From the Faculty of Medical and Human Sciences, University of Manchester and the
Mass Spectrometry Laboratory and || Department of Cell Division, Paterson Institute for Cancer Research, Christie Hospital, Wilmslow Road, Manchester M20 9BX, United Kingdom
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ABSTRACT |
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Currently the most sensitive techniques for protein analysis rely on MS (4). Various MS protocols have been developed to identify sites of phosphorylation. However, the definition of phosphorylation events remains technically challenging due to both the low occurrence of these peptides and their intrinsic physicochemical nature. This results in generally poor ionization and detection using standard MS techniques. These can be moderated using peptide derivatization (e.g. ß-elimination and Michael addition) (5, 6). Derivatization using the same reactions offers specific protein cleavage dependent on phosphorylation (7). However, critical to the success of proteomic protocols is the simplicity and efficiency of these procedures prior to MS (8).
A further criterion for successful adaptation of phosphoproteomics protocols is sensitivity. Enrichment of phosphopeptides can be achieved via IMAC (911), graphite (12), and titanium oxide (13). These can be preceded by protein enrichment using phosphoamino acid-specific antibodies or phosphorylated consensus sequence-specific antibodies (14). Using highly enriched proteins current MS techniques can identify phosphorylation sites. These approaches include neutral loss scanning for the loss of a phosphate moiety (98 Da) from the parent ion (15) and precursor ion scanning either for an ion at m/z 79 in negative ion mode for Ser/Thr phosphorylation or an ion at m/z 216.043 for Tyr(P) in positive ion mode. Comparison of these techniques applied to gel-separated proteins yielded phosphorylation sites on peptides present at <200 fmol (16).
In cases where protein is highly abundant, precursor ion scan-based protocols have identified both novel and previously characterized phosphorylation sites (1719). Notably these protocols have primarily been used for unfocused approaches where a large number of phosphoproteins and peptides have been identified in a complex mixture such as a whole cell lysate.
Where protein phosphorylation events on a specific target protein of interest are sought, however, the sensitivity of current MS techniques often requires overexpression of that protein to achieve quantities commensurate with precursor ion scanning methods. This is of concern to biologists as the protein is expressed at abnormal levels and often in an abnormal cell type. A more relevant method is direct purification from the cell type of interest. However, in a typical study, Zappacosta et al. (16) demonstrated a requirement for 200 fmol of protein (2 x 108 cells for a protein present at 100 copies/cell, assuming complete recovery of the protein during cell lysis, immunoprecipitation, gel separation, and in-gel digestion). This quantity of biological material for the purification of a specific protein target is not always available. Therefore, there is a growing requirement for objective, specific, and more sensitive protocols for discovery of multiple phosphorylation sites on individual proteins generated by standard, relatively rapid biochemical techniques. Previous work using hypothesis-driven analysis of a known protein developed by Chang et al. (20) demonstrated the power of this approach to detect the presence of phosphorylation. However, the approach they describe is unable to determine tyrosine phosphorylation or other post-translational modifications and often assumes that kinases only phosphorylate known consensus sequences.
A novel hybrid quadrupole linear ion trap mass spectrometer with functionalities of both triple quadrupole and linear ion trap instruments is now available (21, 22). This combination offers a unique opportunity for studies on post-translational modifications.
We describe an alternative, information-dependent method for the identification of sites of phosphorylation on target proteins. This technique is specifically designed to comprehensively analyze a single target protein rather than as a general phosphorylation screening tool. Because in most cases the biologist has a specific target protein of interest, the added sensitivity of this type of focused analysis is a great advantage. The first part of each analysis uses the instrument as a highly sensitive triple quadrupole mass spectrometer to perform multiple reaction monitoring (MRM)1 analyses to screen for potential phosphopeptide signatures. The second phase of the experiment uses the third quadrupole as a linear ion trap for product ion detection to identify the peptide and the site of modification. This unique combination allows sensitive detection of phosphopeptides and identification of phosphorylation sites. We demonstrate below how this relatively simple approach can enhance the development of phosphoproteomics for the biologist.
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EXPERIMENTAL PROCEDURES |
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The six-protein mixture sample consisting of bovine serum albumin, alcohol dehydrogenase, apotransferrin, ß-galactosidase, lysozyme, and cytochrome c was diluted from a 100 pmol lyophilized protein mixture digest (LC Packings, Amsterdam, The Netherlands) using 2% (v/v) acetonitrile, 0.1% (v/v) formic acid.
