Stable Isotope N-Phosphorylation Labeling for Peptide de Novo

Article pubs.acs.org/ac

Stable Isotope N‑Phosphorylation Labeling for Peptide de Novo Sequencing and Protein Quantification Based on Organic Phosphorus Chemistry Xiang Gao,†,‡ Hanzhi Wu,† Kim-Chung Lee,† Hongxia Liu,‡ Yufen Zhao,§ Zongwei Cai,*,†,‡ and Yuyang Jiang*,‡ †

Department of Chemistry, Hong Kong Baptist University, Kowloon Tong, Hong Kong, SAR, China The Key Laboratory for Cancer Metabolomics of Shenzhen, Graduate School at Shenzhen, Tsinghua University, Shenzhen, 518055, China § Department of Chemistry and The Key Laboratory for Chemical Biology of Fujian Province, College of Chemistry and Chemical Engineering, Xiamen University, Xiamen, 361005, People’s Republic of China ‡

S Supporting Information *

ABSTRACT: In this paper, we describe the development of a novel stable isotope N-phosphorylation labeling (SIPL) strategy for peptide de novo sequencing and protein quantification based on organic phosphorus chemistry. The labeling reaction could be performed easily and completed within 40 min in a one-pot reaction without additional cleanup procedures. It was found that N-phosphorylation labeling reagents were activated in situ to form labeling intermediates with high reactivity targeting on N-terminus and ε-amino groups of lysine under mild reaction conditions. The introduction of N-terminal-labeled phosphoryl group not only improved the ionization efficiency of peptides and increased the protein sequence coverage for peptide mass fingerprints but also greatly enhanced the intensities of b ions, suppressed the internal fragments, and reduced the complexity of the tandem mass spectrometry (MS/MS) fragmentation patterns of peptides. By using nano liquid chromatography chip/time-of-flight mass spectrometry (nano LC-chip/TOF MS) for the protein quantification, the obtained results showed excellent correlation of the measured ratios to theoretical ratios with relative errors ranging from 0.5% to 6.7% and relative standard deviation of less than 10.6%, indicating that the developed method was reproducible and precise. The isotope effect was negligible because of the deuterium atoms were placed adjacent to the neutral phosphoryl group with high electrophilicity and moderately small size. Moreover, the SIPL approach used inexpensive reagents and was amenable to samples from various sources, including cell culture, biological fluids, and tissues. The method development based on organic phosphorus chemistry offered a new approach for quantitative proteomics by using novel stable isotope labeling reagents.

A

methods including mass-difference labeling and isobaric labeling methodologies have been developed for quantitative analysis in proteomics, peptidomics, and glycomics over the past decade.4−7 Relative quantification is based on the introduction of a chemically equivalent but mass-differential stable isotope tags that can be used to determine accurately the abundance of proteins in one sample to another. Isotope incorporation can be performed at the peptide or protein level by using 2H, 13C, 15N, or 18O as heavy isotopes. On the basis of the characteristic isotope patterns, the “light” and “heavy” isotopic peptide pairs from both samples shared the same physicochemical properties and may be compared and distinguished in the mass spectra. Mass spectrometry-based stable isotope labeling methods are mainly divided by the way

nalysis of proteins on a genomewide scale is increasingly important for providing valuable information on biomarker discovery and metabolic pathway analysis, accelerating the development of powerful new diagnostic tools and new drugs and leading to a better understanding of the molecular mechanisms that control cell behaviors. The systematic analysis of all proteins in biological samples such as a tissue or cell was popularized under the name of proteomics.1 Mass spectrometry plays an important role not only in rapidly identifying proteins and determining details of their covalent structures and posttranslational modifications in a biological mixture, but also in high-throughput proteome-wide quantification of proteins, their variants, and protein−protein interactions in cell or tissue in response to a variety of conditions of interest.2 In parallel with label-free quantification, stable isotope labeling coupled with mass spectrometry has been developed and used extensively in the comparison of relative abundances of expressed proteins for quantitative analysis.3 Numerous © 2012 American Chemical Society

