From the Max Planck Institute of Molecular Plant Physiology, 14476 Potsdam-Golm, Germany
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ABSTRACT |
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Surprisingly the Arabidopsis thaliana genome contains numerous TPS and trehalose-phosphate phosphatase homologues (8). The expression of TPS genes occurs in various tissues and is controlled by numerous stimuli such as abiotic stress, nutritional stress, and circadian cycle (9, 10). The TPS gene At2g18700 is strongly induced during extended night conditions (low sugar levels) (9). Four other genes coding for TPS (At1g60140, At1g68020, At1g70290, and At1g23870) also show induction (9). Furthermore the Arabidopsis TPS1 (At1g78580) mutant, disrupted in the trehalose-6-phosphate synthase gene, has a pronounced phenotype that could be interpreted as the consequence of glycolytic deregulation (8, 11). Recent transcriptomic analysis further revealed gene clusters that correlated with alterations in trehalose 6-phosphate levels (7). Among the genes identified, one was repressed by sucrose addition (7). This gene, AtKIN11, encodes a sucrose non-fermenting 1-related protein kinase (SnRK1) known to be involved in signal transduction affecting sugar utilization (13).
In a manner analogous to the biosynthetic key enzymes sucrose-phosphate synthase (SPS) and nitrate reductase, whose activity states are rapidly altered by protein kinase action in response to environmental changes and/or genetic perturbations present in mutant plants (1417), the TPS enzyme also is thought to be regulated by reversible protein phosphorylation/dephosphorylation (18). The reversible phosphorylation of proteins is recognized as a common and important post-translational modification in eukaryotes. It alters the behavior of proteins and is well known to play a key regulatory role in cellular processes such as plant defense (19, 20), plant growth (21), energy metabolism (22), and stress responses (23, 24). It is estimated that 30% of all proteins expressed in eukaryotic cells exist in phosphorylated forms at any given time (2527). In Arabidopsis, the regulation of these highly dynamic phosphorylation states is achieved by
1100 protein kinases and 100200 protein phosphatases (28, 29).
The Ca2+-independent SnRK1 subgroup is involved in signaling pathways controlling fundamental cellular processes. SnRK1 is the closest plant homologue of yeast sucrose non-fermenting-1 protein kinase and of the AMP-activated protein kinase in animals. The identification of SnRK1 substrates like SPS and nitrate reductase (14, 3032) demonstrated the roles of these kinases in regulating biosynthetic pathways such as sucrose synthesis and nitrogen assimilation. Furthermore SnRK1 is involved in plant growth (33) and was shown to play a role in plant development (34). In Arabidopsis there are three known SnRK1s (35), one of which is thought to be subject to regulation by glucose 6-phosphate (36). Moreover Arabidopsis TPSs possess putative SnRK1 phosphorylation sites.
MS has become the method of choice for profiling post-translational modifications, in particular reversible phosphorylation. Such applications help us gain insight into protein regulation at the cellular level. For phosphopeptide analysis, characteristic marker ions at m/z 63 (PO2) and/or 79 (PO3) are used in precursor ion scan mode (3739). Furthermore phosphopeptide-specific neutral losses of H3PO4 (98 Da) or HPO3 (80 Da) are utilized for identification in neutral loss MS experiments (40, 41). We recently developed a robust non-targeted and stable isotope-based method for the routine quantification of phosphorylation at specific sites (42). The method allows the simultaneous identification and quantification of a multitude of in vitro phosphorylated synthetic peptides. We have observed clear differences in the phosphorylation states of Arabidopsis wild-type and mutant plant SPS (42). Although it is a very homologous system using substrates similar to those used in sucrose biosynthesis, no investigations on the regulation of TPS isoforms via post-translational phosphorylation have been reported.