The cdc13+ gene (23) was cloned into the NdeI site of pREP41PkC (29) and introduced into cdc25.22 leu1.32 cells. Expression was induced by culturing at 25 °C for 31.5 h when the level of Cdc13-Pk equaled that of the native Cdc13 protein. The monoclonal antibody MAb336 was used to enrich for Cdc13 and associated proteins (24). Interphase (G2 arrest) samples were generated by inducing cell cycle arrest through an increase in culture temperature from 25 to 36 °C. Cells were harvested 30 min after this culture had been returned to the permissive temperature of 25 °C.
Enriched fractions were separated by SDS-PAGE, and gels were stained with colloidal Coomassie Blue (www.lrf.umist.ac.uk). In-gel digests were performed as described previously (25).
Liquid Chromatography and Mass Spectrometry
For each experiment, 5 µl of sample were loaded onto a 15-cm x 75-µm inner diameter PepMap C18 3-µm column (LC Packings) using a standard LC Packings UltiMate pump and FAMOS autosampler. Samples were desalted on line prior to separation using a microprecolumn (5-mm % 300-µm inner diameter) cartridge. The washing solvent was 0.1% formic acid delivered at a flow rate of 30 µl min1 for 3 min. Peptides were separated using a solvent gradient determined by the complexity of the sample mixture. Details of the gradients used for each sample is given in Supplemental Methods, Section 1.
Chromatography was performed on line to a 4000 Q-TRAP mass spectrometer (Applied Biosystems, Framingham, MA). All voltages and gas settings used are described in Supplemental Methods, Section 2.
Cdc2, Cyclin B, and Hsp60 were initially identified from 10% of the digested sample by mass spectrometry using a QSTAR XL (Applied Biosystems) as described in Unwin et al. (26). Briefly the mass spectrometer was operated in information-dependent acquisition mode, which involves switching from MS to MS/MS mode on detection of doubly and triply charged species above a preset threshold. Data obtained are then combined and submitted to a protein database such as Swiss-Prot via Mascot (27) for protein identification.
MRM-initiated Detection and Sequencing (MIDAS)
For selective detection of phosphopeptides, a list of MRM transitions of potential phosphopeptides was generated either by manual calculation or by a software script developed by Applied Biosystems. In general, transitions were included for all tryptic peptides (maximum one missed cleavage) containing Ser, Thr, or Tyr residues with either one or two modifications and for doubly and triply charged species for the Q1 mass range 4001600 m/z. The number of MRMs is dependent on various factors, including protein size, number of peptides following digestion, and the number of potential phosphorylation sites. It is, however, important to optimize the cycle time to around or below 10 s. Such a cycle time ensures that if the peak width is around 0.5 min it is highly probable that a peptide is scanned for and analyzed at least twice as it is eluted and that one of these analyses will occur at, or close to, the apex of its elution profile.
A large list of MRMs may be halved, and two separate analyses can be performed. Clearly other modifications can be included in the experiment such as oxidation of methionines or deamidation. Although these should be minimized during sample preparation, using a small proportion of sample in a standard identification experiment should quickly determine whether these modifications are present and whether they should be included in the list of MRM transitions.
The number of MRM transitions, dwell times, and cycle times used for each protein described in the study are provided in Supplemental Methods, Section 3. Collision energies (CE) for Ser(P) and Thr(P) peptides were calculated according to the equation CE = (m/z x 0.045) 6.2 for doubly charged peptides and CE = (m/z x 0.026) + 5.2 for triply charged peptides. CE for Tyr(P) peptides is set at 60eV.
The mass spectrometer was instructed to switch from MRM to enhanced product ion scanning mode when an individual MRM signal exceeded 100 counts. Each precursor was fragmented a maximum of twice before being excluded for 2 min. Enhanced product ion scanning was performed using parameters detailed in Supplemental Methods, Section 2. Data were initially analyzed by submitting the MS/MS data to Mascot with a peptide tolerance setting of ±0.02 Da. This proves successful as the exact parent mass is known and defined in the MRM transition. Hence the search can be performed using the actual mass of the peptide of interest rather than a less accurate mass generated by the instrument. As such, few false positive database "hits" are generated. All Mascot searches were performed against a species-specific NCBInr database with trypsin plus one missed cleavage, phosphorylation as a variable modification, and an MS/MS tolerance of 0.5 Da. In addition, data were examined manually using the Applied Biosystems BioExplore feature of the Analyst 1.4 software package.
