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A synthesis of new, bi-labeled peptides for quantitative proteomics Maciej Modzel, Halina Płóciennik, Martyna Kielmas, Zbigniew Szewczuk, Piotr Stefanowicz⁎ Faculty of Chemistry, University of Wroclaw, F. Joliot-Curie 14, Wroclaw, Poland



Article history:

Isotopically labeled peptides are often used in proteomics as internal reference allowing

Received 8 August 2014

quantification of peptides by isotopic dilution method. Although the synthesis of peptides

Accepted 1 December 2014

labeled with stable isotopes is relatively simple, there are several factors limiting application of these standards in proteomic research: cost of labeled derivatives of amino acids, time needed to obtain labeled peptide and problems with quantification of the


standard. To solve these problems we developed a method of synthesis of peptides labeled

Isotopic labeling

with heavy oxygen and with a dabsyl moiety. The chromophoric group facilitates the

Quantitative proteomics

determination of peptide concentration while sequence of peptide allows enzymatic

Isotopic dilution

cleavage of fragment containing dabsyl from peptide leaving “natural” sequence with




O atoms. The approach proposed herein is based on the “analytical

construct” concept. The experiments performed on model peptides demonstrated that response factors in HPLC analysis of labeled peptides does not depend on the sequence and tryptic hydrolysis of obtained conjugates is completed in minutes producing labeled standards useful in quantitative proteomics. Biological significance The reported method allows for a cheap and efficient synthesis of peptides labeled with heavy isotopes, and for their precise quantification. Peptides of our design are stable, and the isotopic label, which is a part of the peptide backbone, is stable as well. Moreover, they can be quickly quantified in solution at any time, so the possible decomposition of standard or a non-uniform distribution of the peptide in lyophylisate does not pose a problem. Therefore, we deem our synthesis to be useful for a broad range of quantitative proteomics methods. In addition, the procedure described herein allows direct application of crude peptides as the analytical standards. The elimination of expensive and time-consuming chromatographic purification reduces the cost of AQUA peptides and gives the possibility of a rapid preparation of large libraries of proteolytic fragments. © 2014 Elsevier B.V. All rights reserved.

Abbreviations: Dab, dabsyl moiety; TCTU, O-(6-chlorobenzotriazol-1-yl)-N,N,N′,N′-tetramethyluronium tetrafluoroborate, Fmoc, fluorenylomethyloxycarbonyl group; DIEA, diisopropylethylamine; SPPS, solid-phase peptide synthesis. ⁎ Corresponding author at: Faculty of Chemistry, University of Wroclaw, F. Joliot-Curie 14, 50 383 PL Wroclaw, Poland. Tel.: +48 71 375 7213. E-mail address: [email protected] (P. Stefanowicz). 1874-3919/© 2014 Elsevier B.V. All rights reserved.