Received: July 18, 2012 Accepted: November 7, 2012 Published: November 7, 2012 10236

dx.doi.org/10.1021/ac301939v | Anal. Chem. 2012, 84, 10236−10244

Analytical Chemistry

Article

also should be completed with minimal sample handling under mild reaction conditions. Recently, new reagents and reactions have been reported for chemical labeling for the quantitative proteomics, including the soluble polymer-based isotopic labeling47 and the Caltech isobaric tags (CITs) based on click chemistry.48 In our previous work, a novel chemical labeling concept based on organic phosphorus chemistry was developed for peptide sequencing by using electrospray ionization tandem mass spectrometry (ESIMS/MS). As shown in Figure S-2 (Supporting Information), a serial of stable organophosphorus reagents, such as dimethyl phosphite (DMP),49 diethyl phosphite (DEP),50 diisopropyl phosphite (DIPP),51,52 and ethyloxy (phenyl)phosphinate (EPP),53,54 have been synthesized and applied to peptide de novo sequencing. A phosphoramidate P−N bond was formed, and a neutral phosphoryl group can be easily incorporated into the N-terminus of peptides through a one-pot specific reaction with quantitative yield under mild reaction conditions. It was found that the introduction of a phosphoryl group onto the Nterminus of amino acids and peptides could not only improve the sensitivity in ESI-MS and matrix-assisted laser desorption ionization mass spectrometry (MALDI-MS) analyses55,56 but also simplify the fragmentation pathways and interpretation of peptide sequences from the mass spectra of the derivatized peptides, providing a convenient and rapid method for peptide sequencing. In this paper, we present a novel stable isotope Nphosphorylation labeling (SIPL) method that may be applied for both peptide sequencing and quantitative proteomics based on organic phosphorus chemistry (Figure 1). By proceeding

of stable isotope labels introduced into the peptide or protein. Generally, stable isotope tags can be introduced metabolically, enzymatically, or chemically. Labeled synthetic peptides may be spiked into the sample as internal standard at defined concentrations. Metabolic in vivo labeling of proteins is achieved by supplying isotope-enriched compounds, such as 13 C-, 15N-, or 18O-enriched nutrients8,9 or stable isotope labeled amino acids (SILAC),10−15 to an organism or cell culture in a way that can be metabolized and exclusively incorporated into proteins. Another global quantitative method is based on labeling the terminal carboxylic groups of the tryptic peptides with 18O atoms during or after proteolysis in 18O-enriched water.16−19 Applications of various types of stable isotope tagging by in vitro chemical derivatization of functional groups in proteins or peptides have been reported, mainly on the N-terminal amino group, but also on a less extent the C-terminal carboxylic group or specific amino acid residues with reactive side chains, such as cysteine, lysine, and tryptophan, in order to achieve the quantification from mass spectrometry (MS) or tandem mass spectrometry (MS/MS) analysis.20,21 The chemical labeling strategies are particularly suitable for various sources of biological protein samples, such as tissue samples derived from animals or humans where metabolic incorporation is difficult. Since the introduction of the isotope-coded affinity tag (ICAT) in 1999 by Gygi et al.,22 stable isotope labeling strategies have been widely used in quantitative proteomics with many reagents over the past decade. These methods can be divided into two main categories. The first one is mass difference labeling, such as reductive formaldehyde dimethylation,23−26 acetylation (H3/D3) and propionylation (H5/D5),27 multifunctional imidazole label (H4/D4),28 isotope-coded protein label (ICPL),29−31 ICAT, and dual stable isotope coding (DSIC).32 The second group gaining popularity recently involves isobaric tags that allow the determination of multiple pools of proteins in one single analysis, such as tandem mass tags (TMT),33−35 isobaric tags for relative and absolute quantitation (iTRAQ),36−39 N,N-dimethyl leucine (DiLeu),40 deuterium isobaric amine-reactive tag (DiART),41,42 isobaric peptide termini labeling (IPTL)43 and so on. Most of the labeling methods are based on traditional protein chemistry established during the 1960s through 1970s for the coupling of stable isotopic tags with functional group of proteins or peptides (Figure S-1, Supporting Information). For example, the labeling chemistry of ICAT is derived from the protein reduction and alkylation widely used in protein preparations. During the past decade, the chemical structures of the functional group of the tags have been changed and designed rationally to achieve different properties with novel functions, such as the acid-cleavable ICAT (cICAT)44,45 and visible-coded affinity tags (VICAT).46 However, the reactive groups of ICATbased approaches have not been changed. For most of the quantitative tags targeting on the N-terminal amino group, the carboxylic groups of stable isotope tags are traditionally activated to form anhydrides or succinimidyl esters, which can subsequently be attacked by primary amines through SN2 reaction pathways and lead to the formation of a C−N amide bond. Fundamentally, it should be noted that virtually all successful quantitative labeling approaches are dependent on the step of coupling reactions. However, the coupling chemistry for stable isotope labeling still relies on classic reactions because the involved chemical reactions not only need to be specific but