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RESULTS |
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We exploited these features to scan for phosphopeptides and termed this approach MIDAS. The overall scheme of the method is outlined in Fig. 1a. Initially a candidate list of MRM transitions is generated. The masses of theoretical phosphopeptides are calculated by addition of 80 to the peptide mass for every "phosphorylatable" residue present. The Q1 mass is calculated for this peptide in doubly/triply charged form with up to two phosphate additions. The Q3 mass is calculated as either loss of 98 Da for each putative Ser(P) or Thr(P) or generation of a 216.0 ion for tyrosine-containing peptides. An example is shown in Table I. A dwell time and collision energy are then added.
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Assessment of MIDAS Sensitivity Using Standard Proteins
The sensitivity and selectivity of this method were assessed using a model phosphoprotein, -casein.
-Casein was selected to allow comparison with existing phosphorylation analysis methods (19, 28) where limits of detection are published.
A MIDAS experiment comprising 65 MRMs to identify -casein S1 phosphopeptides is presented in Fig. 2. 10 fmol of
-casein spiked with 4 fmol of [Glu]fibrinopeptide B (GluFib), used as a positive control, were loaded onto the column. The addition of GluFib was important at this stage as a control when dilution of
-casein below the limit of detection was performed.
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Using MIDAS, the peptide sequence and number of phosphorylation sites are inferred from the MRM transition trigger. Thus, location of phosphorylation site(s) may require few fragment ions in certain cases. Because the sensitivity of MRM is much higher than for product ion scanning, the sensitivity of phosphopeptide and phosphorylation site detection is improved. Fig. 3, which depicts data from MIDAS analysis of either -casein S1 or S2, illustrates this point. Fig. 3, a and b, shows product ion spectra that define phosphorylation sites on
-casein S1 and confirm phosphorylation of an
-casein S2 peptide when loaded onto the column at 10 fmol of
-casein (a mixture of S1 and S2). Even when the column loading was reduced to 5 fmol, the
-casein S1 (trypsin) missed cleavage phosphopeptide YKVPQLEIVPNpSAEER is identified (Fig. 3c). Fewer y ions were generated, but there remains sufficient information to confirm the site of phosphorylation. For the
-casein S2 peptide (Fig. 3b), the spectrum confirms peptide identity. Phosphorylation was implied from the precursor mass and positive MRM, and the spectrum shows modification on either Ser-6 or Thr-7. A more focused MIDAS analysis to detect only this peptide with a longer acquisition of product ion spectra demonstrates phosphorylation on Ser-6 (see Supplemental Fig. 1) showing the utility of careful MRM experiment design. Two peptides were detectable that contain the same phosphorylated serine due to inefficient tryptic cleavage. Therefore we can characterize phosphopeptides in the low femtomole region, with and without missed cleavage, by MIDAS.
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DISCUSSION |
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In theory and in practice our protocol generated few false positives. Compared with established phosphate scanning methods, definition of a precursor mass equivalent to that of a phosphopeptide vastly reduces false positive results due to generation of b2 ions (216 Da) interfering in a phosphotyrosine peptide analysis. In the cases of phosphoserine and phosphothreonine neutral loss analysis, false positives resulting from generation of a peptide fragment of "diagnostic" m/z or loss of valine (99 Da) are diminished. In addition, because the accurate precursor mass is known, database searching can be performed with very tight tolerances. This specificity is especially important if the protein is present in a mixture.
Even with 20 fmol of phosphoprotein in a mixture of other proteins at 500 fmol (Fig. 4), no false positives were generated. This sample, prepared specifically to test the specificity of the MRM scanning approach, also allowed direct comparison to published methods (28). Because this 20 fmol is a mixture of two -casein proteins and the phosphopeptides found in this experiment are a missed cleavage around the same phosphorylation site, we are effectively detecting peptides present at significantly less than the 20 fmol of total protein loaded onto the column. The sensitivity of this method represents a significant improvement of approximately an order of magnitude compared with analysis of a similar sample on this instrument using precursor ion scanning, which detected
-casein phosphopeptides in a protein mixture at 75 fmol (28). This type of sample represents an extreme "worst case" scenario where the biologist has very high levels of contaminating protein, yet MIDAS is still capable of identifying phosphorylation sites on the target protein. MRM analysis therefore provides a more sensitive and specific scanning method for phosphopeptides from a known target protein than existing methodologies.
