Stable Isotopes in Proteomics

Stable Isotopes in Proteomics
Steven Cockrill, Justin Wildsmith, Kelly Foster, Tom Hassell
Sigma-Aldrich R&D, St. Louis, MO

Proteomics is the systematic study of proteins encoded by the genome. This involves all aspect of protein characterization: identification of the post-translational modifications (PTM), localization, structure and ultimately identification and quantitation. Specifically, quantitation has been described for both protein expression relative to the changes in response to external and internal perturbations, as well as on an absolute scale ^(1,2).

Quantitative proteomics has recently emerged as a complementary technology to mRNA profiling with the ability to comprehensively characterize the structural and functional aspects of protein species ⁽³⁾. The general approach to quantitative proteomics involving selective tagging, stable isotope dilution and mass spectrometry has helped initiate a variety of different isotopic labeling methods. Stable isotope incorporation has been achieved in a variety of ways, including metabolic labeling ⁽⁴⁾, enzymatically directed methods ⁽⁵⁾, and chemical labeling using externally introduced tags. Use of stable isotope incorporation involves the addition to a sample of chemically identical form of the analyte(s), containing stable heavy isotopes (e.g., 2H, 13C, 15N, etc.) as an internal standard. Because of the ionization variability for different peptides, the best internal standard for peptide quantitation is the same peptide labeled with stable isotopes. This way, relative protein quantitation is achieved when two proteins or protein mixtures are compared where one serves as a control sample. The control and test samples differ only by the incorporation of either the heavy stable isotopes, or the native isotopes. In theory, all peptides in the combined sample exist as analyte pairs of identical sequence but differing masses. The peptide have the same properties and behave with essentially identical characteristics under any isolation or separation step. Thus the ratio between intensities of the pairs provides an accurate relative peptide and therefore protein abundance measure. There are several protocols for the use of stable isotopes in proteomics.

The technique that has received the most attention is the use of Isotope-Coded Affinity Tags (ICATTM) methodology ⁽⁶⁾. ICAT reagents contain three moieties: a thiol specific reactive group, a linker region with light or heavy isotopes, and a biotin tag for affinity isolation. One of the major advantages of the ICATTM strategy is that the labeling reaction is compatible with commonly used reagents for 2D electrophoresis. The ICATTM strategy has been adopted into a number of proteomics studies ⁽⁶⁾. The ICATTM technique has some limitations in that it requires cystein-containing proteins and it provides only relative quantitation. A more recent development of ICAT is ITRAQ™, in which an isobaric tag with up to four different stable isotope mass tags is incorporated, permits multiplexing of sample populations. The isobaric nature of the gross tag requires that quantitation occurs in MS/MS rather than MS mode, simplifying analysis, but necessitating higher performance instrumentation. Other labeling strategies include GIST (define) and ICPL (define) techniques, which similarly allow multiplexing but also require MS/MS capability.

Another technique involving comparative proteomics, stable isotope labeling with amino acids in cell culture (SILAC), has recently gained popularity for its ability to compare the expression levels of hundreds of proteins in a single experiment. SILAC was developed in CEBI ⁽⁴⁾ as a simple and accurate approach for MS-based quantitative proteomics. The method relies on the incorporation of ²H, ¹³C, ¹⁵N-labeled amino acids added to the growth media of separately cultured cell lines, giving rise to cells containing either "light" or "heavy" proteins, respectively. Upon mixing lysates collected from these cells, digests of the protein populations are analyzed by mass spectrometry. Using this method, one can compare the expression levels for hundreds of proteins.

Enzymatic labeling involves the use of proteases to incorporate a mass label, ¹⁸O, into virtually all generated peptides ^(1,2). This is accomplished by performing digestion in the presence of isotopically enriched water. This technique offers the advantage of ease of implementation. Because oxygen atoms in a sample do not spontaneously exchange in the absence of the protease, this method is highly specific for stable isotope incorporation into the carboxy terminus of each new peptide. Only the protein C-terminal peptide will not have a stable isotope incorporated. This peptide can then be easily identified in labeled samples by the lack of a mass shift. Depending on the protease utilized for enzymatic labeling, either one or two ¹⁸O atoms can be incorporated into peptides. Trypsin, Lys-C and Glu-C are all capable of incorporating two ¹⁸O atoms per peptide. This is due to the nature of the interaction between peptide and protease. Trypsin digestion in ¹⁸O water, therefore results in mass spectra with ion doublets spaced by either 2 or 4 Da. This provides a means of differentiation from analogous samples digested in natural water. In addition, covalently linked binary peptides will display an 8 Da shift when digested in ¹⁸O water. This allows for the study of protein structure through identification of internal disulfide bonds and the detection of interacting domains in protein complexes using chemical crosslinking methods.

Although the above techniques allow the comparison of expressed proteins, the absolute quantity of a targeted protein could not be readily measured from a complex mixture until the development of the AQUATM (Absolute Quantification) strategy ⁽²⁾. This approach uses ¹³C-labeled reference peptides and tandem MS to measure expression in terms of number of molecules per cell. Reference peptides, corresponding to specific, pre-defined proteolytic fragments of proteins of interest, are synthesized with an incorporated stable isotope. Samples containing the protein of interest are then subjected to complete digestion with trypsin, following the addition of the isotopically labeled peptide in a known amount. LC-MS is subsequently used to quantitate the naturally occurring peptide from the protein of interest by comparison with the level of the corresponding internal standard. This technique is made possible by the use of stable isotopes to generate peptide standards that are structurally identical but occur 2-10 Daltons higher in mass to charge. This enables one to quantitate peptides in a stoichiometric sense using the synthetic peptide as internal standard. The AQUATM method is precise and highly specific.

Quantitative proteomics based on stable isotope tagging and automated mass spectrometry is a maturing technology that can interface with a multitude of biological and clinical research initiatives ⁽⁷⁾. Mass spectrometry-based analysis of protein mixtures is rapidly becoming a mainstream technology in proteomics. With advances in MS and LC-MS technologies, stable isotope labeling has gained increased attention as a crucial tool for differential proteomics. Stable isotope labeled peptides and proteins exhibit near-identical HPLC elution profiles but distinctively different MS spectra, compared to their endogenous peers. This unique feature leads to the possibility of quantitative studies of peptide and protein populations, which is vital in the field of proteomics. Isotec provides scientists with a wide range of isotope labeled reagents and growth media. With LC-MS, genetic engineering techniques, and a commercial stable isotope supplier, the opportunities for today's biochemists are boundless. The proteomics field may be the most prosperous research field in the near future. The only limitation here is scientific imagination.

References

1. R. Aebersold, M. Mann, (2003) Nature 422, 198-207.
2. S. A. Gerber, J. Tush, O. Stemman, M.W. Kirschner, S. P. Gypi, (2003) Proc. Natl. Acad. Sci. USA 100, 6940-5.
3. S. Fields, Proteomics in Genomeland, (2001)Science 291, 1221-4.
4. S.E. Ong et al., Mol Cell Proteomics (2002) 1, 376-86.
5. A. Shevchenko et al., (1997) Rapid Commun. Mass Spectrom. 11, 1015-24.
6. S.P. Gygi et al., (1999) Nat. Biotechno.l 17, 994-9.
7. M.R. Flory, T. J. Griffin, D. Martin and R. Aebersold, (2002) Trends Biotechnol. 20, 23-29.