Figure 1. Scheme of the stable isotope N-phosphorylation labeling (SIPL) procedure for protein quantification based on organic phosphorus chemistry.

through a modified Atherton−Todd reaction, a neutral phosphoryl group was specifically incorporated into tryptic peptides with high yields under mild conditions (Figure S-3, Supporting Information). A deuterium-labeled dimethyl phosphite (D-DMP) was synthesized and employed on a standard peptide, a model protein, and protein mixtures for demonstrating the applicability of the proposed method for relative quantitative proteomics. It was found that this one-pot phosphorylation reaction was simple, highly efficient, and selective, which might provide a rapid, straightforward, and cost-effective mean both for peptide de novo sequencing and for the comparison of the relative abundance of proteins.

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EXPERIMENTAL SECTION Materials and Reagents. Lysozyme, myoglobin, hemoglobin, β-lactoglobulin B, bovine serum albumin (BSA), DL10237



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analytical nano column. The tryptic peptides were separated using a linear gradient of 5−85% solvent B over 60 min at a flow rate of 200 nL/min. The mass spectrometer was performed in positive ion mode. The instrument settings were adjusted during autotune and set to the following: fragmentor voltage 175 V, skimmer 65 V, and the octopole 1 rf voltage 750 V. The temperature and flow rate of drying gas were at 325 °C and 3.5 L/min, respectively. The data acquisition was set to auto MS/MS mode. The precursor ion was isolated based on relative abundances. The doubly charged ions were given the first priority followed by singly charged ions, triply charged, and then other multiply charged ions. The collision energy applied was based on the mass-to-charge (m/z) ratio of the ion. The scan range was generally from m/z 500 to m/z 2000. The data analysis was performed with the Agilent MassHunter qualitative analysis software (Agilent Technologies, Santa Clara, CA).

dithiothreitol (DTT), iodoacetamide (IAA), ammonium bicarbonate, formic acid, trifluoroacetic acid (TFA), triethylamine (TEA), tetrachloromethane, and α-cyano-4-hydroxycinnamic acid (CHCA) were purchased from Sigma (St. Louis, MO, U.S.A.) and used without further purification. Sequencing grade trypsin was obtained from Promega (Madison, WI). The fibrinopeptide A (human, ADSGEGDFLAEGGGVR) was purchased from GL Biochem. Ltd. (Shanghai, China). Dimethyl phosphite (H-DMP) and CD3OD (99.9 atom % D) were obtained from Alfa Aesar Chemical Ltd. (Tianjin, China). Deuterium-labeled dimethyl phosphite (D-DMP) was synthesized using modified Michaelis−Arbuzov reaction according to a reported method (Figure S-3, Supporting Information).57 HPLC grade acetonitrile (ACN) and methanol were purchased from Tedia (Fairfield, OH, U.S.A.). Deionized water (18 MΩ) was produced by a Milli-Q system (Millipore, Bedford, MA, U.S.A.). Unless specified otherwise, all chemicals and solvents were analytical reagents. N-Phosphorylation Labeling of Standard Peptide and Tryptic Peptides. The individual standard peptide (2 μL, 1 mg/mL) or protein digests (2 μL, about 1 mg/mL) were dissolved in a mixture of H2O (10 μL), ethanol (5 μL), and triethylamine (5 μL) in a 0.2 mL PCR tube, gently vortexing and spinning the sample to the bottom of the tube. The reaction mixture was cooled to 0 °C in an ice−water bath for about 10 min. Then, labeling reagents H-DMP or D-DMP (1 μL) in tetracloromethane (5 μL) were added into the above reaction mixture, which was mixed periodically. The reaction was allowed to proceed 40 min at room temperature before evaporating the mixtures to dryness under N2 gas. The residues were dissolved in 50 μL of H2O (0.1% formic acid) and desalted using ZipTip tips (Millipore, Billerica, MA, U.S.A.) according to the manufacture’s instructions. Mass Spectrometry Analysis. MALDI-TOF MS analyses were performed on a Bruker Autoflex II mass spectrometer (Bruker Daltonics, Bremen, Germany) in positive reflection mode. The mass spectrometer was equipped with a pulsed nitrogen laser operated at 337 nm with 3 ns duration pulses and employed stainless steel targets (MTP 384 target ground steel, Bruker Daltonics). ESI-MS analyses were carried out with a quadrupole orthogonal acceleration time-of-flight (TOF) mass spectrometer (API QStar Pulsar, MDS Sciex, Toronto, Canada) with an external nanoelectrospray ion source (Protana, Odense, Denmark). The MS conditions and analysis details are provided in the Supporting Information. Nano LC-Chip/Q-TOF MS Analysis. Quantification of the standard peptide, protein, and protein mixtures was carried out with an Agilent 6200 HPLC-chip/TOF MS system (Agilent Technologies, Santa Clara, CA), which was equipped with an Agilent 1100 nanopump as the analytical pump for sample separation, a capillary pump as the loading pump for sample enrichment and desalting, a microwell-plate autosampler maintained at 6 °C by the thermostat, an HPLC-chip cube as interface, and Agilent 6210 TOF MS as detection. Chromatographic separations were performed on an enrichment column with a volume of 40 nL and an analytical reversed-phase column (Zorbax C18, 5 μm, 43 mm × 75 μm). Mobile phase buffer A was 5.0% ACN/water (v/v) with 0.1% formic acid. Mobile phase buffer B was 95% ACN/water in 0.1% formic acid. A flow rate of 4 μL/min of solvent A was used for sample loading and desalting with 6 μL injection volume for 3 min. Following this, the stream-select module was switched to a positive where the enrichment column became in-line with the

