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Cost-effective isobaric tagging for quantitative phosphoproteomics using DiART reagents† Nikhil Ramsubramaniam,a Feng Tao,b Shuwei Licd and Mark R. Marten*a We describe the use of an isobaric tagging reagent, Deuterium isobaric Amine Reactive Tag (DiART), for quantitative phosphoproteomic experiments. Using DiART tagged custom mixtures of two phosphorylated peptides from alpha casein and their non-phosphorylated counterparts, we demonstrate the compatibility of DiART with TiO2 affinity purification of phosphorylated peptides. Comparison of theoretical vs. experimental reporter ion ratios reveals accurate quantification of phosphorylated peptides over a dynamic

Received 20th August 2013, Accepted 25th September 2013 DOI: 10.1039/c3mb70358d

range of more than 15-fold. Using DiART labelling and TiO2 enrichment (DiART-TiO2) with large quantities of proteins (8 mg) from the cell lysate of model fungus Aspergillus nidulans, we quantified 744 unique phosphopeptides. Overlap of median values of TiO2 enriched phosphopeptides with theoretical values indicates accurate trends. Altogether these findings confirm the feasibility of performing quantitative

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phosphoproteomic experiments in a cost-effective manner using isobaric tagging reagents, DiART.

Introduction Protein phosphorylation, as the most ubiquitous post-translational modification,1 plays critical roles in eukaryotic cells, such as impacting protein stability, modulating catalytic activity, and regulating protein–protein and protein–nucleic acid interactions.2,3 It is estimated that roughly one third of all eukaryotic proteins are phosphorylated at some point of time.1,4 Because of its importance, quantitative assessment of the phosphorylation states of cells, or phosphoproteomic analysis, has recently emerged as a powerful analytical tool, making it possible to elucidate complex gene regulatory networks, as a result of concerted actions among protein kinases and phosphatases, at a global level.5 Phosphoproteomic workflows differ from traditional, shotgun proteomic approaches that have been used in the past decade. a

Department of Chemical, Biochemical & Environmental Engineering, UMBC, Engineering Building, Room 314, 1000 Hilltop Circle, Baltimore, MD 21250, USA. E-mail: [email protected]; Tel: +1-410-455-3439 b Omic Biosystems, 9700 Great Seneca Highway, Room 115, Rockville, MD 20850, USA. Tel: +1-301-850-4949 c Department of Chemistry and Biochemistry, University of Maryland, College Park, MD 20742, USA. Tel: +1-240-314-6330 d Institute for Bioscience and Biotechnology Research, University of Maryland, Rockville, MD 20850, USA. Tel: +1-240-314-6330 † Electronic supplementary information (ESI) available: LC MS/MS data used to calculate false localization rates are organized in Table S1. Peptide identification from Mixture-1 and Mixture-2, obtained post-enrichment and pre-enrichment, is provided in Table S2a and S2b respectively. Quantitative LC MS/MS data obtained post-enrichment, pre-enrichment and spike-in, using the DiART-TiO2 approach to Mixture-1 and Mixture-2, are provided in Table S3a, b and c respectively. Quantitative LC MS/MS data obtained using the DiART-TiO2 approach to Mixtures 3–6 are provided in Table S4a–d respectively. See DOI: 10.1039/c3mb70358d