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1. Introduction Stable isotope labeling of proteins and peptides is useful in quantitative proteomics [1]. Such peptides can be used as internal references when assessing the concentration of a sample by mass spectrometry, for example, using multiple reaction monitoring (MRM) method [2,3]. The following stable isotopes: 2H, 13C, 15N, and 18O are often used for this purpose [1,4,5]. Labeling with 18O has several significant advantages over the other isotopes. Every introduced atom increases the mass by 2 and incorporation of 18O in vitro is easier than 13C or 15 N. On the other hand, unlike deuterium, heavy oxygen does not induce a change in the retention time [6]. Several protocols for 18O labeling in vitro have been published. They can be divided into two main categories: enzymatic [7,8] and acid-catalyzed [9–11]. Isotopically labeled peptides may be also obtained using commercially available amino acid derivatives containing 13C or 15N atoms. Therefore, from synthetic point of view, the preparation of labeled peptides does not pose a problem. However, there still remains the problem of precise determination of the quantity of the standard especially if only a limited amount of sample is available. In case of peptides, it is not possible to determine the amount of the peptide by simply weighting it, as the lyophylisate contains counter ions and possibly also a large amount of solvent [12]. Moreover, the peptides in question usually need to be purified, thus increasing the cost of the analysis, and their concentration can change over time due to decomposition, introducing an additional source of error [13]. Even if the concentration in the dry product is determined, sometimes complete dissolution of the peptide might not be possible, and therefore the concentration in solution does not correspond to that calculated for the amount of peptide used. A quantification of peptides is often performed by the method of amino acid analysis, which has been described as the “golden standard” [12]. However, the error of this method sometimes exceeds 10% [14]. In addition, amino acid analysis requires a relatively large sample of pure compound. The synthesis of labeled substances at a large scale is expensive, so the quantification of the standards by amino acid analysis can be prohibitively expensive. On the other hand, milligram quantities of labeled peptide may be sufficient to perform thousands of analyzes. It is clearly visible that a new analytical method for absolute quantification of peptides in solution is desired. We have, therefore, decided to develop an approach based on the concept of analytical construct [15,16]. This term was introduced by Geysen et al. [17]. The strategy relies on two orthogonal linkers within the synthesized compound: one links the proper compound to an “analytical enhancer,” the other one links the whole compound to the resin. Such constructs have found their use mainly in the analysis combinatorial libraries when each of many compounds was synthesized in a very small amount. Therefore, the construct contained an analytical enhancer, for example, a chromophore group for UV detection, or a quaternary ammonium salt for MS analysis. Depending on the cleavage conditions, either the whole construct, containing the enhancer, could be cleaved, or just the desired compound without the enhancer.

The general structure of analytical constructs described herein is depicted in Fig. 1. They are composed of the following elements: the sequence corresponding to the tryptic peptide, cleavable linker and chromophore. The desired peptide, which has the same sequence as a peptide derived from a protein of interest during tryptic digestion, is attached directly to a Wang resin. This part is labeled with heavy oxygen. Then at the N-terminus of this part, a linker containing a cleavage site for trypsin is attached, followed by dabsyl moiety at its N-terminus. The idea of the experiment is also shown in Fig. 1. Briefly, the synthesized peptide, labeled with 18O, is quantified by HPLC, mixed with the protein solution and subjected to enzymatic hydrolysis. As dabsyl moiety is cleaved together with the linker, a shorter isotopically labeled peptide, the analytical standard, is liberated. The whole mixture is then subjected to LC-MS analysis.

2. Experimental section 2.1. Materials and reagents 1,4-Dioxane was purchased from POCh (Gliwice, Poland), concentrated sulfuric acid (96% H2SO4) from Stanlab (Lublin, Poland), sodium chloride from Chempur (Piekary Śląskie, Poland), TFA from Merck (Darmstadt, Germany) and Fmoc-protected amino acids and preloaded Wang resins (0.54–0.77 mmol/g) from NovaBiochem (Merck, Darmstadt, Germany). All the remaining chemicals, including 18O labeled water (97% 18O), were purchased from Sigma-Aldrich (St. Louis, MO, USA). Water was purified using a Hydrolab purification system (Hydrolab, Poland).

2.2. Peptide synthesis Safety warning: as coupling reagents are toxic and potentially carcinogenic, gloves and protective glasses must be worn for peptide synthesis. Peptides were synthesized by standard SPPS Fmoc strategy on Wang resin, as described [18]. The synthesis was manually performed in polypropylene syringe reactors (Intavis AG) equipped with polyethylene filters. The coupling reactions were performed in DMF, with TCTU (3 Eq) in presence of DIEA (6 Eq). The Fmoc protecting groups were removed with 25% piperidine solution in DMF. The sequences chosen were model peptides basing on sequences of tryptic peptides of human serum albumin (HSA) and fibrinogen.

2.3. Fmoc-amino acid labeling procedure The conditions for these reactions have been published previously [19]. Briefly, 5 mg of the Fmoc-protected amino acid derivatives were dissolved in 2 M HCl in dioxane, with 5% H18 2 O and either left overnight, or treated with microwaves (110 °C) for 15 min. The solvent was then evaporated in a stream of dry nitrogen. Then the protected amino acids were used directly for regular solid-phase peptide synthesis with no purification.

