III. | Stable Isotope‐Labeling With ^18^O, And Disulfide Analysis | 191 |
| | A. Incorporation of ^18^O Into Peptides | 191 |
| | B. Identification of Disulfide‐Linked Peptides | 193 |
| | C. Comparison of Tryptic and Peptic Cleavage in 50% H~2~^18^O | 196 |
| | D. ^18^O Isotope Profiles of Single‐Chain and Disulfide‐Linked Peptides | 198 |
| | E. Stability of ^18^O Isotope Profiles During Chromatography and Storage | 201 |
| | F. Application of Pepsin‐Mediated ^18^O Incorporation to a Large Disulfide‐Linked Protein | 203 |
| IV. | Tandem Mass Spectrometry of ^18^O‐Labeled Disulfide‐Linked Peptides | 205 |
| V. | Ancillary Methods and Considerations | 206 |
| VI. | Conclusions | 211 |
| Acknowledgments | 212 |
| References | 212 |
The determination of disulfide bonds is an important aspect of gaining a comprehensive understanding of the chemical structure of a protein. The basic strategy for obtaining this information involves the identification of disulfide‐linked peptides in digests of proteins and the characterization of their half‐cystinyl peptide constituents. Tools for disulfide bond analysis have improved dramatically in the past two decades, especially in terms of speed and sensitivity. This improvement is largely due to the development of matrix‐assisted laser desorption/ionization (MALDI) and electrospray ionization (ESI), and complementary analyzers with high resolution and accuracy. The process of pairing half‐cystinyl peptides is now generally achieved by comparing masses of non‐reduced and reduced aliquots of a digest of a protein that was proteolyzed with intact disulfide bonds. Pepsin has favorable properties for generating disulfide‐linked peptides, including its acidic pH optimum, at which disulfide bond rearrangement is precluded and protein conformations are likely to be unfolded and accessible to cleavage, and broad substrate specificity. These properties potentiate cleavage between all half‐cystine residues of the substrate protein. However, pepsin produces complex digests that contain overlapping peptides due to ragged cleavage. This complexity can produce very complex spectra and/or hamper the ionization of some constituent peptides. It may also be more difficult to compute which half‐cystinyl sequences of the protein of interest are disulfide‐linked in non‐reduced peptic digests. This ambiguity is offset to some extent by sequence tags that may arise from ragged cleavages and aid sequence assignments. Problems associated with pepsin cleavage can be minimized by digestion in solvents that contain 50% H~2~^18^O. Resultant disulfide‐linked peptides have distinct isotope profiles (combinations of isotope ratios and average mass increases) compared to the same peptides with only ^16^O in their terminal carboxylates. Thus, it is possible to identify disulfide‐linked peptides in digests and chromatographic fractions, using these mass‐specific markers, and to rationalize mass changes upon reduction in terms of half‐cystinyl sequences of the protein of interest. Some peptides may require additional cleavages due to their multiple disulfide bond contents and/or tandem mass spectrometry (MS/MS) to determine linkages. Interpretation of the MS/MS spectra of peptides with multiple disulfides in supplementary digests is also facilitated by the presence of ^18^O in their terminal carboxylates. © 2002 Wiley Periodicals, Inc., Mass Spec Rev 21:183–216, 2002; Published online in Wiley InterScience (www.interscience.wiley.com). DOI 10.1002/mas.10025