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October 2001
Vol. 10, No. 10,
pp 30–32, 34–35.
 
 
 
Today's Chemist at Work
Focus: Organic Spectroscopy

FEATURE

Proteins on mass (spectrometry, that is)

Single and tandem MS using ESI or MALDI techniques are providing the means to bring protein analysis into the 21st century, paving the road to proteomics.

Genomes are passé. Proteins rule. That is the current mantra of biomedical research. Not because all genomes are a completed enterprise by any means, but rather because proteins are where the action is—where the biochemical rubber meets the physiological road. And in instrumental analysis, the same seems true as well.

DNA is child’s play compared with proteins when it comes to using automated analytical techniques. DNA has its polymerase chain reaction (PCR) to make mass quantities of gene sequences for study. Genes are clonable, sequencers automated, and even the majority of research in bioinformatics has been tagged to the analysis of genomes from the start. But as for proteins and the proteome (an organism’s complete complement of proteins), the technologies are still emerging. Because so many proteins are unknowns and exist in such small quantities with no PCR-analogous means for scale-up, more sophisticated analytical techniques must be developed. As part of such an enterprise, mass spectrometry (MS) is key.

When coupled to 2-D electrophoresis (2DE), and more recently to single or multidimensional HPLC or capillary electrophoresis (CE), a variety of MS techniques—including electrospray ionization (ESI)-MS, ion-trap instruments, and matrix-assisted laser desorption/ionization (MALDI) MS—are showing promise for providing unique and rapid identification of a multiplicity of proteins (and even glycoprotein moieties). This explosive growth in MS research dedicated to solving the protein problem is moving biomedical science ever closer to the degree of automation and sensitivity that the proteomics revolution demands.

Problematic Proteins
In the 1950s, a key breakthrough in protein sequencing occurred with the development of the Edman degradation, which uses phenylisothiocyanate to cleave off the N-terminal amino acid from a protein. As the reaction runs, amino acids are cleaved off in order, one by one, and can be purified and identified using standard techniques. By the late 1960s, the Edman reaction was adopted as the basis for the first automated amino acid sequencers. But such automated sequencers are proving too slow for the demands of the biotech revolution. For this reason, MS techniques are being adapted as the ultimate replacement technology for protein sequencing and identification.

But how to get the proteins to study? Which ones are worth analyzing?

“Out Damned Spot!”
Perhaps the most significant breakthrough in protein analysis as it relates to biological systems has been the development and refinement of 2DE, which separates proteins from one another so that they can be studied individually. Typically, the proteins are first separated on an isoelectric focusing gel, which is then saturated with sodium dodecyl sulfate and placed on an SDS-PAGE gel, which then is run to size the proteins by molecular weight. The first dimension is thus a separation by pI along the x-axis and by molecular weight along the y-axis of the final 2DE gel. Finding the unique proteins that vary in differential 2DE displays has led to a host of breakthroughs in animal and plant physiology. One major benefit of this technique is the ability to excise the proteins of interest from their visualized spots (labeled radioactively with coumassie blue, fluorescent markers, or silver stains) for subsequent analysis.

Differential 2DE imaging is an extraordinarily powerful technique with numerous commercial imaging systems that make it ever more effective. With differential display, proteins are isolated from treated or untreated cells, healthy or cancerous—any two states or tissues that it is desirable to compare—and run on separate gels using the same conditions. Upon staining, visual inspection or imaging analysis is used to reveal the presence or absence of protein spots that show a differential between the samples being analyzed. These are the spots that are excised for further study.

Unfortunately, gels have the drawback of being slow, and they are often highly inefficient for the isolation of very low concentration proteins. The stains typically used, along with the gel matrix itself, can sometimes interfere with subsequent analysis. Gel techniques until recently proved difficult to automate, although there has been some progress. For example, the Proteomeworks System developed through an alliance between Micromass, Ltd. (U.K.) and Bio-Rad Laboratories, Inc. (USA), was designed to automate various steps in the processes of making, running, and analyzing the results of 2DE.

