mass spectrometry: Towards peptide and protein modification, separation and identification

Journal of Chromatography A, 1479 (2017) 153–160

Contents lists available at ScienceDirect

Journal of Chromatography A journal homepage: www.elsevier.com/locate/chroma

Dual reductive/oxidative electrochemistry/liquid chromatography/mass spectrometry: Towards peptide and protein modification, separation and identification Lars Büter a,b , Lisa M. Frensemeier a , Martin Vogel a , Uwe Karst a,b,∗ a b

Westfälische Wilhelms-Universität Münster, Institut für Anorganische und Analytische Chemie, Corrensstraße 30, 48149 Münster, Germany Westfälische Wilhelms-Universität Münster, NRW Graduate School of Chemistry, Wilhelm-Klemm-Straße 10, 48149 Münster, Germany

a r t i c l e

i n f o

Article history: Received 27 May 2016 Received in revised form 22 November 2016 Accepted 5 December 2016 Available online 6 December 2016 Keywords: Disulfide reduction Quinones Electrochemistry Adduct formation Protein labeling

a b s t r a c t A new hyphenated technique based on on-line dual (oxidative and reductive) electrochemistry coupled to liquid chromatography and high resolution electrospray mass spectrometry is presented. Two liquid streams are combined, with one containing a disulfide, which is reduced to the respective thiol in an electrochemical cell based on a titanium working electrode. The other stream contains phenol, which is electrochemically activated to benzoquinone on a boron-doped diamond working electrode. Upon combination of the two streams, a Michael addition takes places, leading to the covalent coupling of thiol to quinone. In continuous flow, the reaction mixture is transferred into an injection valve and the products are separated by reversed phase liquid chromatography and detected by electrospray-high resolution mass spectrometry. Proof of concept is demonstrated for low molecular mass disulfides and peptides, but further optimization will be required in future work to achieve efficient protein labelling. © 2016 Elsevier B.V. All rights reserved.

1. Introduction The identification of proteins can either be carried out by identifying proteolytic peptides (bottom-up approach) or by analyzing intact proteins (top-town approach). One crucial step in both respective workflows is related to the reduction of disulfide bonds. These covalent dimers of the amino acid cysteine are the most common posttranslational modification of proteins (∼19% of all proteins contain multiple disulfide bonds [1]) and are important for their tertiary structure [2,3]. Within bottom-up experiments, the disulfide bonds have to be reduced quantitatively prior to enzymatic digestion in order to obtain the highest digestion yield possible [4]. In the top-down approach, reduction of disulfide bonds is necessary to achieve sequence information obtained from collision-induced dissociation (CID) experiments [5]. In contrast, methods as electron capture dissociation (ECD) [6] and electron transfer dissociation (ETD) [7] are capable of cleaving both disulfide bonds and the peptide backbone. Usually, the reduction step is performed using chemical reducing reagents such as dithio-

Abbreviations: BDD, boron-doped diamond; FA, formic acid; GSH, glutathione; GSSG, glutathione dimer; H2 Q, hydroquinone; Q, benzoquinone; Q-GSH adduct, benzoquinone-glutathione adduct. ∗ Corresponding author at: University of Münster, Institute of Inorganic and Analytical Chemistry, Corrensstraße 30, 48149 Münster, Germany. E-mail address: [email protected] (U. Karst). http://dx.doi.org/10.1016/j.chroma.2016.12.008 0021-9673/© 2016 Elsevier B.V. All rights reserved.

threitol (DTT) [8] or tris(2-carboxyethyl)phosphine (TCEP) [9,10]. However, since an excess of these reagents is used, the protein sample has to be purified afterwards in order to prevent a modification and an activity loss of the respective applied protease. Due to this time-consuming, multi-step method, alternative, less laborious approaches are required. Electrochemistry as a purely instrumental setup, which allows fast and selective reduction of disulfide bonds by applying negative potentials to the working electrode can serve as an alternative. Kraj et al. have shown a proof of principle by optimizing the parameters applied for the electrochemical reduction of insulin, which contains two intermolecular and one intramolecular disulfide bond [11]. They used a commercially available flow-through thin-layer cell equipped with titanium working and counter electrodes and a Pd/H2 reference electrode. Square-wave pulses were applied to the working electrode in order to achieve a reduction efficiency of 100% for all three disulfide bonds. By varying the parameters of the square-wave pulses, Kraj et al. were able to control the reduction efficiency, which is advantageous for experiments where no quantitative reduction is required. Reduction of somatostatin (14 amino acids, one disulfide bond) allowed the online detection of almost the full series of y- and b-type ions by means of CID experiments. Without reduction, only one y-ion was identified. Finally, partial to complete reduction of ␣lactalbumin (123 amino acids, 4 disulfide bonds) was performed, thus demonstrating that their developed method is also capable