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RESULTS AND DISCUSSION Stable Isotopic N-Phosphorylation Labeling of Standard Peptide and Proteins. To examine the labeling efficiency and selectivity of N-phosphorylation labeling, a synthetic peptide fibrinopeptide A with the sequence ADSGEGDFLAEGGGVR was investigated by nano ESI-MS. As shown in Figure 2A, ions for the native peptide were observed at m/z

Figure 2. Nano ESI-MS spectra of the standard fibrinopeptide ADSGEGDFLAEGGGVR: (A) native peptide; (B) N-terminal HDMP labeling; (C) N-terminal stable isotope D-DMP labeling. 10238



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Figure 3. MALDI-MS spectra of the tryptic peptides of lysozyme: (A) native; (B) H-DMP labeling; (C) stable isotope D-DMP labeling. (*, the number of labeled phosphoryl groups).

1536.3761 and 768.7190 corresponding to the protonated ion [M + H]+ and doubly charged ion [M + 2H]2+, respectively. After labeling, an m/z shift of 108.2746 and 114.4987 from m/z 1536.3761 was found for the H-DMP (light) and D-DMP (heavy) labeled peptide ions, respectively (Figure 2B). This is consistent with the fact that the model peptide has one site available to be labeled at the N-terminal amine of alanine residue. Moreover, Figure 2C shows that [M + H]+ and [M + H + Na]2+ ions of the D-DMP-labeled peptide at m/z 1650.8748 and 836.9596 are 6 and 3 mass units higher relative to the H-DMP-labeled ions at m/z 1644.6570 and 833.8426, respectively, indicating that the mass difference imparted by isotopic labeling (at least 3 or 4 Da mass shift is needed) was enough to minimize quantitative errors resulting from isotopic overlapping. The obtained data also showed that complete labeling was achieved because no peaks at the m/z corresponding to native peptides were observed in mass spectra of H-DMP- and D-DMP-labeled peptides. Figure 3 shows the MALDI-TOF MS spectra of the native tryptic peptides as well as H-DMP- and D-DMP-labeled digest of standard lysozyme. The assigned peptide sequences are summarized in Table S-1 (Supporting Information) based on database search. As shown in Figure 3, 13 peptides were observed both for H-DMP- and D-DMP-labeling as compared to 10 native peptides detected without N-phosphorylation in the same mass range from m/z 700 to 2500. It is worth noting that all of the signals are protonated species and no metal ion adducts are found. Three additional tryptic peptides were observed at m/z 822.5003, 1090.5111, and 1265.6172 for HDMP-labeling and at m/z 834.5506, 1102.5773, and 1277.6785 for D-DMP-labeling, respectively, indicating that there were two dimethyl phosphoryl group attached to each peptide because of the 12 Da mass shifts between H-DMP- and D-

