Protein disulfide bond determination by mass spectrometry
Authors: Gorman Jeffrey J. , Wallis Tristan P. , Pitt James J. 
Published: December 2002

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 18O, And Disulfide Analysis
191

A. Incorporation of 18O Into Peptides
191

B. Identification of Disulfide‐Linked Peptides
193

C. Comparison of Tryptic and Peptic Cleavage in 50% H218O
196

D. 18O Isotope Profiles of Single‐Chain and Disulfide‐Linked Peptides
198

E. Stability of 18O Isotope Profiles During Chromatography and Storage
201

F. Application of Pepsin‐Mediated 18O Incorporation to a Large Disulfide‐Linked Protein
203

IV.
Tandem Mass Spectrometry of 18O‐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% H218O. Resultant disulfide‐linked peptides have distinct isotope profiles (combinations of isotope ratios and average mass increases) compared to the same peptides with only 16O 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 18O 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