As a complement to the non-targeted approach, we developed a stable isotope-free method for the highly selective detection and absolute quantification of target phosphopeptides using LC coupled to tandem mass spectrometry in MRM mode. Although phosphopeptides are generally not observed as intense peaks due to ionic suppression, especially in the presence of non-phosphorylated peptides (43, 44), our MRM-based method allows a significant enhancement of signal-to-noise (S/N) ratios in phosphopeptide analysis and compensates for the low ionization efficiency of phosphopeptides in positive ionization mode by the high selectivity of the mass spectrometer using MRM. This enabled us to look at TPS isoform phosphorylation states in Arabidopsis for the first time
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EXPERIMENTAL PROCEDURES |
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Extraction of Protein Kinases from Arabidopsis Leaves
Frozen Arabidopsis wild-type leaf tissue (usually 300 mg fresh weight) was ground in a chilled mortar. After addition of 400 µl of extraction buffer containing 50 mM Tris-HCl, pH 7.5, 20% (v/v) glycerol, 5 mM dithiothreitol, 1 mM benzamidine, 0.3 µM microcystin, 60 mM sodium fluoride, 2 mM 4-(2-aminoethyl)benzenesulfonyl fluoride, 100 µM bestatin, 20 µM pepstatin A, 20 µM leupeptin, 30 µM (2S,3S)-3-(N-{(S)-1-[N-(4-guanidinobutyl)carbamoyl]3-methylbutyl}carbamoyl)oxirane-2-carboxylic acid, and 5 mM 1,10-phenanthroline, the crude extract was placed on ice for 20 min and was then centrifuged at 15,000 x g for 5 min. The supernatant was subsequently desalted on a Sephadex G-25 column (1 x 5 cm) previously equilibrated with extraction buffer containing no glycerol. The volume of eluent from the Sephadex G-25 column was 1 ml.
Determination of Protein Content
Protein concentrations were determined via the dye-binding method of Bradford as described previously (45) using bovine serum albumin as a standard. Each measurement was made in triplicate, and the mean values were used. 300 mg of Arabidopsis wild-type leaf tissue (fresh weight) used in each assay yielded 3 mg of total protein.
In Vitro Kinase Activity Assay
Typically each 100-µl kinase assay reaction mixture contained 50 µl of desalted Arabidopsis crude extract, 25 µM synthetic TPS peptide, 10 mM MgCl2, 5 mM ATP, and 1 mM CaCl2. Where specified, CaCl2 was replaced by 4 mM EGTA for the Ca2+-independent kinase activity assay. The reaction was initiated by the addition of desalted Arabidopsis crude extract. Following a 15-min incubation at 25 °C, the reaction was stopped by adding 500 µl of ice-cold EtOH. After standing on ice for 20 min, the mixture was centrifuged at 15,000 x g for 2 min. The supernatant was then evaporated in a centrifugal vacuum system (SpeedVac), reconstituted in 10 µl of 0.1% formic acid (FA) in water, and finally injected into the LC-MS/MS system.
Preparation of Calibration Curves with Phosphorylated TPS Peptide Standards
1 mM stock solutions of synthetic phosphorylated TPS peptides were prepared in 50 mM Tris-HCl, pH 7.5. These stock solutions were diluted further to make working solutions and stored at 20 °C. Aliquots of working solutions were spiked into kinase activity assay reaction mixtures instead of their non-phosphorylated counterparts. Following incubation and sample preparation as described above, the reconstituted TPS phosphopeptides were used for the calibration curves. The construction of calibration curves with phosphopeptide standards was done on a daily basis.
Liquid Chromatography
The HPLC system consisted of a Surveyor autosampler and Surveyor MS pump with an integrated degasser (ThermoFinnigan, San Jose, CA). The chromatographic separation of phosphopeptides was performed on a 100 x 1-mm, 3-µm, Luna C18 (2) column (Phenomenex, Aschaffenburg, Germany) that was directly interfaced to the ESI source of the mass spectrometer. Mobile phase A (0.1% FA in Milli-Q water (v/v)) and mobile phase B (0.1% FA in acetonitrile (v/v)) were used for elution. The gradient transitioned from 0 to 40% phase B during the first 25 min followed by 4080% phase B over a 1-min period. It was held for 10 min at 80% phase B and then transitioned back to 0% phase B over a 2-min period with a flow rate of 40 µl/min. Re-equilibration was performed at 0% B for 15 min with a flow rate of 60 µl/min. The injection volumes were 10 µl.