DMP-labeling. Indeed, the newly observed peaks were determined by nano ESI-MS/MS, and the sequences of the peptides were identified as KVFGR (13), HGLDNYR (12), and CELAAAMKR (11), respectively (Figure S-4, Supporting Information). Compared to the peptide sequence coverage found from the unlabeled peptide digest, the N-phosphorylation labeling can increase the coverage by 15% from 61% to 76% searched by Mascot. The N-phosphorylation labeling of several other test proteins with dimethyl phosphite showed that the labeling reaction was highly efficient. For example, the sequence coverage of BSA with the higher molecular weight than lysozyme could also be increased by 13% from 41% to 54% (Figure S-5, Supporting Information). It is worth noting that all N-terminal primary amines can be labeled quantitatively with H-DMP or D-DMP reagents in 40 min by a one-pot reaction under mild reaction conditions. In most cases of protein digestion, some interfering components, such as the primary amine-containing molecules, needed to be removed by desalting the samples prior to labeling or by performing the digestion in buffers without primary amines. However, under our conditions, the ammonium bicarbonate remaining in the reaction mixture after in-solution protein digestion did not affect the labeling efficiency, indicating a significant improvement in method development with the DMP-labeling. The peak of the unmodified peptide disappeared completely in the spectra of labeled lysozyme and BSA digests. Furthermore, the introduction of the neutral phosphoryl group with high proton affinity might not only provide improved ionization efficiency and concomitant signal enhancement for the ESI-MS analysis of the labeled compounds but also increase their retention capability on the reversed-phase C18 column. For example, the peak intensity of the [M + H]+ ion of HDMP-labeled fibrinopeptide A peptide is increased about 50 10239



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Figure 4. Nano ESI-MS/MS of tryptic peptide CKGTDVQAWIR derived from lysozyme: (A) MS/MS spectrum of native peptide (2+, m/z 667.3344); (B) MS/MS spectrum of H-DMP-labeled peptide (2+, m/z 775.3282).

times than that of unlabeled peptide in nano ESI-MS analysis under the same conditions (Figure 2, parts A and B). For MALDI-MS analysis, the signal intensity of most labeled small peptides was slightly enhanced as compared to that of the native peptides. Tryptic peptide KVFGR derived from lysozyme could not be detected without N-phosphorylation labeling. However, after the incorporations of two phosphoryl groups at the N-terminus and side chain amino group of lysine, a very intensive peak corresponding to KVFGR was observed due to two possible reasons. First, the labeled peptides might be easier to be trapped on a desalting C18 column (SPE using

ZipTips) than native peptides because two primary amine groups that could form the salt under acidic desalting conditions have been protected by phosphoryl groups, which can increase the retention capacity of the peptides. Second, the existence of phosphoryl groups could significantly improve the ionization efficiency in ESI-MS analysis because of its intrinsic high surface affinity for the surface of ESI droplets and proton affinity. In addition, the hydroxyl group of tyrosine could also be derivatized dependently on the peptide structure as a side reaction, which might increase the complexity of the peptide mixtures but provide some useful information for peptide 10240



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Figure 5. (A) Extracted ion chromatogram for a linearity assay of H-DMP- and stable isotope D-DMP-labeled fibrinopeptide (A) standard peptide ADSGEGDFLAEGGGVR (2+, m/z 822.82 and m/z 825.83) under various ratios (1:0.5, 1:1, 1:2, 1:3, 1:4, 1:5, 1:10, and 1:20). (B) The linearity plot of the H-DMP- and D-DMP-labeled peptide ADSGEGDFLAEGGGVR. The ratios are plotted by their expected values on the x-axis and their measured ratios on the y-axis. A red diagonal line indicates the ideal distribution. Every value was analyzed in triplicate. The error bars denote the minimum and maximum range of the data acquired from three individual experiments.

more, a mass shift of 216 Da was observed for b ions but not for y ions, which is consistent with the fact that two dimethyl phosphoryl groups are attached to the N-terminus and ε-NH2 of the lysine residue, respectively. Interestingly, in the MS/MS spectrum of the deaminated peptide CKGTDVQAWIR (2+, m/z 712.3379),58 the b ions were not enhanced because of the loss of the N-terminal amine group of cysteine residue resulting in only one phosphoryl group attached at the ε-NH2 of the lysine residue, which suggested that the N-terminal phosphoryl group played an essential role on the fragmentation pathways of peptides (Figure S-6, Supporting Information). The b ions could also be enhanced by N-phosphorylation labeling even through the sites of phosphoryl group labeling were at the Cterminus of the peptides, such as the H-DMP-labeled HGLDNYR (2+, m/z 547.7138) and CELAAAMKR (2+, m/ z 633.2594) (Figure S-4, parts B and C, Supporting Information). In contrast to the doubly charged ions, the y ions did not exist or were weak in the MS/MS of singly charged KVFGR peptide, while the yields of a and b ions were greatly increased (Figure S-4A, Supporting Information). Major advantages of N-phosphorylation labeling included the great enhancement of b ions and reduction of the complexity of the MS/MS fragmentation patterns of peptides, which is very helpful for interpreting ESI-MS data of peptides with completely unknown amino acid sequence. The incorporation of the neutral phosphoryl group might increase the relative proton affinities (PA) of the modified N-terminus, which was important for determination of the relative abundance of b and y ions formed through a charge-direct fragmentation mechanism.59 According to the “pathways in competition model”,60 the N-terminal phosphoryl group may compete with other basic sites, such as arginine, histidine, or lysine, to localize the charges favorably. For singly charged peptides in which the charge may be mainly retained at the N-terminal phosphoryl group, the C-terminal fragments will be neutral. Thus, a relatively high abundance of b ions may be formed since the ytype cleavage is suppressed. On the other hand, for multiply charged peptides, y ions may be formed because C-terminal fragments will be charged due to the remaining charge at the Cterminal basic residues in the sequence. In conclusion, by using the N-phosphorylation labeling method discussed above, the peptide sequence can be rapidly extrapolated from the