2.4. Chromophore tagging Dabsyl: 2 Eq of dabsyl chloride was dissolved in 2 ml of DMF, and 4 Eq of DIEA was added. The mixture was added to a

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Fig. 1 – The scheme of the experiment: a solution of the construct is subjected to HPLC analysis to determine its quantity (A). A known amount of that solution is then mixed with a solution of the protein and subjected to enzymatic hydrolysis (B), and then to LC-MS analysis ©.

syringe reactor containing the peptide and left rotating for 48 h. Methyl red: the coupling conditions were the same as for a regular Fmoc-protected amino acid derivative. DNP: 10 Eq of dinitrofluorobenzene together with 2 Eq of DIEA was dissolved in 2 ml of DMF. The mixture was added to a syringe reactor containing the peptide and left rotating for 48 h.

2.5. Purification Before NMR measurements, peptides were purified by preparative HPLC on a Varian ProStar, column—Tosoh Biosciences ODS-120 T C18 (ODS 300 × 21.5 mm). Water–acetonitrile gradients containing 0.1% TFA at a flow rate of 7 ml/min were used for purification with UV-Vis detection at 210/473 nm.

2.6. MS analysis The measurements were conducted on an Apex-Qe Ultra 7 T instrument (Bruker Daltonics, Bremen, Germany) equipped with a dual ESI source and a heated hollow cathode dispenser. The instrument was operated in the positive-ion mode and calibrated with the TunemixTM mixture (Bruker Daltonics). The mass accuracy was within 5 ppm. For the measurement, 1 mg of the sample was dissolved in 1 ml of H2O:MeCN: HCOOH 50:50:1 mixture and then diluted 100 times and injected to the ion source at a rate of 3 μl/min. Analysis of the obtained mass spectra was carried out using a Biotools (Bruker Daltonics) software. The instrumental parameters were as follows: scan range, 100–1600 m/z; dry gas, nitrogen;


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temperature, 200 °C; potential between the spray needle and the orifice, 4.2 kV.

2.7. NMR analysis

over 62 min (flow rate 0.05 ml/min, room temperature). Data Analysis 4.0 program (Bruker, Germany) was used for generation of mass list and extracted ion chromatograms. The mass accuracy was within 10 ppm.


H NMR spectra were recorded on high-field spectrometer (Bruker Avance III 500 MHz, Bremen, Germany) equipped with broadband inverse-gradient probe heads. Spectra were referenced to the residual solvent signals (methanol-d6: δ = 3.31, 4.78 ppm). Two-dimensional NMR spectra were recorded with 2048 data points in the t2 domain and up to 1024 points in the t1 domain, with a 1 s recovery delay. The samples were prepared by weighting 0.4–1.3 mg of the peptide or dabsylated alanine and dissolving them in 600 μl dioxane:deuterated methanol 0.15:1000 v:v mixture. Right after the NMR measurements, the samples were transferred to HPLC vials and subjected to HPLC analysis.

2.8. HPLC analysis The analysis was performed on Dionex chromatography system equipped with Dionex ICS-Series Variable Wavelength Detector 064654. The column was a Jupiter 4 μ Proteo 90A, 250 × 4.6 mm. The eluents were as follows: A—0.1% TFA in H2O; B—0.1% TFA in H2O:MeCN 20:80 v:v mixture. The gradient program used was as follows: from 100% A to 80% B in A in 40 min.

2.9. Proteolytic digestion The enzymatic hydrolysis was performed as described previously by Kijewska et al. [20] Briefly, HSA (3.07 mg) and/or the dabsylated peptide (1.23 mg) was dissolved in NH4HCO3 buffer solution (3 ml, 10 mM, pH 8.0). After the addition of dithiothreitol (45 μl, 45 mM in water), the mixture was incubated at 50 °C for 15 min. The final concentration of dithiothreitol in the mixture was approximately 0.67 mM. Then two samples were prepared: one consisted of 1 ml of HSA solution and 100 μl of peptide solution, the other consisted of 200 μl of peptide solution. Aliquots of trypsin stock solution (1.5 mg and 500 μl in water) were added to the samples: 100 μl to the first and 20 μl to the second. The reaction mixtures were incubated for 24 h at 37 °C. The reaction was quenched by the addition of 10% aqueous trifluoroacetic acid (TFA, 240 μl). The resulting digest was lyophilized and used for LC-MS experiments.