Mass Spectrometry
Mass spectrometry was performed on a TSQ Quantum Discovery MAX mass spectrometer (ThermoFinnigan) equipped with an Ion Max ESI source with a 34-gauge metal needle operated under XcaliburTM software (version 1.4 SR1, ThermoFinnigan) in the positive ion mode. The collision cell was pressurized with argon. Mass spectrometric detection of positively charged phosphopeptides was achieved with MRM for which the following tune parameters were set: sheath gas pressure of 30 arbitrary units; spray voltage set to 3.7 kV; temperature of the heated transfer capillary, 270 °C; collision gas pressure, 1.5 millitorrs. The scan width for all MRMs was 0.7 mass units. The resolution for Q1 was 0.3 mass units; the resolution for Q3 was set to 0.7 mass units. The collision energies used for the recorded transitions are shown in Table I. The dwell time per transition was 50 ms. To demonstrate the impact of the Q1 resolution setting on peak purity and selectivity of the detection the mass spectrometer was set at Q1 resolutions of 0.7, 0.5, and 0.3 Da full width at half-maximum (FHWM), respectively, and at a Q3 resolution of 0.7 Da FHWM.
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For the identification of phosphorylation sites MS3 spectra were recorded on an LTQ linear ion trap mass spectrometer (ThermoFinnigan). The isolation width was 2.0 mass units, the activation was set to 0.25, and the activation time was 30 ms. TPS phosphopeptides and phosphorylation sites were identified by automatic data-dependent acquisition consisting of a MS2 of the known precursor ions (TPS phosphopeptides) and a subsequent MS3 scan on the neutral loss fragments (m/z = 49, H3PO4 for the doubly charged ions).
Quantification
For the quantification of TPS phosphopeptides formed in the in vitro kinase assay, the two most abundant MRM transitions for each phosphopeptide (Table I) were used based on their observed fragmentation behavior. Subsequent processing of product ion chromatograms and peak integration was performed using the ICIS algorithm in LCquanTM 2.0 (Xcalibur software, version 1.4 SR1, ThermoFinnigan). Absolute amounts of phosphopeptides were calculated from the slope of peak area ratios of the in vitro formed phosphopeptides and calibration standards using the least square linear regression (weighting factor 1/concentration).
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RESULTS |
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Selectivity of Detection
The selective detection and identification of target TPS phosphopeptides formed in the multiparallel in vitro kinase assay were achieved by collision-induced dissociation, which allows the MS/MS system to be operated in the positive ionization MRM mode. By applying the optimum tuning parameters for each individual selected precursor/product ion transition, the two most abundant fragments (product ions) of each TPS phosphopeptide were used for quantification, thus verifying the selectivity of the method. As a result, identical amounts of phosphorylated analytes were obtained using calibration curves with both product ions. The phosphopeptide level ratios determined from the two MRM transitions were close to unity and proved stable for the calibration standards and the plant samples indicating that the approach was selective for the detection and quantification of TPS phosphopeptides formed in the multiparallel in vitro kinase activity assay.
A comparison of the MRM-based LC-MS/MS strategy with full-scan detection (in the mass range m/z 300900) is shown in Fig. 1. As can be seen, the S/N ratios for the in vitro formed TPS phosphopeptides detected in MRM mode are considerably higher than those observed using full-scan detection, demonstrating the high selectivity and specificity of the MRM-based method (Fig. 1 and Table I). The S/N gains ranged from 3.2 to 8.0. In Fig. 1, B and C, the extracted ion chromatograms for the phosphorylated TPS peptides 1 and 8 are shown for the mass spectrometric detection using the MRM mode and full-scan detection, respectively.
