Posttranslational modification of proteins is a fundamental regulatory mechanism, and characterization of protein modifications is paramount for understanding protein function. MS is one of the most powerful tools for the analysis of protein modifications because virtually any type of protein modification can be identified. Although we focus here on protein phosphorylation, the analysis of other types of protein modification by MS has been described (25). Protein phosphorylation is one of the most common of all protein modifications and has been found in nearly all cellular processes (74, 88, 153). MS can be used to identify novel phosphoproteins, measure changes in the phosphorylation state of proteins in response to an effector, and determine phosphorylation sites in proteins. Identification of phosphorylation sites can provide information about the mechanism of enzyme regulation and the protein kinases and phosphatases involved. A proteomics approach to protein phosphorylation has the advantage that instead of studying changes in the phosphorylation of a single protein in response to some perturbation, one can study all the phosphoproteins in a cell (the phosphoproteome) at the same time. A common approach to studying protein phosphorylation events is the use of in vivo labeling of phosphoproteins with inorganic ³²P. The phosphoproteomes of cells that differ in some way (e.g., normal versus diseased) can be analyzed by growing cells in inorganic ³²P and creating cell lysates. Changes in the phosphorylation state of proteins can then be examined by 2-DE and autoradiography. Proteins of interest are excised from the gel and microsequenced by MS. A major limitation of this approach is that while many phosphorylated proteins can be visualized by autoradiography, they cannot be identified because of their low abundance. One solution to this problem is enrichment of the phosphoproteome.

Phosphoprotein enrichment. Enrichment of the phosphoproteome of a cell can allow the identification of low-copy phosphoproteins that would otherwise go undetected. In one approach, phosphoproteins were enriched by conversion of phosphoserine residues to biotinylated residues (118). This method is an extension of techniques originally developed by Hielmeyer and colleagues (108) and more recently by our laboratory (51) for the identification of phosphorylation sites using Edman sequencing. Following derivatization, proteins that were formerly phosphorylated can be isolated by avidin affinity chromatography (118). Proteins immobilized on avidin beads can then be eluted with biotin, theoretically resulting in the isolation of the entire phosphoserine proteome (Fig. 13). By increasing the amount of cell lysate used for avidin affinity chromatography, low-abundance phosphoproteins can be enriched. However, this technique does not work for phosphotyrosine and the reactivity of phosphothreonine by this method is very poor (118). Tyrosine-phosphorylated proteins can be isolated by the use of antiphosphotyrosine antibodies (124). As an alternative, another method for phosphopeptide enrichment was devised to allow the recovery of proteins phosphorylated on serine, threonine, and tyrosine (179). In this method, a protein or mixture of proteins is digested to peptides with a protease and then subjected to a multistep procedure for the conversion of phosphoamino acids into free sulfhydryl groups. To capture the derivatized peptides, the free sulfhydryl groups in the peptides are then reacted with iodoacetyl groups immobilized on glass beads. Using this method, several phosphopeptides were recovered from ß-casein and from a yeast cell extract, although it was unclear whether all the proteins isolated from the yeast extract were phosphoproteins (179).

View larger version (31K): [in this window] [in a new window]   FIG. 13. Phosphopeptide and phosphoprotein enrichment. (A) Enrichment of phosphopeptides. Phosphoproteins are digested with a protease, and the phosphate groups are converted to biotin tags (119). Once biotinylated, the peptides can be selectively recovered with avidin-Sepharose and analyzed by MS. (B) Differential display of phosphoproteins. The phosphate groups present in proteins derived from two different samples are converted to biotin tags, and the phosphoproteins are purified on avidin-Sepharose in an identical manner. The phosphoproteins are then compared by 1- or 2-DE, and the target proteins are digested and analyzed by MS.

 
Enrichment of the phosphoproteome can also be combined with protein profiling by 1- or 2-DE. In this way, changes in protein amount observed on electrophoresis will reflect the level of protein phosphorylation (Fig. 13). Recently, the principle of protein quantitation by ICAT has been combined with phosphoprotein enrichment (60). This was accomplished by the introduction of isotopic label into ethanedithiol, the reagent used to convert the alkene created by ß-elimination of phosphoserine into a free sulfhydryl group. In this way, the differences in the amount of phosphoproteins in extracts can be analyzed quantitatively in the mass spectrometer (60). It should be noted that because of the chemistry used in both of these methods, these techniques are relatively insensitive and require tens of picomoles of phosphoprotein. As a result, we have found that these methods as currently designed are impractical for the isolation and enrichment of low-abundance phosphoproteins.