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The low abundance of phosphoproteins, along with the low stoichiometry of phosphorylation events requires additional enrichment and fractionation steps.6,7 For example, performing affinity purification using different resins dramatically improves the selectivity of phosphopeptide identification.8,9 Moreover, to identify a larger subset of the phosphoproteome, it is often necessary to fractionate samples before or after affinity enrichment of phosphopeptides.10 A typical phosphoproteomic workflow consists of two steps. The first step involves prefractionation, which is aimed at reducing the complexity of a tryptic peptide mixture. This is typically accomplished using a chromatographic technique such as SCX, ERLIC or HILIC.9 Finally, enriched phosphopeptides are desalted using a C18 column to ensure compatibility with subsequent MS analysis. Each of these steps can involve significant peptide loss, which results in the need for a significant amount of protein starting-material. For example, ‘‘large-scale’’ phosphoproteomic workflows use between 2 and 10 mg of protein startingmaterial.10 While these quantities of protein are often available, the cost associated with isobaric tagging of this material (i.e., based on dollars spent per experiment) can be prohibitive.11 One approach to reduce cost is to add isotopic tags to peptides, after performing phosphopeptide enrichment.12 However, experimental reproducibility associated with manually implemented affinity enrichment is difficult to control among various samples, in part compromising the benefit gained through the use of isotopic labeling. There are two types of isotopic labeling approaches, massshift and isobaric tags. Isobaric tags such as TMT and iTRAQ have certain advantages over mass-shift tags, including concurrent analysis of more samples (up to eight with currently available

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isobaric reagents like iTRAQ) and enhanced sensitivity. Yet, TMT and iTRAQ can be prohibitively expensive to be used for large-scale quantitative phosphoproteomic assays.11,13 A simple solution to this problem is to adopt a low-cost replacement for iTRAQ or TMT such as Deuterium isobaric Amine Reactive Tags (DiART).14,15 DiART offers up to 6-plex quantitation, which doubles the number of samples that can be analyzed concurrently by dimethyl labeling, a mass-shift tag.11,14 Using higher collision energy dissociation (HCD) as a method of fragmentation, researchers have shown that DiART tagged peptides are easier to fragment as compared to iTRAQ, leading to significantly higher reporter ion intensities available for quantification.15 The synthesis of DiART reagents involves fewer steps and has higher yields, thereby reducing synthesis cost by almost 10-fold.14 Even when additional costs influencing sales (e.g., marketing costs, overhead, profit, volume discounts, etc.) are accounted for, the sale price of DiART is approximately 75% lower than other commercially available isobaric tagging reagents (e.g., iTRAQ). Thus, the high cost problem that has so far prevented the use of isobaric reagents for large-scale quantitative phosphoproteomic workflows can be significantly alleviated. The broad goal of our work is to examine the applicability of DiART reagents in quantitative phosphoproteomic workflows involving TiO2 enrichment. Initially, we performed proof-ofconcept studies using custom mixtures of synthetic peptides and low amount of synthetic phosphopeptides. These mixtures were tagged with DiART reagents, enriched for phosphopeptides using TiO2 and analyzed by mass spectrometry. The same strategy was used to examine the capability of DiART reagents to quantify phosphopeptides obtained from complex cell lysates of model fungus, Aspergillus nidulans.

Results and discussion Initial experiments employed two alpha-casein synthetic peptides to optimize phosphorylation–localization criteria for phosphopeptides identified using mass spectrometry. These peptides were Table 1

dual (Peptide-App) and mono (Peptide-Bp), phosphorylated at known positions (see Experimental section). To ensure confident identification of the phosphorylation sites, both peptides (Peptide-App and Peptide-Bp) were injected into the mass spectrometer and LC-MS/MS data were acquired using collision induced dissociation (CID). Pseudo MS3, or multistage activation, was used in order to fragment ions arising from phosphorylation losses. Several software packages (e.g., Ascore,16 MD Score,17 PhosphoRS18) are available to localize phosphosites. We used PhosphoRS (available as a module in Proteome Discoverer 1.3) to confidently localize phosphosites. A localization probability of 95% on PhosphoRS was used to count the total number of localizations, which were then categorized as either correct or incorrect. The false localization rate was calculated as the ratio of the total number of incorrect localizations to that of localizations, multiplied by 100. Using this approach, we obtained a false localization rate (FLR) of 1.05% (see Table S1, ESI†), a threshold routinely used for phosphoproteome analysis.19,20 This validates our ability to confidently localize phosphorylated residues in phosphopeptides. To assess the applicability of DiART reagents for quantitative phosphoproteomic analysis, we made two custom mixtures (Mixture-1 and Mixture-2, Table 1). These were composed of the phosphorylated peptides described above (Peptide-App and Peptide-Bp) and non-phosphorylated versions of the same peptides (Peptide-A and Peptide-B) labeled by DiART tags (114, 115, 118, 119). Using Mixture-1 and Mixture-2, six different reporter ion ratios were obtained for each phosphorylated peptide. To ensure phosphopeptides were low abundant species in the mixtures, the ratio of each tagged non-phosphorylated peptide to that of the phosphorylated counterpart (A/App or B/Bp) was kept in the range of 3 to 10 in each channel (see Table 1). For instance, when Mixture-1 was analyzed by LC-MS/MS, approximately 10% of the peptide spectral matches were phosphopeptides (Fig. 1). This corroborates the under-representation of these phosphopeptides in our custom mixtures.