identification and improve intensity. Overall, N-phosphorylation labeling based on phosphorus chemistry with quantitative yield, one-pot reaction, minimal sample preparation, and the enhanced ESI-MS signal intensity provided advanced performance in derivatization of the tryptic peptide samples. Peptide de Novo Sequencing. Several unique advantages of N-phosphorylation labeling for de novo peptide sequencing were demonstrated using some small synthetic peptides with different structures. Specifically, for singly charged peptides, the N-phosphorylation labeling approach can significantly enhance a and b ions series that are usually missing in MS/MS spectra and simplify the MS/MS fragmentation patterns, thus providing more confident sequence assignment and facilitating peptide identifications. Furthermore, it can be used to effectively distinguish the isobaric amino acid residues glutamine (Q, 128.0585 Da) versus lysine (K, 128.0949 Da) and differentiate the overlapping masses with one amino acid from the combination of two amino acids, such as glycine and alanine (G + A, 128.0585 Da). Although N-phosphorylation labeling appears to be promising for de novo sequencing, it is needed to be further validated using tryptic peptide mixtures of protein samples. Unlike the synthetic model peptides being singly charged in ESI-MS analysis, the peptides derived from protein digests might contain basic residues of either arginine or lysine at the C-terminus and thus be multiply charged easily, which might have a significant effect on the fragmentation pathways of Nphosphoryl labeled peptides. As an example, Figure 4 shows MS/MS sequencing of tryptic peptide CKGTDVQAWIR derived from lysozyme by nano ESI-MS analysis. The native peptide displayed intensive y ions and complex fragment ions with low abundance in the mass range of m/z 100−700 (Figure 4A). In contrast, the incorporation of phosphoryl group resulted in a much cleaner fragmentation spectrum as shown in Figure 4B. The b ion series were greatly enhanced, providing complete sequence coverage despite the presence of a Cterminal arginine that strongly favors the formation of y ions. It is also worth noting that most of the small fragments in the low-mass range are disappeared as well as the two intense ions shown in Figure 4A; namely, the y1−H2O−CO ion at m/z 129.0993 and y2−NH3 ion at m/z 271.1187, were not observed in Figure 4B after the N-phosphorylation labeling. Further10241



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Table 1. Quantification of the Protein Mixture under Various Ratios by Nano LC-Chip/TOF MS ratio protein BSA

β-casein

hemoglobin

β-lactoglobulin

lysozyme

myoglobin

peptide position

peptide sequence

charge state

no. of labeling

exptl

obsd

205−209 360−371 469−482 48−63 192−198 199−217 32−41 97−105 134−145 108−117 158−164 165−178 52−63 80−86 135−143 2−17 18−32 135−140

IETMR RHPEYAVSVLLR MPCTEDYLSLILNR FQpSEEQQQTEDELQDK AVPYPQR DMPIQAFLLYQEPVLGPVR LLVVYPWTQR LHVDPENFR VVAGVANALAHK VLVLDTDYKK ALPMHIR LSFNPTQLEEQCHI FESNFNTQATNR WWCNDGR GTDVQAWIR GLSDGEWQQVLNVWGK VEADIAGHGQEVLIR ALELFR

1 3 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 1

1 1 2 1 1 1 2 1 1 2 1 1 1 1 1 1 1 1

1.00 1.00 1.00 2.00 2.00 2.00 10.00 10.00 10.00 4.00 4.00 4.00 0.50 0.50 0.50 1.00 1.00 1.00

1.10 1.05 0.90 2.04 2.12 1.82 9.65 8.74 10.31 4.12 3.53 3.54 0.44 0.53 0.43 1.02 0.95 0.91

mean ± SD

error (%)