154

L. Büter et al. / J. Chromatogr. A 1479 (2017) 153–160

to assign and localize disulfide bridges in proteins. Nicolardi et al. described an improved Fourier transform ion cyclotron (FTICR)-CID and −ETD sequence coverage of the disulfide containing peptides oxytocin and hepcidin after online electrochemical reduction [12]. In the group of Chen, electrochemical reduction of biomolecules in combination with desorption electrospray ionization (DESI)-MS as detection technique was investigated [13–19]. In a study by Zhang et al. based on the bottom-up approach, electrochemical reduction of enzymatically digested peptides containing disulfide bonds was performed using an amalgam working electrode [15]. They were able to identify disulfide containing enzymatic peptides based on the changes of their relative ion abundance during DESI-MS detection, when the cell was either switched on or off. In addition, several peptides and the protein ␣-lactalbumin were electrochemically reduced and subsequently labelled with selenium reagents, which are reactive towards free thiol functions. Thus, it was possible to distinguish between intra- (+2 tags) and intermolecular (+1 tag) disulfide bonds. Apart from these bottom-up experiments, Zhang et al. [19] investigated the electrochemical reduction of disulfide bonds in top-down proteomics experiments. Here, a significantly enhanced sequence coverage of the two proteins ␤-lactoglobulin A (24 vs. 75 backbone cleavages before and after electrochemical reduction) and lysozyme (5 vs. 66 backbone cleavages) could be obtained. Van Berkel et al. [20] developed a method, which allows to perform disulfide reduction, benzoquinone generation and follow-up reactions within only one experiment. They modified an ESI interface by applying porous flow-through electrodes instead of stainless-steel electrodes at the upstream grounding point and emitter. At the upstream grounding point a disulfide containing peptide was electrochemically reduced and the obtained free thiol groups were subsequently tagged with benzoquinone, which was generated by electrochemically oxidizing hydroquinone at the emitter. Another important aspect in protein analysis is related to the determination of the number of free and disulfide bound cysteines in proteins, which provides additional information for protein characterization [21]. Free cysteines contain a nucleophilic thiol group, which acts as a target for several drug compounds such as platinum(II) anti-cancer drugs [22,23]. Beyond this, thiol groups can be modified with reactive electrophilic intermediates or reactive oxygen and nitrogen species, which can be formed during the metabolism of a xenobiotic or during oxidative stress, respectively [24–27]. These modifications may lead to an activity loss or to toxic side effects. Therefore, analytical methods are required allowing the determination of the number and position of free and disulfide bound cysteine moieties. In the last years, electrochemistry in combination with mass spectrometry has shown to be a suitable tool for the specific modification of cysteine residues in peptides and proteins [20,28–38]. Within this study, a new method was developed, which combines for the first time a dual electrochemical approach (oxidative and reductive) on-line with reversed phase liquid chromatographic separations and electrospray mass spectrometric detection of the formed products. While a proof of principle of this method can be demonstrated, the approach should be suitable for automated labelling approaches after further optimization.