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Enhanced Q1 Resolution and Peak Purity in MRM
To test and compare the sensitivity and selectivity of the mass spectrometer under typical conditions for quantitative investigations, the TPS phosphopeptides were injected on-column and analyzed in MRM mode using three different resolution settings for Q1 (0.7, 0.5, and 0.3 Da FHWM) and fixed Q3 resolution (0.7 Da FHWM). As an example, Fig. 2 shows the MRM chromatograms of phospho-TPS peptide 8 obtained after chromatographic separation. As can be seen, enhanced resolution (Q1 = 0.3 Da FHWM, Q3 = 0.7 Da FHWM) is a valuable tool to eliminate interfering substances having the same nominal mass as the target phosphopeptide and background noise. Under enhanced resolution, the peak area of the target phosphopeptide was 5-fold lower than that observed at Q1 resolution 0.7 Da FHWM (Fig. 2). The S/N ratio, however, only decreased by a factor of 2. The ratio of S/N and peak area (S/N:peak area) was used as an indicator of peak purity. An increasing ratio signifies higher peak purity. Increasing the resolution of Q1 from 0.7 to 0.3 Da FHWM resulted in an increase of (S/N:peak area) from 0.001 to 0.0024. Thus, the peak purity significantly increased under enhanced resolution making the detection considerably more selective. Changing Q3 resolution from 0.7 to 0.5 Da FHWM under enhanced resolution did not further improve the selectivity of the measurement (data not shown). This indicates the predominant role of the Q1 resolution setting (rather than Q3) in peak purity and therefore selectivity of the mass spectrometric detection in MRM mode. To avoid interference peaks that can hamper the quantitative analysis of phosphorylation levels and therefore to achieve high peak purity, the mass spectrometer was always operated under enhanced resolution.
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Linearity of Quantification
To quantify phosphorylation reliably, method linearity validation is essential. To this end, calibration curves with synthetic standard TPS phosphopeptides were constructed, and the linearity of the method from 0.5500 pmol for all TPS phosphopeptides was investigated. Each measurement was done in triplicate. Using the least square linear regression (weighting factor 1/concentration) the coefficients of regression (r2) were determined to be 0.98830.9964 confirming a linear dependence of the amounts of TPS phosphopeptides and peak areas of the corresponding MRM transition. In Fig. 3 the calibration curves for two phosphorylated TPS peptides, p-TPS1 and p-TPS8, are depicted as examples.
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In Vitro Phosphorylation of TPS Peptides: Ca2+ Dependence and Independence
After establishing the described MS-based kinase assay system, we started to investigate the in vitro phosphorylation of TPS homologues in Arabidopsis. Beyond verifying TPS protein phosphorylation, three other questions have been central. (i) Based on transcriptomic analyses a Ca2+-independent kinase is known to cluster with trehalose accumulation (see the Introduction). Consequently we investigated the Ca2+ dependence of protein phosphorylation. (ii) We analyzed further whether the different TPS isozymes in Arabidopsis are differentially phosphorylated and (iii) to what extent multisite phosphorylation occurs for different TPS protein isozymes. To address these questions we first needed to determine whether the method is sensitive and selective enough to see differences among the phosphorylation levels of substrate TPS peptides.
The limits of detection (LODs) for all TPS phosphopeptides were determined by spiking matrix (Arabidopsis crude extract) with phosphorylated TPS standard peptides of known concentration and subsequently subjecting the mixture to LC-MS/MS analysis with MRM as described above. The S/N ratios were calculated from the ratio between analyte peak signal to base-line noise. The estimated LODs at 3 times S/N are shown in Table I.
Target peptide sequences were obtained by aligning the sequences of six different Arabidopsis TPS isozymes with the consensus sequence and alternatives for recognition of the SnRK1 subfamily (Fig. 4A) (47). The corresponding TPS genes At2g18700, At1g60140, At1g68020, At1g70290, and At1g23870 show (strong) induction under extended night conditions (9), and the Arabidopsis TPS1 (At1g78580) mutant, disrupted in the trehalose-6-phosphate synthase gene, exhibits a pronounced phenotype (8, 11).