1.01 ± 0.10

1.0

1.99 ± 0.15

0.5

9.57 ± 0.78

4.3

3.73 ± 0.33

6.7

0.47 ± 0.05

6.0

0.96 ± 0.05

4.0

level and accuracy might be obtained if more tryptic peptide pairs could be selected for quantification. Quantification of Protein Mixtures by Nano LC-Chip/ TOF MS. To test the applicability of the stable isotope Nphosphorylation labeling for quantification in a complex sample, the relative abundances of six purified proteins in two samples were analyzed by using full-scan MS, and the sequencing for protein identification was obtained based on data-dependent LC/MS/MS on a nano LC-chip/TOF MS system and a database search. Two tryptic digests of the six standard proteins were combined in different amounts and different ratios to give two sets of samples, in which the protein abundance ratios (H-DMP- and D-DMP-labeling) between the mixtures were 1, 2, 10, 4, 0.5, and 1 for BSA, β-casein, myoglobin, β-lactoglobulin, lysozyme, and hemoglobin, respectively. Three peptide pairs, including double and triply charged ions, were selected manually to calculate peptide ratios for each protein (Figure S-8, Supporting Information). Meanwhile, the means and standard deviations of peptide ratios were used to quantify proteins in the mixture. As shown in Table 1, the N-phosphorylation labeling strategy was able to determine the relative abundance ratios of the six target proteins in the sample and showed excellent correlation of the measured ratios with theoretical ratios with errors that range from 0.5% to 6.7% and relative standard deviation of less than 10.6%, which were sufficient for many proteomic applications. Additionally, the isotopic mass pair of phosphopeptide FQpSEEQQQTEDELQDK (2+, m/z 1085.4145 and 1088.4350) with one phosphoric acid group on the serine residue was selected for the quantification of β-casein as the phosphorylated protein (Figure S-8B, Supporting Information). It was found that the measured ratio (H-DMP-/D-DMP-, 1:2.04) precisely reflected the true ratio (1:2.00) and no side reactions occurred at the phosphoryl group on the serine residue during labeling, indicating that this labeling strategy could also be used for the analysis of posttranslational protein phosphorylation. In addition, all N-terminal primary amine could be first labeled quantitatively with H-DMP or D-DMP reagents under mild reaction conditions. N-Phosphorylation labeling of the side chain amino group of the lysine residue and

interpretation of MS/MS spectra. Thus, the developed method may be a useful tool for the de novo peptide sequencing. Quantification of Standard Peptide and Protein. In order to investigate the feasibility of N-phosphorylation labeling for quantitative analysis, fibrinopeptide A was analyzed by using LC/ESI-TOF MS (Figure 5). Two solutions of the peptide standard with different concentration ratios within a linear range from 1:0.5 to 1:20 were labeled using H-DMP and DDMP, respectively. The samples were mixed and analyzed by LC/MS in triplicate. One challenge associated with stable isotope labeling is the presence of peptides overlapping that may contribute to quantitative errors. In the ESI-MS analysis, the peptide ADSGEGDFLAEGGGVR with one labeled site at the N-terminus (6 mass units for each labeling) was doubly charged. However, the light-labeled (2+, m/z 822.82) and heavy-labeled (2+, m/z 825.83) peptides were completely resolved due to the detected mass difference of 3 Da. The average from the three experimentally obtained ratios was plotted against their expected ratios (Figure 5), indicating that the N-phosphorylation labeling method was linear across a 20fold range of concentration ratios from 0.5 to 20 with an R2 value of 0.9984. Furthermore, the relative standard derivations were in the range of 2.5−9.4%, implying that the labeling method was reproducible and precise for peptide quantification. Furthermore, lysozyme was studied as a protein model for validation of the SIPL method for the relative protein quantification (Figure S-7, Supporting Information). Two sets of samples containing different ratios (1:0.5, 1:1, 1:2, 1:3, 1:4, 1:5, and 1:10) of the tryptic peptides were labeled with lightand heavy-labeled phosphorus reagents, respectively. The lightand heavy-labeled samples were mixed together and analyzed by using LC/ESI-MS. Two tryptic peptides, namely, FESNFNTQATNR (2+, one labeling, 3 Da of mass shift) and WWCNDGR (1+, one labeling, 6 Da of mass shift), were selected manually as examples to estimate the linearity of this method. The measured ratios of the light- and heavy-isotopic pairs were consistent with the theoretical ratios, resulting in a linear dynamic range with R2 values of 0.9951 and 0.9919, respectively. Rationally, it is believed that higher confidence 10242