2. Experimental section 2.1. Chemicals Glutathione (GSH), oxytocin, insulin from bovine pancreas and ammonium formate were obtained from Sigma-Aldrich Chemie GmbH (Steinheim, Germany). Formic acid was ordered from Fluka Chemie (Buchs, Switzerland) and acetonitrile from VWR Chemicals

(Darmstadt, Germany). Phenol was purchased from ABCR (Karlsruhe, Germany). All chemicals and solvents were used in the highest quality available. Water was purified before utilization with an Aquatron A4000D system (Barloworld Scientific, Nemours, France). 2.2. Electrochemical reduction of glutathione dimers and oxytocin by means of EC/MS The reduction of oxidized glutathione (GSSG) was performed using an amperometric thin-layer cell (ReactorCell, Antec Leyden, Zoeterwoude, The Netherlands) equipped with a titanium working electrode, a graphite-doped Teflon counter electrode and a Pd/H2 reference electrode. The potential was controlled by a home-made potentiostat. A schematic overview of the applied setup can be found in SI-1, Supporting information. In order to detect GSSG and its reduced form glutathione (GSH), the cell was directly coupled to a high-resolution mass spectrometer (Exactive Orbitrap, Thermo Fisher Scientific, Bremen, Germany) with an electrospray ionization (ESI) source. With a flow-rate of 10 ␮L/min, which was controlled by a syringe pump (model 74900, Cole Parmer, Vernon Hills, USA) a 50 ␮M solution of GSSG in 1% (v/v) formic acid (FA) in purified water containing 10% acetonitrile (ACN) (v/v) was pumped through the electrochemical cell. For reduction, a potential ramp from 0 to −3.0 V versus Pd/H2 with a scan rate of 10 mV/s was applied. Mass spectra were recorded in the positive ion mode and were plotted as three-dimensional mass voltammograms (for detailed MS parameters see SI-1, Supporting information). Electrochemical reduction of oxytocin was performed using a preparative electrochemical thin-layer cell (␮PrepCell, Antec Leyden) equipped with working and counter electrodes made of titanium and a Pd/H2 reference electrode. The potential was controlled by a home-made potentiostat. A solution with a concentration of 10 ␮M (in 1% FA (v/v) and 10% ACN (v/v)) was introduced into the EC cell, where a constant potential of −3.0 V vs. Pd/H2 was applied. The effluent of the cell was continuously analyzed by means of ESI-MS using a time-of-flight (ToF) mass analyzer (micrOTOF, Bruker Daltonics, Bremen, Germany). Mass spectra were recorded in the positive ion mode (for detailed MS parameters see SI-1, Supporting information). 2.3. Electrochemical reduction of oxytocin and insulin by means of EC/LC/MS Oxytocin (50 ␮M) and insulin (10 ␮M) were electrochemically reduced using the previously described preparative cell which was equipped with a titanium working electrode. The reduction potential was set to constant value of −1.2 V vs. Pd/H2 using a homemade potentiostat. An LC separation using a C5 wide pore column ® (Discovery BIO Wide Pore C5, 300 Å, 150 × 2.1 mm, 5 ␮m particle size, Supelco, Steinheim, Germany) and a binary gradient using 0.1% (v/v) FA and ACN was performed prior to MS analysis (for a schematic overview of the used setup see SI-1, Supporting information) in order to achieve a separation of the intact biomolecule and its reduced form. For online EC/LC/ESI-MS analysis, the cell effluent was collected in a 5 ␮L injection loop, which was mounted in a ten-port switching valve. By switching the valve to the injection position, the solution was transferred into the column. For this purpose, an Alexys LC system (Antec Leyden) and a microTOF-MS (Bruker Daltonics) equipped with an ESI source were used. The LC system included two LC 100 pumps, an OR 110 organizer rack with a degasser and a pulse dampener, an AS 100 autosampler, and a Roxy column oven. The flow-rate was set to 300 ␮L/min and the oven temperature was adjusted to 40 ◦ C. The gradient profiles for reduced oxytocin and insulin are shown in Table 1. In order to prevent salts from entering the MS, the effluent of the first 2 min of the LC separation was discarded. Mass spectrometric

L. Büter et al. / J. Chromatogr. A 1479 (2017) 153–160

155

Table 1 Gradient profiles for the LC separations of intact and reduced oxytocin (a) and insulin (b). a) Oxytocin: t [min] %B b) Insulin: t [min] %B

0 10

2 10

10 40

11 40

12 10

15 10

0 25

2 25

10 65

12 65

13 25

16 25

Table 2 Observed m/z and charge states for investigated analytes and their reduction as well as labelling products. Analyte