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In a control experiment, ATP was omitted in the reaction mixture. Here apparent in vitro phosphorylation of TPS peptides most likely resulted from remaining endogenous (kinase-bound) ATP in the desalted crude extract. This "background phosphorylation" was subtracted from the amounts of phosphopeptides formed in the kinase activity assay upon addition of exogenous ATP. The absolute quantities of TPS phosphopeptides formed (taking into account the combined quantitative data obtained from the two most abundant MRM transitions for each TPS phosphopeptide) corresponding to extracted in vitro kinase activities are shown in Fig. 5. Reproducibility of absolute quantification of several independent replicates was 20% mean coefficient of variance. This level of precision is useful for most biological problems.
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Furthermore the in vitro phosphorylation of six TPS peptides (1, 4, 7, 9, 11, and 15) corresponding to the TPS isozymes At2g18700, At1g70290, At1g23870, At1g60140, and At1g68020 exhibited a (partially) strong Ca2+ dependence (Fig. 5 and Table I). When CaCl2 in the reaction mixture was replaced with 4 mM EGTA, the formation of the corresponding phospho-TPS peptides was significantly reduced. This indicates that Ca2+-dependent kinases in the crude extract also recognize these peptides as substrates. As can be seen, Ca2+-dependent kinases represent the dominant phosphorylating activity; Ca2+-independent kinase(s) phosphorylated a smaller portion of TPS peptides. In this context, the Ca2+ dependence of phosphopeptide formation is particularly pronounced for TPS peptides 7, 11, and 15 (TPS isozymes At1g70290, At1g23870, At1g60140, and At1g68020) where the formation of the corresponding phosphopeptides is reduced by 100% in the absence of CaCl2.
Identification of Phosphorylation Sites
The determination of the phosphorylated sites of the TPS peptides was done by the generation of neutral loss-driven MS3 spectra using a linear ion trap mass spectrometer (see "Experimental Procedures"). As a result of MS3 sufficient peptide backbone fragmentation of the TPS phosphopeptides was achieved to identify the correct phosphorylation sites. It could be shown that the full MS3 spectra of standard TPS phosphopeptides were identical to the full MS3 spectra obtained from in vitro formed TPS phosphopeptides thus providing evidence that the phosphorylated site of TPS peptides phosphorylated in the in vitro kinase assay was indeed the SnRK1 phosphorylation site (Fig. 6).
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DISCUSSION |
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Although absolute signal intensity and peak area obtained with this enhanced resolution were 5-fold lower compared with Q1 and Q3 0.7 Da FWHM (Fig. 2), the selectivity of detection was greatly increased. This is a clear advantage in eliminating interfering matrix compounds. At enhanced resolution, only the desired precursor ions pass through Q1. The resulting MRM mass chromatograms contain peaks of higher purity due to the exclusion of co-eluting ions that have the same nominal mass as the target peptides and that yield product ions of the nominal same mass as the target peptides. Thus, the resolution setting of Q1 has a much higher impact on peak purity than that of Q3.
Using this strategy, we have provided the first evidence for in vitro phosphorylation of TPS gene family peptides. This data makes possible the selective screening of upstream kinases potentially involved in TPS regulation via phosphorylation in vivo. The targeted approach described here complements our recently developed non-targeted stable isotope labeling strategy for the relative quantification of phosphopeptides in different biological samples. The use of the MRM mode for the mass spectrometric analysis allows highly selective detection and quantification of target phosphopeptides, whereas differential labeling allows identification of novel phosphorylation sites via de novo sequencing. The low ionization efficiency of phosphopeptides in positive ionization mode often hampers mass spectrometric analysis of phosphopeptides, resulting in rather low signal intensities, especially in the presence of non-phosphorylated peptides. A major advantage of our LC-MS/MS-based strategy is that it overcomes this drawback by the high selectivity of the triple quadrupole instrument in the MRM mode.