Quantities of different components of peptide mixtures used in the study

Peptide load Mixtures

114 channel (mg)

Expected reporter ion ratios 115 channel (mg)

118 channel (mg)

119 channel (mg)

115/114

118/114

119/114

Mixture-1 App Bp A B Total

5 1 29 15 50

2.5 2 25.5 20 50

5 5 20 20 50

1.5 3 15.5 30 50

0.50 2.00 0.88 1.33

1.00 5.00 0.69 1.33

0.30 3.00 0.53 2.00

Mixture-2 App Bp A B Total

1.5 3 15 30.5 50

3 1.5 30 15.5 50

7.5 3 22.5 17 50

4.5 1 22.5 22 50

2.00 0.50 2.00 0.51

5.00 1.00 1.50 0.56

3.00 0.33 1.50 0.72

— — — —

— — — —

0.3 1.0 2.0 4.0

— — — —

— — — —

Mixtures 3–6 Lysate (Mixture-3) Lysate (Mixture-4) Lysate (Mixture-5) Lysate (Mixture-6)

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1500 1000 666 400

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Fig. 1 Percentage of peptide spectral matches (PSMs) containing at least one phosphosite identified before and after TiO2 enrichment. LC-MS/MS data were acquired before and after performing TiO2 enrichment. PSMs were identified with Proteome Discoverer 1.3 and S/T/Y sites annotated with 95% or higher phosphorylation localization probability by phosphoRS were counted as phosphosites for analysis. Percentage of spectra containing at least one phosphosite was averaged between Mixture-1 and Mixture-2.

Next, we performed affinity enrichment for DiART tagged phosphopeptides using TiO2 resin (hereafter called DiARTTiO2). In DiART reagents, deuterium is strategically placed next to hydrophilic groups to eliminate deuterium-related chromatographic effects in reverse phase HPLC.14 To ensure that there were no interfering interactions between deuterium and TiO2 resin, LC-MS/MS data were acquired for DiART tagged peptide mixtures prior to, and after, TiO2 enrichment. Peptides were identified using CID spectra, while quantification was performed using pulsed Q dissociation (PQD) spectra (see Fig. S1, ESI†). We observed a remarkable enrichment of phosphopeptides, after which 74% of the peptide spectral matches showed a confident phosphosite localization using collision induced dissociation (Fig. 1; Table S2a and b, ESI†). Further, we calculated reporter