Detected m/z

Detected charge state

Benzoquinone (Q)

not detected (n.d.) neutral mass: 108.0 Da n.d. neutral mass: 110.0 Da 307.1 613.2 635.1 308.1 330.1 416.1 438.1 1007.4 1009.5 1117.5 1225.5 1147 1170 1223 1277 850 877 904

–

Phenol GSSG

GSH Fig. 1. Instrumental setup for the reduction of disulfide bonds in biomolecules and subsequent online labelling of the generated free cysteines with electrochemically generated reactive intermediates. For electrochemical oxidation, an analytical thin-layer cell equipped with a boron-doped diamond working electrode was used. Electrochemical reduction was performed in different types of thin-layer cells (analytical and preparative EC cell), both equipped with a titanium working electrode. The effluents of the two cells were combined using a T-piece and after passing a reaction coil, the solution was directly infused into an ESI-MS.

detection was performed in the positive ion mode (for detailed MS parameters see SI-1, Supporting information). 2.4. Electrochemical reduction of disulfides and subsequent labeling with electrochemically generated benzoquinone Labeling of reduced biomolecules was performed by combining the effluents of two, one reductive and one oxidative, electrochemical cells (Fig. 1). Analogous to the previously described reduction experiments, GSSG (50 ␮M, 10 ␮L/min) was electrochemically reduced in an analytical thin-layer EC cell (−3.0 V), whereas oxytocin (50 ␮M, 20 ␮L/min) and insulin (5 ␮M, 4 ␮L/min) were reduced using the preparative thin-layer EC cell (−3.0 V). For electrochemical reduction, a home-made potentiostat was used. To the effluent of the reductive cell, a solution of electrochemically oxidized phenol (50 ␮M (GSSG, oxytocin)) and 100 ␮M (insulin) in 50/50 (v/v) aqueous ammonium formate (10 mM, pH 7) and ACN was added via a T-piece. Therefore, phenol was electrochemically oxidized in an analytical thin-layer cell equipped with a borondoped diamond working electrode. The oxidation potential was controlled by a Roxy potentiostat (Antec Leyden). The flow-rate was adjusted to 10 ␮L/min. The applied potential was set to 2.5 V. The mixture containing the reduced and oxidized solutions was allowed to pass a reaction coil. Finally, the mixture was analyzed by means of ESI-MS in the positive ion mode. In case of GSSG, MS detection was performed with an ion-trap mass analyzer (Esquire6000, Bruker Daltonics). For oxytocin and insulin a ToF-MS (micrOTOF, Bruker Daltonics) was used. For detailed MS parameters see SI-1, Supporting information. 3. Results and discussion 3.1. Electrochemical reduction of disulfide containing biomolecules First, electrochemical reduction as well as LC separation of the native and reduced forms of different biomolecules was studied. Therefore, the optimum potential for electrochemical reduction

GSH + Q Oxytocin (native) Oxytocin (reduced) Oxytocin (red.) + Q Oxytocin (red.) + 2Q Insulin (native) A-Chain Insulin A-Chain + Q A-Chain + 2Q B-Chain Insulin B-Chain + Q B-Chain + 2Q

– +2 +1 +1 (sodium adduct) +1 +1 (sodium adduct) +1 +1 (sodium adduct) +1 +1 +1 +1 +5 +2 +2 +2 +4 +4 +4

was determined by applying a potential ramp to glutathione dimers (GSSG). Second, electrochemical reduction of the peptide oxytocin was carried out, and an LC separation was integrated into the setup. Finally, the method was transferred to insulin in order to verify whether electrochemical reduction and LC/MS analysis can also be used for larger biomolecules containing intermolecular disulfide bonds. Glutathione (GSH) is a tripeptide present in almost all cells in living organisms and is particularly important for the detoxification of electrophilic intermediates as well as of reactive oxygen and nitrogen species. Furthermore, it serves as an endogenous nucleophile during phase II metabolism. GSH conjugation can either be spontaneous or catalyzed by glutathione-S-transferases. Since glutathione also serves as antioxidant, the ratio between reduced and oxidized glutathione provides information about the state of possible oxidative stress [39]. GSSG was electrochemically reduced by applying a potential sweep from 0 V to −3.0 V (vs. Pd/H2 ) using a titanium working electrode, which was placed in an analytical thin-layer electrochemical cell (ReactorCell). Titanium was chosen as cathode material analogous to the work of Kraj et al. who reported increased conversion on titanium compared to e.g. boron-doped diamond for disulfide containing molecules [11]. The obtained mass spectra were plotted versus the applied potential. From the obtained mass voltammogram, the optimum reductive potential can be derived. In Fig. 2, the reaction scheme of the electrochemically induced reduction of GSSG (a) as well as the recorded mass voltammogram (b) are shown. The mass voltammogram depicts the recorded mass spectra in dependency of the applied potential. Detected species as well as their m/z and charge states are listed in Table 2. At a m/z of 307.1, a signal for doubly charged GSSG ([GSSG + 2H]2+ ) with decreasing signal intensity at decreasing potentials was detected, indicating a reduction of GSSG. The signal detected at m/z 307.6 corresponds to the 13 C isotope peak of doubly charged GSSG. At