Studies on the recovery of phosphorylated reaction products of in vitro incubations have revealed enormous differences among TPS phosphopeptides, making quantification of phosphorylation levels a difficult task. Low phosphopeptide recoveries are probably due to nonspecific binding of in vitro formed phosphopeptides to reaction mixture proteins. The addition of EtOH to the reaction mixture results in the precipitation of proteins thus terminating the reaction. However, protein-bound TPS phosphopeptides co-precipitate, thereby reducing individual recoveries. Therefore, to absolutely quantify phosphopeptides the construction of calibration curves including the individual recoveries of TPS phosphopeptides is essential. We achieved this by substituting phosphorylated TPS calibration standards for TPS substrate peptides in the in vitro kinase activity assay reaction mixture, thus subjecting them to all steps of sample incubation and preparation that potentially diminish their recovery. This calibration procedure compensated for the different phosphopeptide recoveries and therefore allowed absolute quantification of TPS phosphopeptides formed during in vitro incubations.
As can be seen in Fig. 5 and Table I, the multiparallel kinase activity assay in the presence of synthetic TPS peptides clearly discriminates among phosphorylation specificities, thus providing evidence for different kinase substrate affinities. (Such discrimination may be particularly useful for experiments involving multiple types of factors like time points, treatments, and tissue types and for biological replicates.) TPS peptides 2, 12, 13, and 15 exhibit a proline residue at position 1 (adjacent to the target serine/threonine). Except for TPS peptide 15, which acted as a kinase target (formation of about 2 pmol of corresponding phospho-TPS peptide), the in vitro incubation of the other three TPS peptides yielded none or only very low amounts (below 0.5 pmol) of the phosphorylated counterparts (see Fig. 5 and Table I). The presence of a proline residue can significantly affect polypeptide structure with the result that sequences may be kinked or bent (48). Although TPS peptides 2 and 12 exhibit a preferred consensus motif designating them as rather good substrates, their phosphorylated analogues did not form. This might be due to the proline residue at 1, which, by producing kinks in these peptides, may impair kinase binding. Studies of target peptide variants are needed to identify the effect of this proline residue on substrate targeting.
TPS peptides 5, 6, 13, and 16 contain the less preferred consensus motif for SnRK1s (47). In contrast to the preferred consensus sequence where the requisite basic residue is located in position 3 with respect to the target serine/threonine, here the basic residue is in 4 (see Fig. 4). These peptides might therefore be suspected to make poor SnRK1 substrates. In vitro incubations revealed that kinases did not phosphorylate TPS peptides 5, 13, and 16 in either the presence or absence of Ca2+. TPS peptide 6, on the other hand, was phosphorylated. However, the rather low phosphorylation level of this substrate peptide remains consistent with the presence of the less preferred consensus motif for SnRK1s.
The observation that phosphopeptide formation for six of the TPS substrate peptides is Ca2+-dependent (Fig. 5 and Table I) implies that additional, Ca2+-dependent kinases in the crude extract also recognize these peptides. In fact, the consensus sequence for SnRK1s includes the minimal recognition motif for calcium-dependent protein kinases (CDPKs), generally given as -X-Basic-X-X-Ser-X-X-X-
where
is a hydrophobic residue and X is any residue (15). That CDPKs and SnRK1s have similar recognition motifs is consistent with the fact that both protein kinases are members of the same family (12). The complete inhibition of the in vitro phosphorylation of TPS peptides 7, 11, and 15 by substitution of CaCl2 with EGTA, however, excludes a role of SnRK1s in the phosphorylation of these peptides and strongly suggests the phosphorylation seen is exclusively the work of CDPKs. Interestingly all TPS peptides with in vitro phosphorylation not dependent on Ca2+ exhibit a target threonine instead of a serine.
Not only were differences in the degree of and the Ca2+ dependence of phosphorylation observed among various members of the TPS gene family, variation was also seen among TPS peptides originating from the same TPS isozymes. For example, TPS peptides 9, 10, 11, and 12, all representing TPS isozyme At1g60140, exhibited diverse behavior in the in vitro kinase assay. TPS peptide 10 was phosphorylated in a Ca2+-independent manner, whereas the phosphorylation of TPS peptides 9 and 11 was Ca2+-dependent. TPS peptide 12 did not act as a kinase substrate at all. TPS isozyme At1g68020 peptides 13, 14, and 15 showed little or no phosphorylation, offering this isozyme a potential function as a negative in vitro kinase activity control for future studies.