Method ion ratios from LC-MS/MS data after performing TiO2 enrichment (Fig. 2; Table S3a, ESI†). Reporter ion ratios for different channels were taken with respect to the 114 tag. Data in Fig. 2 show that both Peptide-App and Peptide-Bp exhibited linear trends (R2 > 0.95) on a plot of expected versus theoretical reporter ion ratios, over 15-fold change. Fold change was calculated as the ratio of highest theoretical fold change to the lowest theoretical fold change, used in these experiments. This demonstrates the capability of a DiART-TiO2 approach to quantify differences in phosphopeptide mixtures. In addition, the slopes obtained for phosphorylated peptides after performing TiO2 enrichment did not differ significantly, when compared to that obtained prior to TiO2 enrichment (Peptide-App, P-value = 0.906; Peptide-Bp, P-value = 0.139; see Table S3b, ESI†), which implies TiO2 resins do not detrimentally hinder quantitation of DiART labelled phosphopeptides. To examine the feasibility of using our DiART-TiO2 approach to quantify differential expression of phosphopeptides in a complex mixture, we carried out ‘‘spike-in experiments in which the peptide mixtures described above were diluted with a complex protein mixture—A. nidulans, cell lysate. In complex biological samples, phosphopeptides are typically present at a relatively low abundance, which can cause significant variability in quantitation using isobaric mass tags. Furthermore, contributions from the background (e.g., co-elution of peptides or peptides of equivalent m/z values) can alter the feasible dynamic range, thus affecting the accuracy of quantitation.21,22 To determine if these challenges could be overcome, we performed spike-in experiments with 40 mg Mixture-1 and Mixture-2, each containing 5 mg of phosphopeptides. Each of these was diluted 10-fold with the A. nidulans cell lysate (i.e., 400 mg added) tagged with DiART reagents 114, 115, 118, 119 and mixed in the ratio 1 : 1 : 1 : 1 (see Fig. S2, ESI†). This was followed by TiO2 enrichment. Thus, the post-enrichment mixture should theoretically contain

Fig. 2 A plot of experimental versus theoretical reporter ion ratios for synthetic phosphopeptides: (A) DIGSpESpTEDQAMEDIK (App) and (B) TVDMESpTEVFTK (Bp). They were labelled by DiART tags (114, 115, 118, 119) and mixed with their non-phosphorylated counterparts in amounts shown in Table 1. These mixtures were further subjected to TiO2 enrichment and LC-MS/MS data were recorded before (B) and after (U) TiO2 enrichment. Tagged peptide mixtures were also diluted ten-fold with mass tagged (114, 115,118 and 119) A. nidulans cell lysate and subjected to TiO2 enrichment – spike-in experiments (n).

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Method phosphopeptides from Mixture-1 or Mixture-2, as well as background phosphopeptides in A. nidulans having reporter ion ratios of unity. Post-enrichment LC-MS/MS data are shown in Fig. 2 (see Table S3c, ESI†). Both Peptide-App and Peptide-Bp exhibited linear trends on a plot of expected versus theoretical log reporter ion ratios. Furthermore no statistically significant difference was observed when we compared slopes from the spike-in experiment, to that obtained from peptide mixtures after TiO2 enrichment (Peptide-App, P-value = 0.417; Peptide-Bp, P-value = 0.973). This suggests that our DiART-TiO2 approach is capable of providing quantitative assessment of phosphopeptide abundance in the presence of a complex background. To confirm our findings above, we carried out a large-scale labeling study using our DiART-TiO2 approach to quantify phosphopeptides in the A. nidulans cell lysate. A total of 8 mg cell lysate was trypsinized and then divided into four 2 mg aliquots (Table 1, Mixture-3, -4, -5, and -6). Each sample was further divided into two aliquots with different amount of proteins and labeled with DiART reagents 114 and 115 respectively. Labeling was performed using chemicals and methodology similar to that described previously,14,15 where labeling efficiency >98% was achieved.15 Enrichment was then performed using TiO2 resin. These enriched fractions were then further separated using ultralong gradients in an online LC-MS/MS system. This approach has been used recently to identify thousands of proteins with sensitivity on par with 2D LC-MS/MS approaches.23,24 In our experience, as little as 20–30% ACN was adequate to elute most phosphopeptides from the C18 column (see the ESI,† Fig. S3). Using the 95% phosphosite localization probability described above, we identified and quantified 744 unique phosphopeptides. Quantitative data from these enriched mixtures were plotted against theoretical values (Fig. 3). Data show strong correspondence between observed ratios (logarithm of 115/114) of quantified phosphopeptides in