Recently, multidimensional HPLC techniques have been designed that have the promise of supplanting the use of 2DE in many applications while providing the substantial benefits of working exclusively in liquids rather than gels. The drawback to HPLC is the relative inability to visualize from simple image inspection those proteins that are of particular interest for subsequent analysis.

Regardless of whether a protein spot is eluted from a gel or from a protein peak separated on an HPLC, however, there remains one final choice before entering an MS system—to digest or not digest. Typically, the first step in analyzing a 2DE protein spot (or a particular HPLC peak) has been digestion of the protein into peptide fragments using the enzyme trypsin, which cuts proteins specifically on the C-terminal side of every lysine and argine in the amino acid sequence (except when these bonds are to prolines). For such analysis, protein spots from 1- or 2DE are dissected out, then subjected to the tryptic digest, passed through capillary HPLC (or affinity-HPLC for fragments of posttranslationally modified proteins), and followed by injection into a mass spectrometer.

On a Wing and a Sprayer
Until the mid-1980s, molecular MS as applied to the biomed industry “was still only a fringe technique”, according to Fred W. McLafferty in his foreward to Interpreting Protein Mass Spectra. “To the total surprise of many of us, all of this changed a decade ago with the discovery of matrix-assisted laser desorption by Franz Hillenkamp and electrospray ionization (ESI) by John Fenn, increasing by orders of magnitude the molecular-weight range of MS. To me this has been an astounding revolution.” Of the two techniques, McLafferty considers ESI the best for analyzing impure samples because its “gentle ionization” minimizes concomitant fragmentation so that more representative ions of the mixture’s components are revealed.

Whether one is using a full-blown protein (alone or in mixture) or peptides derived from a trypsin or other enzyme digest, the principle behind using ESI-MS is the same. In ESI, an electrical field at the tip or outlet of a spray capillary imparts charge to the spray droplets with typical flow rates of between 1 and 100 mL per minute (or 1–100 nL/min for nanospray or microelectrospray ionization). Initially, larger droplets disperse into an ionized mist that feeds into the actual MS, usually by passing through a nitrogen curtain, which serves as a drying stream to remove solvent or water molecules from the charged protein or peptide.


Coming Out on TOF (MALDI, that is)
MALDI-MS uses a nitrogen UV laser (337 nm) on high-mass, nonvolatile samples such as proteins and peptides embedded in a matrix such as alpha-cyano-hydroxy cinnamic acid. In such a matrix, the proteins ionize rather than decompose and are passed down a flight tube into the mass spectrometer using a magnetic field. A detailed discussion of the use of MALDI-MS can be found at the Keck Foundation Biotechnology Resource Laboratory Web site (http://info.med.yale.edu/wmkeck/procmald.htm). Recently, MALDI quadrupole TOF-MS has been shown capable of identifying proteins from 2DE runs at the femtomole level (1). Because the MALDI technique generates gaseous ions from analytes deposited on surfaces, it has even been adapted for use directly with 2DE gels and membrane blots, the dried gel band or blot itself acting as the holding matrix.

Reference

  1. Shevchenko, A.; Loboda, A.; Schevchenko, A; Werner, E.; Standing, K. G. Anal. Chem. 2000, 72, 2132–2141.
Identification on Mass

Mass spectrometers analyze ions in the gaseous state. Therefore numerous methods are used to gasify and ionize sample compounds before MS analysis can be performed, including electrospray ionization (ESI) (see “On a Wing and a Sprayer” at right) and MALDI techniques (see “Coming out on TOF (MALDI that is)” below).

MS then separates the entering ions according to their m/z (the mass/charge ratio). The charge is generally due to the addition of variable numbers of protons to the proteins or peptide fragments.

Tandem mass spectrometers operate by using this separation of ions as a first fractionation step. Before entering the second mass spectrometer, ion fractions from the first are collisionally dissociated by passage through a neutral gas to induce fragmentation. These fragments exist as a family of subset ions of the original parent ion. Computerized analysis of the m/z spectrum of these subset ions can be used to determine the structure of the parent ion. In protein analysis, the parent ions are whole proteins or peptides, and the subsets consist of amino acid chains of varying length that have characteristic m/z fingerprints.