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hydroxyl group of tyrosine residue might be dependent on the reaction conditions, amounts of phosphorus reagents, and chemical structures of the tryptic peptides. The overall data demonstrated that differential N-phosphorylation labeling could be applied to quantify tryptic peptides present at different concentration ratios within the biological sample for quantitative proteomics. Validation of the Deuterium Atom Effect of Labeled Peptides. Deuterium labeling has many attractive features, such as being less expensive and simpler for preparing coding agents and allowing smaller coding agents. However, it is known that the deuterium atom is slightly more hydrophilic than the hydrogen atom. Thus, separation of the light- and heavy-labeled peptides in reversed-phase chromatography is often observed, which may affect the accuracy of quantitative analysis. However, it has been reported that the deuterium effect can be minimized when the deuterium atoms are labeled on polar functional groups.61 To examine the chromatographic elution profiles of both H- and D-labeled N-phosphorylated labeled peptides, two small peptides, KVFGR and HGLDNYR, derived from lysozyme with 12 deuterium atoms labeled on the two phosphoryl groups were investigated (Figure S-9, Supporting Information). It was found that the extracted chromatograms of the D12-DMP- and H12-DMP-labeled KVFGR ions at m/z 834.4260 and 822.3497 almost coeluted with the comparative signal intensity at retention times of 16.28 and 16.32 min, respectively, indicating that the isotope effect from the deuterium-labeled peptide was minimal. Even through a small isotope effect would be observed, the quantitative ratios could be determined based on the integration of the entire extracted ion peaks of the heavy and the light peptides. Furthermore, the isotope effect may be neglected for the labeled peptides with higher molecular weight and small number of deuterium atoms because of two possible reasons. The first is that the coding agent size of the dimethyl phosphate group is moderately small, which is considered to be correlated inversely with the magnitude of the isotope effect.61 The second possible reason is that the deuterium atoms are placed adjacent to the neutral phosphoryl group with higher electrophilicity, which may minimize the hydrophilicity difference between the deuterium atom and hydrogen atom under acidic separation conditions and, therefore, reduce the differential interactions with the reversed-phase chromatography that leads to chromatographic resolution.62

efficiency for peptide mass fingerprints but also greatly enhance the intensities of b ions and reduce the complexity of the MS/ MS fragmentation patterns of peptides, which allows more confident peptide de novo sequencing and protein identification. N-Terminal phosphorylation adds a moderately hydrophobic phosphoryl group that may increase retention on the stationary phase of reversed-phase chromatography, which can increase the detection capability of short or polar tryptic peptides that might otherwise elute near the void volume and escape detection. Third, the labeling reagent is inexpensive compared to other current labeling reagents and can be easily synthesized by a one-step reaction at high yield with a commercially available isotope reagent. Finally, the Nphosphorylation labeling tags have small molecular weight of 108 Da with simple chemical structures, which may not generate fragments interfering the MS/MS sequencing. Additionally, the isotope effect was insignificant in most cases even though six deuterium atoms were incorporated for each labeling. The isotope effect might be further minimized by introducing other isotope atoms, such as 13C, 15N, and 18O, into the coding reagents. Further investigations may be conducted to design series of phosphorus reagents with different chemical structures as well as novel functions and to optimize the coupling conditions of the reactions in order to improve the applicability of the N-phosphorylation labeling for quantitative proteomics.

CONCLUSIONS We have successfully developed a novel stable isotopic Nphosphorylation labeling method for peptide de novo sequencing and protein quantification based on organic phosphorus chemistry. The N-phosphorylation labeling reaction was performed easily within 40 min in a one-pot reaction without additional cleanup procedures reducing sample handling and thus the potential of experimental errors. The new approach has several advantages. First, the reaction is a global labeling with quantitative yield and high selectivity. The primary amine group of the N-terminus of peptides may be labeled efficiently. The labeling reagent is more stable than most of the other stable isotopic labeling reagents, in which the tags need to be activated in advance. By changing the reaction conditions, the phosphite may be activated in situ to form a labeling intermediate with high activity for peptide coupling. Second, the N-terminal labeled phosphoryl group with high relative proton affinity may not only improve the ionization

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ASSOCIATED CONTENT

S Supporting Information *

Additional information as noted in text. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected] (Z.C.); [email protected]. cn (Y.J.). Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors acknowledge financial support from the National Natural Science Foundation of China (Nos. 20928005 and 21172129) and the Ministry of Science and Technology of China (2011DFA30620).

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REFERENCES

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Stable Isotope N-Phosphorylation Labeling for Peptide de Novo

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