Fig. 3 Experimental versus expected reporter ion ratios for large-scale labeling experiments. Data obtained from A. nidulans cell-lysate tagged with DiART reagents 114 and 115 and mixed in specific ratios and subjected to TiO2 enrichment. Expected (U) and experimental median value (*). The ends of open boxes extend from the first (Q1) to third quartile (Q3). The whiskers extend up to one standard deviation or up to the maximum/minimum if they lie within this range.

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Molecular BioSystems each mixture and those expected theoretically. A slightly higher deviation of the reporter ion ratios is seen with higher theoretical ratios, likely due to fold change under/overestimation, often seen in isobaric tagging experiments.25,26 These data demonstrate the applicability of our DiART-TiO2 approach for large-scale quantitative phosphoproteomic experiments. Not only are DiART reagents able to perform quantitative phosphoproteomic analysis with complex cell lysate, but can do so in a cost-effective manner. At the time of writing, the DiART sale price was approximately 25% of another commercially available isobaric mass tag (i.e., iTRAQ), making the cost of such large-scale labeling experiments much more reasonable.

Conclusion A DiART-TiO2 approach was successfully implemented to provide quantitation of phosphopeptides enriched from a complex biological background. Our data indicate that low cost DiART tagging is compatible with TiO2 enrichment for phosphoproteomic analysis. DiART demonstrates successful quantitation of both singly and doubly phosphorylated phosphopeptides over a 15-fold dynamic range. Using this DiART-TiO2 approach, more than 700 unique phosphopeptides were quantified from a cell lysate of the model filamentous fungus A. nidulans.

Experimental section A. nidulans cell growth Aspergillus nidulans parent strain A1166 (Fungal Genetics Stock Center, Kansas City, Missouri, USA) was grown at 30 1C for 3–7 days on potato dextrose agar (Difco, Detroit, USA). Spores from agar plates were stored in 20% w/v glycerol (Sigma-Aldrich, St Louis, USA) at 80 1C. Growth was carried out as described previously.27 Briefly, spore suspension containing 8  106 spores was thawed for 1 h. The suspension was vortexed for 30 seconds and inoculated into 50 ml of complex media (MAGUU) containing 2% (w/v) malt extract (Becton Dickinson, MD, USA), 0.2% peptone (Becton Dickinson, MD, USA), 0.122% uridine (Acros, NJ, USA), 0.112% uracil (Acros, NJ, USA) and 7.5 g L 1 glucose at pH 3.3. MAGUU also contained the following components per liter, 6 g NaNO3, 0.52 g KCl, 0.82 g KH2PO4, 1.05 g K2HPO4 and 0.52 g MgSO47H2O (Fisher, NJ, USA) and 1000 ppm of filter-sterilized trace-element solution (per liter, 5 g FeSO47H2O, 22 g ZnSO47H2O, 11 g H3BO3, 5 g MnCl24H2O, 1.6 g CoCl26H2O, 1.6 g CuSO45H2O, 1.1 g (NH4)6Mo7O244H2O, and 50 g EDTA). Spores were incubated in 50 ml of low pH MAGUU for 12 h at 30 1C at an agitation speed of 250 rpm. This was then used as inoculum for a 2.8 L baffled flask containing 1.2 L of MAGUU containing 7.5 g L 1 glucose, 0.5% malt extract, 0.2% peptone, 0.122% uridine, 0.112% uracil at pH 6.5. Other components of the MAGUU were present in the same concentration as that used to germinate spores. After cells were cultivated for 20 hours at 30 1C at 250 rpm in an oscillating shaker, they were harvested using non-gauze milk filters (Ken AG, Ohio, USA) and stored at 80 1C for protein extraction.