2.10. LC-MS analysis The LC–MS analysis was performed in the Laboratory of Mass Spectrometry at the Faculty of Chemistry, University of Wroclaw, using an Agilent 1200 HPLC system coupled to a micrOTOF-Q mass spectrometer (Bruker Daltonics, Germany). The micrOTOF-Q instrument equipped with an ESI source with ion funnel was operated in positive-ion mode and calibrated before each analysis with the Tunemix™ mixture (Bruker Daltonics, Germany) in a quadratic method. For separation, an Aeris PEPTIDE, Phenomenex (50 × 2.1 mm, 3.6 μm) column was used, with gradient elution of 0–100% B in A (A, 0.1% HCOOH in water; B, 0.1% HCOOH in acetonitrile)

3. Results and discussion To develop a simplified method of the preparation of isotopically labeled peptides of a defined concentration, we solved the following problems: 1 The synthesis of doubly labeled analytical construct containing peptide sequence (with isotopic label), spacer susceptible to enzymatic hydrolysis and chromophore. 2 Testing the series of model peptides to check if response factor in HPLC experiment is sequence independent 3 Testing the chromatographic properties of the hydrolyzed construct.

3.1. Synthesis of labeled peptides In order to obtain a labeled construct, we have used a previously described method of labeling Fmoc-protected amino acids. This method bases on using a mixture of 2 M HCl in dioxane with H18 2 O (95:5, v:v), either for 24 h or for 15 min, but with microwave assistance (110 °C). Thus, we can obtain peptides regioselectively labeled—the label is at the carboxyl group of the labeled amino acid. While not all of the amino acids can be easily labeled using this protocol—only those that do not require side chain protection—a majority of tryptic peptides obtained from proteins contain at least one such amino acid.

3.2. Chromophore tagging and construct susceptibility to enzymatic hydrolysis We have tested three reagents suitable for introducing a chromophoric groups: methyl red, fluorodinitrobenzene and dabsyl chloride. Tagging with methyl red followed a standard protocol for coupling an amino acid to peptide on solid support, as this reagent possesses a carboxyl group. However, the efficiency of labeling was low; therefore, only the remaining two tags were selected for further analysis. The tagged peptide contained alanine and lysine moieties as the two N-terminal amino acids, and the tagging was carried out on the N-terminal alanine. After cleavage of labeled peptides from the resin, they were purified by preparative HPLC. The examples of HPLC chromatograms for crude peptides are presented in supplementary data (Fig. 6 and Fig. 7). The homogeneity and identity of obtained products were tested by analytical HPLC and HRMS. Analytical data are presented in Table 1. We have noticed that dabsylated peptides usually have a retention time between 18 and 26 min. We have also checked the difference of the retention time for several peptides and their dabsylated analogues—in all the investigated cases, the retention time was significantly extended. The comparison of retention times for peptides and their dabsylated analogues is presented in the supplementary material Table 3. Even if some by-products are observed, they are usually well resolvable, and purification of the peptides is

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possible (see supplementary data Fig. 5)—a more detailed study concerning the detection of dabsylated peptides confirms this observation [21]. To test susceptibility to enzymatic hydrolysis, the labeled peptides were dissolved in ammonium bicarbonate solution and trypsin was added. TFA was then added to quench the reaction, and the reaction mixture was analyzed by HPLC and MS. Both methods proved that the hydrolysis of dabsylated peptides was quantitative within up to several minutes, while peptides tagged with DNP moiety did hydrolyze, albeit much slower. We have also tried to shorten the construct by removing the N-terminal alanine and dabsylating the N-terminal lysine α-amino group, but such peptides proved resilient to hydrolysis. Therefore, only dabsyl was used to tag peptides in the further experiments and constructs which were to be hydrolyzed had dab-Ala-Lys fragment at their N-termini. For sample chromatograms of dabsyl-tagged peptides subjected to enzymatic hydrolysis (hydrolysis performed according to experimental section, the incubation time 1 h), see Supporting Information Figs S8, S9 and S10.