We confirmed the effectiveness of the method for biochemical samples prepared in a standard format. Phosphorylation sites on proteins from Schizosaccharomyces pombe isolated by immunoprecipitation and one-dimensional gel electrophoresis were taken for successful identification of a novel site of serine phosphorylation on mitotic Cyclin B (argued to be a completely phosphor-mapped protein) and two previously observed phosphorylation sites in a Cyclin B-interacting protein, Cdc2. In addition, a novel serine phosphorylation site was identified on Hsp60. These data demonstrate that MIDAS can be used to obtain information regarding novel phosphorylation sites using quantities of protein that are feasibly obtained from real biological model systems.
Creating the MRM experiment peptide list is relatively simple and can be tailored to search for specific predicted phosphorylation sites, at even greater sensitivity, by increasing the dwell time or the number of MS/MS scans performed on a specific peptide precursor. For a protein that generates a large number of MRMs, the experiment is easily fractionated keeping the cycle time below 10 s. Because we did not use the entire sample for the identification of phosphopeptides in our yeast protein samples, this is a real solution to this problem. It should be mentioned, however, that an experiment comprising 172 MRMs was performed and successfully identified sites of Cyclin B phosphorylation. It is important to note that for the samples analyzed here not all potential phosphorylation sites and charge states were considered; rather a degree of intelligent filtering was used to limit the number of MRMs to a reasonable level. The reasoning behind this filtering is described in the supplemental methods for each protein. For a complete screen of multiple sites and charge states the MRM transition list often becomes large and requires splitting between multiple injections. For example, Cdc2 would require 374 MRM transitions for both +2 and +3 charge states with Ser/Thr/Tyr being potentially phosphorylated (maximum of three per peptide), assuming one missed cleavage and possible methionine oxidation. This would require four injections of the sample. If zero missed cleavages are considered this is reduced to one injection of 124 MRMs. The level of missed cleavages and methionine oxidation can be assessed in a standard protein identification experiment on the sample and can therefore be included or excluded from the MRM analysis as required.
Clearly MIDAS is not a panacea. The intrinsic biochemical nature of phosphopeptides that causes their adherence to surfaces during LC-MS/MS or their poor ionization is not addressed in this study. However, further experiments such as phosphopeptide enrichment by IMAC or strategies to improve ionization efficiency of phosphopeptides will increase the sensitivity of this approach further. Use of other proteases will also improve protein coverage and may also improve ionization of peptides containing phosphorylated residues. Where sufficient sample is available, several techniques should be combined to fully characterize all sites of phosphorylation on a protein of interest. However, given the improvement in sensitivity and selectivity that MIDAS offers over other LC-MS/MS-based methodologies, it is clear that this method is successful where small amounts of protein are available.
None of the S. pombe proteins analyzed were overexpressed, making MIDAS a highly valuable tool in rapidly assessing phosphorylation in a target protein of interest without the need for lengthy cloning, overexpression, and purification techniques. Furthermore MIDAS is also applicable to the analysis of other key post-translational modifications such as acetylation (results not shown). Given the semiquantitative nature of MRM, MIDAS can also theoretically be extended to perform relative quantitation of modified peptides in a protein from different samples without the need for radioactivity, stable isotopes, or the production of modification-specific antibodies. Therefore, MIDAS provides a sensitive and flexible approach to protein post-translational modification identification and analysis.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Published, MCP Papers in Press, May 27, 2005, DOI 10.1074/mcp.M500113-MCP200
1 The abbreviations used are: MRM, multiple reaction monitoring; MIDAS, MRM-initiated detection and sequencing; TIC, total ion chromatograph; XIC, extracted ion chromatograph; CE, collision energy; GluFib, [Glu]fibrinopeptide B.
* This work was supported by the Leukemia Research Fund, UK; Cancer Research UK; and Biotechnology and Biological Sciences Research Council. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
S The on-line version of this article (available at http://www.mcponline.org) contains supplemental material.
¶ Both authors contributed equally to this work.
** To whom correspondence should be addressed. Tel.: 44-0-161-446-8247; Fax: 44-0-161-446-3109; E-mail: Tony.Whetton{at}Manchester.ac.uk
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REFERENCES |
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