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Synthetic protein mixtures Mixtures of four 4 synthetic peptides were used in the proof-ofconcept experiments (Table 1). This was comprised of phosphorylated and unphosphorylated versions of two synthetic peptides, Peptide-A and Peptide-B (Thermo Scientific, Rockford, Illinois, USA). Peptide-App has phosphorylation at two serine residues whereas Peptide-Bp has phosphorylation at one serine residue. Peptide-A, Peptide-App, Peptide-B and Peptide-Bp have sequence DIGSESTEDQAMEDIK (M.W 1767.9 Da), DIGSpESpTEDQAMEDIK (M.W 1926.0 Da), TVDMESTEVFTK (M.W 1386.6 Da) and TVDMESpTEVFTK (M.W 1466.6 Da). Protein extraction and digestion To extract the cell-lysate protein mixture, frozen mycelia were finely crushed using liquid nitrogen in a mortar and pestle.27 Thereafter they were suspended in lysis buffer comprised of 8 M urea, 4% 3-[(3-cholamidopropyl) dimethylammonio]-1-propanesulfonate (CHAPS), 10 mM dithiothreitol (DTT), and 40 mM Tris–HCl, pH = 8.0. They were then sonicated in ice 3 times, at pulses of 10 seconds each, for a total of 30 seconds. The proteins were reduced using 10 mM DTT at 60 1C for 1 h and further alkylated with 55 mM iodoacetamide, in the dark, at room temperature, for 45 minutes. The proteins were then precipitated with 6 volumes of ice cold acetone at 20 1C for 6 hours and redissolved in 0.5 M tetraethylammonium bicarbonate (TEAB), 0.1% sodium dodecyl sulfate (SDS), pH 8.5. The solution was well vortexed, centrifuged and the supernatant was analyzed using a BCA assay to determine protein concentration. For trypsin digestion the protein was dissolved in trypsinization buffer (0.5 M TEAB, 0.1% SDS) to a final concentration of 5 mg ml 1. The proteins were digested by proteomic grade trypsin (Sigma-Aldrich, St Louis, USA) with a protein to enzyme ratio of 50 : 1 for 18 hours at 37 1C. DiART labeling A tube containing lyophilized DiART reagents was thawed briefly for 2–4 minutes. Anhydrous isopropanol was added to make a 50 mM DiART solution. This was added to a microcentrifuge tube containing trypsinized protein solution in the lysis buffer (pH 8.5). The volume of DiART solution added was calculated assuming a ratio of DiART solution to trypsinized protein solution of 9 : 4 (2.25 : 1). The labeling reaction was performed for 2 hours at 30 1C. The reaction was stopped by acidifying the mixture to pH = 3 using trifluoro acetic acid. Phosphopeptide enrichment To enrich phosphopeptides from the various synthetic peptide mixtures (Table 1), TiO2 spin-tips (Thermo Fisher scientific, Rockford, Illinois) were used. 200 mg of Mixture-1 or Mixture-2 (Table 1) was lyophilized to dryness. This was followed by desalting with C18 EWP SPE cartridges (Agilent, CA, USA), the flow through of which was desalted using C8 hypersep SPE cartridges (Thermo Fisher Scientific, Rockford, Illinois). The eluents from the SPE cartridges were pooled and lyophilized to dryness. TiO2 enrichment was performed three times using