3.3. Quantitative analysis We then concentrated on quantitative analysis of dabsylated peptides to test, if the response factors in HPLC analysis for various peptides (sequences and experimental data provided in Table 1) depend on their sequences. However, it required an another independent method for quantification of the peptides. NMR was selected as the reference method. The measurements were performed in deuterated methanol, as it dissolves the peptides well and is compatible with HPLC. Therefore, no solvent evaporation step is necessary, and the sample used for NMR experiment can be directly used for HPLC analysis, thus reducing the potential for errors. At first, we analyzed the NMR spectrum of dabsylated alanine (the structural formula of dab-Ala and its 1H NMR spectrum is presented in supplementary data Fig. 3 and Fig. 4) in order to find signals, which could be used for quantitative analysis. The peak of dimethylamine group was selected as the best candidate—its signal is a singlet, well separated from all the other peaks. Its identity was confirmed by integration (relative to the signals in the aromatic range, corresponding to 8 protons total) and by analyzing COSY spectrum. We have then prepared a stock solution of deuterated methanol with dioxane (0.15 μl/ml) and subjected a series of peptides to analyzys by NMR. Between 0.5 and 1.5 mg of lyophilisate of each peptide was dissolved in 750 μl of “spiked”


methanol, and 1H NMR spectra were recorded. The ratio of areas of peak corresponding to dimethylamino group to that corresponding to dioxane was denoted as “NMR concentration.” The solution was then directly used for HPLC analysis. The total area under the peaks registered at 473 nm was denoted as HPLC concentration. The results were plotted on a chart and a linear regression line was fitted to the experimental points. The correlation coefficient was good (0.97). Assuming that a perfect calibration curve must pass through the origin we have simplified its equation to the form of y = ax and calculated the “a” parameter (slope) for all the peptides— the mean error was approximately 6% (all the statistical data are provided in the Supplementary Information). This proves that the response factor for the dabsyl labeled compound does not depend on the peptide sequence, and therefore dabsyl is a suitable tag for quantitative analysis of peptides by HPLC. In addition to that to allow an easy transfer of analytical procedure between different HPLC systems, we have run the same experiment on various amounts of commercially available dabsylated alanine. The mean slope for the points representing these samples is very similar to mean slope obtained for the whole set of peptides. The combined results for the peptides and the various samples of dabsylated alanine are presented in Fig. 2. It is worth noting that the R2 coefficient exceeds 0.97, and the regression line passes very close to the origin. Moreover, a two-tailed Student's t-test performed on the slope coefficients calculated for the set of peptides and separately, for several samples of dabsylated alanine proved that there is no statistically significant difference between them (p score of 0.5). The quantitative data are presented in Tables 1 and 2, supplementary data. Therefore, the proposed protocol for analysis using our method is to either prepare an HPLC calibration curve using known weighted amounts of dabsylated alanine, or to mix a known amount of dabsylated alanine with the peptide sample before HPLC analysis and assess the concentration of the peptide basing on the relative areas of peaks corresponding to the peptide and to dabsylated alanine.

Table 1 – Experimental data for peptides. Peptide sequence dab-AKAAR dab-AKTGK dab-KRGF dab-AK dab-AKALAR dab-TGK dab-AKASSAK dab-AKVVER dab-AKGGGW dab-GDKKA

m/z calc 402.2032 396.1976 397.6925 505.2233 458.7452 592.2553 475.2322 988.5038 862.3670 805.3666

[M [M [M [M [M [M [M [M [M [M

+ + + + + + + + + +

H]+ 2H]2+ 2H]2+ H]+ 2H]2+ H]+ 2H]2+ H]+ H]+ H]+

m/z exp

Retention time (min)

402.2035 369.1967 397.6963 505.2287 458.7435 592.2533 475.2321 988.4992 862.3680 805.3624

22.44 21.44 24.03 23.92 25.06 22.84 21.36 24.45 23.16 21.73

Fig. 2 – The comparison of HPLC and NMR data obtained for peptides and dab-Ala. The separate plots for dab-Ala and set of dabsylated peptides are presented in supplementary data (Fig. 1 and Fig. 2).