Besides their SnRK1/CDPK phosphorylation sites many TPS substrate peptides have additional serine and/or threonine residues in their sequence that are potential phosphorylation targets of other kinase families. Because the MRM approach used in this study is not suitable for the identification of the specific phosphorylation sites of the TPS phosphopeptides, MS3 spectra from standard TPS phosphopeptides and phosphopeptides formed in the in vitro kinase activity assay were recorded using a linear ion trap mass spectrometer. MS2 spectra generated from phosphopeptides often do not provide enough peptide backbone fragmentation necessary for unambiguous identification of phosphorylation sites because of a dominant neutral loss of phosphoric acid during MS2 fragmentation (40). Here we made use of the neutral loss peak derived from serine/threonine phosphorylated TPS peptides to trigger a data-dependent MS3 fragmentation step. However, a chromatographic separation of the in vitro phosphorylated TPS peptides in front of mass spectrometric analysis as described under "Experimental Procedures" is indispensable for good quality MS3 spectra. This ensures that other compounds and contaminations present in the reaction mixture are clearly separated thus limiting unwanted ionization suppression by the matrix. The resulting neutral loss-driven MS3 spectra of phosphorylated TPS peptides are informative enough to identify the correct phosphorylation site. It could be shown that the MS3 spectra of the synthetic standard TPS phosphopeptides were identical to the MS3 spectra generated from in vitro formed TPS phosphopeptides. Consequently an in vitro phosphorylation of alternative residues within the TPS peptide sequences could be ruled out. Fig. 6 exemplarily shows a comparison of full MS3 spectra generated from standard phospho-TPS peptides 1, 3, 4, 6, and 15 (all containing additional serine/threonine residues in their sequences) and their counterparts formed in vitro.
With regard to the selectivity of detection the triple stage quadrupole instrument operated in MRM mode yielded significantly higher S/N ratios (up to 50-fold) for the TPS phosphopeptides than the linear ion trap mass spectrometer in the neutral loss-driven MS3 scanning mode when using the same chromatographical conditions. Theoretically the neutral loss-driven MS3 approach is more selective than MS2-based MRM because of one additional fragmentation step in the detection method. However, the enhanced resolution achieved by the quadrupole mass filters of the triple stage quadrupole instrument compensates for that. The relatively low mass accuracy of ion trap instruments does not allow for highly specific precursor selection. As a result, the S/N of detection is greatly increased using the MRM-based approach. This is reflected by the LODs for the TPS phosphopeptides that ranged from 20 fmol to 1.1 pmol for MRM detection (Table I), whereas the LODs using the neutral loss-driven MS3 approach were estimated to be in the picomolar range.
In the near future, the comparison of wild-type and mutant plants harvested at different points of the day/night cycle will allow a deeper insight into the phosphoregulation of TPS. With the simultaneous analysis outlined here, it is possible to cross-correlate TPS isozyme phosphorylation data with quantitative expression data. The utilization of peptide libraries will enable the fast and simultaneous analysis of hundreds of peptides. Especially important is the use of peptides containing putative phosphorylation sites predicted by bioinformatic analysis of the proteome and prediction of consensus motifs (non-random peptide libraries) as demonstrated in this study. In this regard, the method has potential implications for the fast and robust testing of new peptide substrates to distinguish activities of CDPKs from those of SnRK1s (even in crude extracts) and for the screening of new phosphorylation sites in plant proteins.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Published, MCP Papers in Press, July 19, 2005, DOI 10.1074/mcp.M500134-MCP200
1 The abbreviations used are: TPS, trehalose-6-phosphate synthase; CDPK, calcium-dependent protein kinase; FA, formic acid; FWHM, full width at half-maximum; LODs, limits of detection; MRM, multiple reaction monitoring; S/N, signal-to-noise; SnRK1, sucrose non-fermenting 1-related protein kinase 1; SPS, sucrose-phosphate synthase.
* This work was supported by the Max Planck Society.
To whom correspondence should be addressed. Tel.: 49-331-5678109; Fax: 49-331-5678134; E-mail: weckwerth{at}mpimp-golm-mpg.de
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REFERENCES |
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