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Method 40 mg of desalted Mixture-1, Mixture-2, leading to three replicates. The following buffers were prepared: Buffer A: 80% acetonitrile, 0.4% TFA; Buffer B: 400 ml lactic acid in 1.4 ml Buffer A; Elution Buffer 1: 740 mM ammonium hydroxide; Elution Buffer 2: 5% pyrrolidine in water. TiO2 spin tips were activated by centrifuging with 20 ml Buffer A for 2 minutes at 3000g. This step was repeated with 20 ml Buffer B. Lyophilized peptide mixtures were dissolved in 150 ml Buffer B. Phosphopeptides were bound to TiO2 in spin-tips by centrifuging this mixture twice at 1000g for 10 minutes. The spin tips were washed once with 20 ml Buffer B followed by washing 3 times with Buffer A. Each of the washing steps were performed at 3000g for 2 minutes. This was followed by elution steps using 50 ml Elution Buffer 1 followed by 50 ml Elution buffer 2. Elution step was performed by centrifuging at 1000g for 5 minutes. The eluents were pooled together, lyophilized and cleaned with C18 SPE cartridge followed by C8 SPE cartridge in series as described earlier. This was lyophilized and dissolved in LC-MS/ MS buffer (0.1% formic acid). Three repetitive enrichments were performed and used for HPLC MS/MS analysis. To enrich phosphopeptides from the complex cell lysate, the lyophilized DiART tagged trypsinized cell lysate was desalted using C18 SPE cartridges (Agilent). The flow through was cleaned up using C8 hypersep SPE cartridges (Thermo Fisher Scientific, Rockford, Illinois). The eluents from the SPE cartridges were pooled and lyophilized to dryness. TiO2 beads (Glygen, Columbia) were preincubated first in 200 mL loading buffer (80% ACN, 2% TFA, 30 mg ml 1 2,5-dihydroxybenzoic acid). The desalted peptide samples were dissolved in 200 mL loading buffer, and then incubated with 12 mg TiO2 beads (1 : 6 peptide : beads ratio). Phosphopeptide binding was performed again, wherein the supernatant was incubated with another aliquot of freshly prepared TiO2 beads for sequential enrichment. The incubated beads were then washed once with 800 mL wash buffer 1 (65% ACN/0.5% TFA) followed with one wash using wash buffer 2 (65% ACN/0.1% TFA). The bound peptides were eluted once with the 200 mL elution buffer I (300 mM NH4OH/50% ACN) and twice with 200 mL elution buffer II (500 mM NH4OH/60% ACN). All incubation, washing and elution procedures were rotated end-over-end at 1000 rpm for 20 min at room temperature. All eluates were acidified by adding 8 mL of TFA acid each. This was further subjected to cleanup using C18 SPE column, and C8 Hypersep SPE column (Thermo Fisher) in series. The eluates were dried down and reconstituted in 0.1% formic acid/H2O. Three repetitive injections of the sample were used for HPLC MS analysis. Reversed phase nano LC ESI MS/MS The LC-MS/MS setup comprised of Dionex Ultimate 3000 nano high-performance liquid chromatography coupled to an LTQ Xl mass spectrometer (Thermo Electron, San Jose, CA). Peptides were loaded onto a trap column cartridge, Acclaim Pepmap C18 (300 Å, 5 mm 300 mm I.D  5 mm, Dionex, Sunnyvale, CA) at 30 mL min 1. Bound peptides were washed for 3 minutes with 0.1% formic acid (FA) and 5% acetonitrile, following which peptides were injected into an analytical column, Acclaim