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3.4. LC-MS analysis The final goal was to investigate the chromatographic properties of the labeled peptide enzymatically liberated from the construct. For our research, we have utilized labeled Fmoc-alanine, which was introduced to ADLAK peptide. This is a peptide resulting from performing a tryptic cleavage of HSA protein. Both alanines were substituted by their isotopologues, and additionally the peptide was elongated by a lysine, an alanine and a dabsyl moiety at the N-terminus. The final sequence was dabsyl-AKA*DLA*K, where asterisk denotes a labeled alanine. The label is inside the peptide chain so as to prevent back exchange when the peptide is subjected to hydrolysis conditions [22]. Alanines of isotopic purity of approximately 90% were used, and the isotopic purity of the peptide itself is 86%. Two samples—one consisting of just the construct, the second of a mixture of the construct and HSA—were subjected to hydrolysis catalyzed by trypsin to liberate the ADLAK peptide, both in its labeled and non-labeled form. After quenching the reaction with TFA, the solutions were lyophilized, resuspended in water and subjected to LC-MS analysis. The extracted ion chromatograms presented in Fig. 3 prove that the peptide cleaved from the construct retains its label, and that the two isotopologues have the same retention time. While some additional peaks are observed in the extracted ion chromatograms, they can be easily attributed to isobaric species stemming from the hydrolysis of HSA and from the autolysis of trypsin (for example, EQLK peptide has the same mass as ADLAK). The mass spectrum for the peak corresponding to them also shows the two overlapping peptides, one non-labeled and one labeled with two heavy oxygen atoms. No signal corresponding to non-digested analytical construct can be observed. This serves as a proof that the hydrolysis of the peptide is quantitative, and that the labeled

peptide liberated from the analytical construct has the same chromatographic properties as the non-labeled peptide liberated from the protein.

4. Conclusions We have developed a method of synthesis of peptides which can be utilized for quantitative proteomics. Our peptides work in a similar way to analytical constructs: when they are cleaved from resin, they contain a dabsyl moiety, which facilitates their detection and quantification by HPLC; then, when they are subjected to enzyme-catalyzed hydrolysis the dabsyl moiety together with the linker is lost, leaving a peptide labeled with 18O, and therefore able to act as reference standard. The method is fairly universal. Any peptide that contains at least one amino acid which does not require side chain protection can be synthesized using this method. The average error of correlation between NMR and HPLC is below 10%, better than for amino acid analysis. The hydrolysis of the peptide is quantitative. Moreover, LC-MS experiment proves that there is no observable effect of isotopic label on retention times. Thus, we can obtain a desired peptide in a precisely known concentration in a mixture with the protein hydrolysate. A potential advantage of the procedure described herein is a possibility of utilization as a standard a crude peptide obtained by a solid-phase synthesis, without any purification, which reduces significantly time needed for the preparation of isotopically labeled standard.

Conflict of interest statement The authors certify that there is no conflict of interest with any financial organization regarding the material discussed in the manuscript.

Acknowledgment This work has been supported by grant number UMO-2013/11/ N/ST4/01019 from the National Science Centre of Poland.

Appendix A. Supplementary data Supplementary data to this article can be found online at

REFERENCES Fig. 3 – Panel A: mass spectrum of the ADLAK peptide resulting from hydrolysis of the construct. Panel B: mass spectrum of the ADLAK peptide resulting from hydrolysis of the mixture of the construct and HSA. Panel C: extracted ion chromatograms for m/z 517.28 (non-labeled peptide, black) and 521.29 labeled peptide, red). Note the overlap of the red and black peaks at t = 14 min. The broad peak at t = 12 min corresponds to an isobaric peptide, EQLK.

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A synthesis of new, bi-labeled peptides for quantitative proteomics.

Isotopically labeled peptides are often used in proteomics as internal reference allowing quantification of peptides by isotopic dilution method. Alth...
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