Fragments can be sequenced by automated inspection and comparison of the mass spectra obtained to the limited number of available m/z bands capable of being produced by relatively short combinations of the 20 amino acids. Although not all are unique combinations, cross-comparisons with other breakdown products can narrow possibilities, especially when trying to identify a protein with a known sequence.

Signature peptides. In some cases, a specific peptide fragment (or a few fragments in rare combination) proves unique to a particular type or class of protein in a species being studied. This fragment or combination of fragments can be used as “signature peptides” whose mass spec pattern can be used to instantly identify the presence of the particular protein in a trypsin digest of multiple unknown proteins, such as occurs in whole cell analysis (1). One limitation, of course, is that this approach can only work in systems in which a significant amount of background information on the proteome of the cell or species being studied is known. Still, in studies involving human physiological responses to drugs or diseases, the ability to rapidly inspect cell or plasma extracts for the up- or down-regulation of specific proteins or suites of known proteins should prove invaluable in the development of new therapeutics. As greater knowledge of the human proteome unfolds, so too will greater potential benefits reveal themselves.

Going Sweet on MS. One of the most complex problems involved in the analysis of proteins is their variable glycosylation. Posttranslational modifications, which add sugar side-chains to proteins, not only add an entirely different kind of chemistry to the analytical mix, they also create a potential nightmare of complications in what would otherwise be a relatively “simple” amino acid-based MS spectrum.

For example, the molecular weight of the amino acid chain of recombinant human tissue plasminogen activator (rt-PA) is 64 kD, but this is not counting the linked glycosylation groups. Trypsin digest yields at least 51 peptide fragments, including glycosylated forms. The ionization pattern as revealed in one ion-trap MS run of rt-PA is shown in the opening art. Although complex, the use of bioinformatics tools can make such MS displays readily analyzable.

Researchers at the Proteomic Division of ThermoFinnigan demonstrated one such method of identifying complex peptide mixtures and their posttranslational glycosylation patterns using a combination of trypic digests, and LC coupled to ion-trap MS, in this case for rt-PA. Figure 1 shows the tandem MS results on one of the selected ions from this first run that was determined through computerized analysis to be glycosylated.

Spectrometry Speciation
Even the most casual glance at the protein-related abstracts of the 49th Annual American Society of Mass Spectrometry conference of 2001 (see “Industry Facts and Figures”) shows the prominence of creativity and variety in the attempt to juxtapose technologies. Researchers eager to identify biologically significant proteins are adapting myriad forms of electrophoretic or chromatographic techniques for separation with every form of ESI and MALDI and ion trap technique known, and coupling them to TOF or FT-based MS of every description and setting. Such a rapid speciation of experimental approaches in separation and analysis (far more than even hinted at in this article) is a hallmark not of confusion, but of exuberant growth.

Doubtless certain paths will lead to evolutionary dead ends, and a few major technical combinations will come to dominate in specific application areas. But for now the field of protein analysis seems wide open and bounded only by the imagination of researchers and instrument makers. The only overwhelming aspect of the future is that it will include ever smaller sample sizes, ever faster and multidimensional runs, and ever more fully automated and computerized controls from setting up the initial samples to interpreting the final results. And what final results they will be—when the world of the proteome is fully revealed—to be used by medical science for drug discovery and design, and to be explicated by biologist–philosophers on what it is biochemically to be alive.

References

  1. Geng, M.; Li, J.; Regnier, F. J. Chromatogr. A, 2000, 8, 295–313.

Further Reading

  • Chalmers, M. J.; Gaskell, S. J. Curr. Opin. Biotechnol., 2000, 11, 384–390.
  • Kinter, M.; Sherman, N. E. Protein Sequencing and Identification Using Tandem Mass Spectrometry; Wiley-Interscience: New York, 2000.
  • Synder, A. P. Interpreting Protein Mass Spectra: A Comprehensive Resource; Oxford University Press: Washington, DC, 2000.


Mark S. Lesney is a senior editor of Today’s Chemist at Work. Send your comments or questions regarding this article to tcaw@acs.org or the Editorial Office 1155 16th St N.W., Washington, DC 20036.

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