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Method PepMAP300 C18 (75 micron I.D  15 cm, 5 mm 300 Å), and separated using reversed phase chromatography and a gradient buffer system composed of buffer A (0.1% FA) and buffer B (0.1% FA, 99.9% acetonitrile). For the peptide mixtures HPLC runtime was fixed at 4 hours, with the following gradient, 3–15 min 4% B to 4% B, 15–90 min 4% B to 55% B, 90–100 min 55% B to 100% B, 100–110 min 100% B to 100% B, 110–120 min 4% B to 4% B. For phosphopeptides enriched from the A. nidulans cell lysate the HPLC run-time was fixed at 12 hours with an ultralong gradient, 3–220 min, 4% B to 20% B; 550–650 min, 20% B to 30% B, 650–680 min 30% B to 70% B, 680–690 min 70% B to 100% B, 690–700 min 100% B to 100% B, 700 –710 min 100% B to 4% B, 710–720 min 4% B to 4% B. All ultra long-gradient runs were performed in triplicate. The LTQ XL mass spectrometer (Thermo Electron, San Jose, CA) was operated in positive ion mode with data-dependent MS/MS acquisition. The full mass scan range for the instrument was fixed at 400–2000 m/z. This was followed by data dependent scan of the 2 most abundant ions in alternating CID/PQD mode of fragmentation. Dynamic exclusion was enabled for MS/MS analysis with a repeat count of 2 within a repeat duration of 30 seconds. Exclusion duration was set to 10 s with an m/z window of 0.5 to + 1.5 Da. Parameters for CID were 35% CE, an activation Q of 0.25, and an activation time of 30 ms whereas parameters set for PQD were collision energy of 32%, an activation Q of 0.65 and an activation time of 0.1 ms. Data analysis Raw files were analyzed using Proteome Discoverer 1.3. A fasta file containing 12 470 entries corresponding to A. nidulans was downloaded from Uniprot (http://www.uniprot.org) on 02.08.2010 and incorporated into Proteome Discoverer. This database was used to search for A. nidulans proteins. Accession numbers corresponding to alpha casein proteins were acquired from Uniprot and a fasta file was created. Unimod files containing necessary information pertaining to DiART reagents were acquired from Omic Biosystems, Rockville, MD, USA and were imported into Proteome Discoverer 1.3. Only CID spectra were used for identifications. Prior to performing Sequest search, all CID spectra were preprocessed by TopNPeak nodes to include the top ten most abundant peaks in 100 Da intervals, thereby reducing noise in the MS/MS spectra. The following search parameters were used for Sequest search: DiART modification of the N-terminus as a static modification, DiART modification of lysines, oxidation of methionine, carbidomethyl modification (57 Da) on cysteines and phospho modification of 79 Da on S,T,Y as dynamic modifications. Two missed cleavages by trypsin, parent ion tolerance of 1.4 Da and fragment ion tolerance of 1 Da, were permitted. For identification score (Xcorr: 1.5, 2, 2.25) vs. charge state (+1, + 2, + 3) and peptide rank of 1 were used as filters. These settings were validated using Scaffold 4.0. Peptide FDRs of 1.28% were obtained using these settings on Aspergillus nidulans cell lysates. Furthermore, the phosphoRS node was used to select for phosphopeptides. Peptides having at least one phosphosite (having 95% or greater phosphorylation localization probability) were considered for analysis. Only PQD

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Molecular BioSystems spectra were used for quantitation. Extreme reporter ion ratios above 30 were neglected. Outlier removal was done at the peptide level wherein reporter ion intensities below the 10th percentile and above the 90th percentile for each channel were considered as outliers. Such an approach to remove outliers has been performed before.28 Final reporter ion ratios for each peptide were calculated using summed intensity averaging29 i.e. by taking the ratio of sum of intensities in the numerator channel (115/118/119) to that in the denominator channel (114), after discarding intensities which contributed to outliers in each channel.

Acknowledgements We thank Joshua Wilhide for maintaining LC-MS/MS equipment and the associated computational setup, used for data analysis. We also thank Bill Moss for developing reproducible methods to fractionate peptides tagged with isobaric reagents. Funding was provided by the Maryland Technology Enterprise Institute (MIPS Project #4913). This work was partially supported by the National Science Foundation under grant no. 1159973 to MRM. Any opinions, findings, and conclusions or recommendations expressed in this manuscript are those of the authors and do not reflect views of National Science Foundation.

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Mol. BioSyst., 2013, 9, 2981--2987

2987

Cost-effective isobaric tagging for quantitative phosphoproteomics using DiART reagents.

We describe the use of an isobaric tagging reagent, Deuterium isobaric Amine Reactive Tag (DiART), for quantitative phosphoproteomic experiments. Usin...
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