Phosphorylation site determination by Edman degradation. Edman sequencing is still a widely used method for determining phosphorylation sites in proteins labeled with ³²P, either in vitro or in vivo (5, 22, 164). This is because sites can be determined at the sub-femtomolar level if enough radioactivity can be incorporated into the phosphoprotein of interest. In our hands, this can be as little as 1,000 cpm (not ideal). Briefly, a ³²P-labeled protein is digested with a protease and the resulting phosphopeptides are separated and purified by reverse-phase HPLC or thin-layer chromatography (TLC) (Fig. 14). The isolated peptides are then cross-linked via their C termini to an inert membrane (e.g. Immobilon P; PerSeptive Biosystems). The radioactive membrane is subjected to several rounds of Edman cycles, and radioactivity is collected after the cleavage step. The released ³²P is counted in a scintillation counter. This method positionally places the phosphoamino acid within the sequenced phosphopeptide. Of course, this is meaningful only if the sequence of the phosphopeptide is already known. In addition, the analysis ceases to become quantitative beyond 30 Edman cycles (even with efficient, modern Edman machines) due to well-understood issues with repetitive yield associated with Edman chemistry.

View larger version (23K): [in this window] [in a new window]   FIG. 14. Strategies for determination of phosphorylation sites in proteins. Proteins phosphorylated in vitro or in vivo can be isolated by protein electrophoresis and analyzed by MS. (A) Identification of phosphopeptides by peptide mass fingerprinting. In this method, phosphopeptides are identified by comparing the mass spectrum of an untreated sample to that of a sample treated with phosphatase. In the phosphatase-treated sample, potential phosphopeptides are identified by a decrease in mass due to loss of a phosphate group (80 Da). (B) Phosphorylation sites can be identified by peptide sequencing using MS/MS. (C) Edman degradation can be used to monitor the release of inorganic 32P to provide information about phosphorylation sites in peptides.

 
Recently, our laboratory has extended the usefulness of phosphorylation site characterization by Edman chemistry through the development of the cleaved radioactive peptide (CRP) program (J. A. MacDonald, A. J. Mackay, W. R. Pearson, and T. A. J. Haystead, submitted for publication). In CRP analysis, one requires only that the sequence of the protein be known. Purification and sequencing of individual peptides is not required. Radiolabeled proteins (isolated following immunoprecipitation from ³²P-labeled cells, for example) are cleaved at predetermined residues by the action of a protease. The phosphopeptides are then separated by HPLC or TLC (if only one site is present, no peptide separation is required), cross-linked to the inert membrane, and carried through 25 to 30 Edman cycles. The sequence of the target protein is entered into the CRP program. This program predicts how many Edman cycles are required to cover 100% of all the serines, threonines, and tyrosines from the site of cleavage. Generally, one round of CRP analysis narrows the number of possible sites to 5 to 10 for most proteins. Phosphoamino acid analysis can be used to reduce the number of possibilities still further. The CRP analysis is then repeated following cleavage with a second protease (usually one cutting at R, but M and F are alternatives). The second round of CRP usually unambiguously localizes the phosphoamino acid to one possible site. The technique does not work if sites are more than 30 amino acids away from all possible cleavage sites. The finding that CRP analysis is not applicable may in itself confine a phosphorylation site to a segment of the protein that is likely to produce very large proteolytic fragments. The Cleavage of Radioactive Proteins (CRP) program is accessible at http://fasta.bioch.virginia.edu/crp/ and was written in collaboration with Aaron Mackey and Bill Pearson of the University of Virginia (MacDonald et al., submitted).

Phosphorylation site determination by mass spectrometry. Because of its sensitivity, MS can allow the direct sequencing of phosphopeptides, resulting in unambiguous phosphorylation site identification. Below, a brief overview of some common methods for phosphorylation site determination by MS are given. A more complete discussion of this topic is provided by Mitchelhill and Kemp (110). Identification of phosphorylation sites in proteins provides several unique challenges for the mass spectrometrist. For example, unlike in protein identification, where analysis of any peptide within the protein can be informative, phosphorylation site analysis requires that the phosphorylated peptide be analyzed. This means that considerably more protein is required for analysis. In addition, phosphorylation can alter the cleavage pattern of a protein and the resulting phosphopeptides may require different purification methods. To isolate and purify the phosphopeptides of interest, it may be necessary to alter the way in which the phosphoprotein is digested and to alter the pH or the chromatographic material used for peptide purification (27, 110, 116).

(i) Phosphopeptide sequencing by MS/MS. In our laboratory, we have found that a combination of HPLC, Edman degradation, and phosphopeptide sequencing by MS/MS provides the best results for phosphorylation site determination (Fig. 14). Following excision and digestion of a ³²P-labeled protein, the peptides are resolved by HPLC. By monitoring HPLC fractions for radioactivity, the phosphopeptides can be selected for analysis. This reduces the complexity of the peptide mixture before MS is performed and facilitates phosphopeptide identification (Fig. 14).

Phosphopeptides can be identified from a mixture of peptides by a method known as precursor ion scanning (116). In this method, the second mass analyzer in the mass spectrometer is set at the mass of the reporter ion for the phospho group (PO₃^-) of m/z = 79. Peptides are sprayed under neutral or basic conditions, and phosphopeptides are identified in the precursor ion scan only if their fragmentation yields an ion of m/z = 79. Once a phosphopeptide is identified, the peptide mixture is sprayed under acidic conditions and the phosphopeptide is sequenced by conventional tandem MS/MS. On fragmentation of the phosphopeptide, phosphoserine can be identified by the formation of dehydroalanine (69 Da), the ß-elimination product of phosphoserine. Similarly, phosphothreonine can be identified by the formation of its ß-elimination product, dehydroamino-2-butyric acid at 83 Da (116).

(ii) Analysis of phosphopeptides by MALDI-TOF. MALDI-TOF mass spectrometry can also be used to identify phosphopeptides (81, 130, 177, 178). When phosphorylated peptides are subjected to ionization by MALDI, phosphate groups are frequently liberated from the peptides. This is the case for phosphoserine- and phosphothreonine-containing peptides, which can liberate HPO₃ or H₃PO₄, resulting in a neutral loss of 80 and 98 Da, respectively. Careful examination of the TOF spectrum for differences in peptide masses of 80 Da that are not found in the unphosphorylated peptide control can identify phosphopeptides. Phosphopeptides can also be identified by treating one of two identical samples with protein phosphatase to liberate phosphate groups (Fig. 14). Once a phosphopeptide is identified, it can be sequenced by MS/MS for identification of the phosphorylation site (178).

One of the most exciting applications of proteomics involves combining this technology with the power of yeast genetics to delineate signaling events in vivo. Our laboratory has published two papers using this strategy to identify in vivo targets for protein phosphatases (9, 40). In one study (9), we identified physiological substrates for the Glc7p-Reg1p complex by examining the effects of deletion of the REG1 gene on the yeast phosphoproteome. In S. cerevisiae, PP-1 (Glc7p) and its binding protein, Reg1p, are essential for the regulation of glucose repression pathways. The target for this phosphatase complex was not known. Analysis by 2-D phosphoprotein mapping identified two distinct proteins that were greatly increased in phosphate content in reg1 mutants. Mixed-peptide sequencing identified these proteins as hexokinase II (Hxk2p) and the E1 subunit of pyruvate dehydrogenase. We then went on to validate these findings in a comprehensive biochemical study. Consistent with increased phosphorylation of Hxk2p in response to REG1 deletion, fractionation of yeast extracts by anion-exchange chromatography identified a Hxk2p phosphatase activity in wild-type strains that was selectively lost in the reg1 mutant. Having carried out these studies, we attempted to rescue the reg1 phosphoprotein phenotype by overexpressing both wild-type and mutant Reg1p in the deletion strains. Here, both the phosphorylation state of Hxk2p and Hxk2p phosphatase activity were restored to wild-type levels in the reg1 mutant by expression of a LexA-Reg1p fusion protein. In contrast, expression of a LexA-Reg1p protein containing mutations at phenylalanine in a putative PP-1C (the catalytic subunit) binding site motif (K/R)(X)(I/V)XF was unable to rescue Hxk2p dephosphorylation in intact yeast or restore Hxk2p phosphatase activity. These results demonstrate that Reg1p targets PP-1C to dephosphorylate Hxk2p in vivo and that the peptide motif (K/R)(X)(I/V)XF is necessary for its PP-1 targeting function. These studies therefore demonstrate how a proteomics approach can be used to first identify enzyme targets in cells and then direct all further analysis to verify the findings. It should be pointed out that often 6 to 12 months of work ensues following the initial sequencing of the targeted proteins. Nevertheless, clearly a combined proteomics and genetics approach greatly enhances one's ability to directly answer key biological questions. We believe that a similar strategy could be adopted with transgenic or knockout mouse work, particularly in cases where there is no obvious phenotype.

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