156

L. Büter et al. / J. Chromatogr. A 1479 (2017) 153–160

Fig. 2. a) Schematic reduction pathway of GSSG under formation of GSH. b) Mass voltammogram of the electrochemical reduction of GSSG.

the same time, an increasing signal for the singly charged monomer ([GSH + H]+ , m/z 308.1) with a maximum at −3.0 V was observed. Its 13 C isotope signal was detected at m/z 309.1, also showing increasing signal intensity. The detection of a GSH signal already before applying a potential to the working electrode can be explained by the presence of GSH in the initial solution or by partial reduction of GSSG during the ionization process. Nevertheless, the increasing signal intensity shows that the electrochemical reduction of disulfide bonds is possible. In order to investigate the applicability of the electrochemical reduction also for other disulfide containing molecules, the small hormone and neurotransmitter oxytocin was investigated in the following. Oxytocin contains nine amino acids, two of which are cysteines that form a disulfide bond between position 1 and 6 (see Fig. 3a). This intramolecular disulfide bond fixes the secondary as well as the tertiary structure of the peptide [40]. Electrochemical reduction was performed using a slightly modified system. In contrast to GSSG, a preparative electrochemical thin-layer cell was used. The major differences to the analytical cell are the larger cell volume and electrode surface. Consequently, the residence time of the respective compound is increased thus resulting in higher yields and giving the possibility of reducing even structurally hindered, e.g., knotted or enclosed disulfide bonds. In Fig. 3b, the extracted ion traces of intact (m/z 1007) and reduced (m/z 1009) oxytocin are shown and Table 2 summarizes the observed m/z as well as the corresponding charge states of the molecules. After 1.5 min, a potential of −3.0 V (vs. Pd/H2 ) was switched on for 2.8 min. Immediately, the signal intensity of intact oxytocin decreases, while a signal for the reduced form appears. Based on the extracted ion trace of intact oxytocin, the electrochemical reduction resulted in high yields even if no quantitative

Fig. 3. a) Structure of oxytocin and b) extracted ion traces of intact (m/z 1007) and reduced (m/z 1009) oxytocin. After 1.5 min, the electrochemical reduction (−3.0 V vs. Pd/H2 ) was switched on.

conversion was observed. However, also re-oxidation during electrospray ionization could have taken place. Nevertheless, it could be shown that the reduction of intramolecular disulfide bonds is possible. In an additional experiment, an LC separation was integrated in order to identify potential re-oxidation of reduced oxytocin during the ionization step. Following electrochemical reduction, the different forms of oxytocin were separated online by applying a binary gradient consisting of 0.1% (v/v) formic acid and acetonitrile using a C5 wide pore reversed phase column. The obtained LC/MS chromatograms are shown in Fig. 4a and b. Without electrochemical reduction (Fig. 4a), a signal for oxytocin (m/z 1007) with a retention time of tR = 6.00 min was detected. The additional signal (m/z 1009) showing the same retention time is also caused by intact oxytocin and can be traced back to an isotope peak of oxytocin with a relative abundance of 25.9%. This observation is confirmed by the exact mass (see SI-2, Supporting information, for comparison of the detected and calculated m/z) of the isotope peak and the relative abundances of the two signals. When applying a reductive potential to the working electrode (Fig. 4b), an additional signal at tR = 6.65 min for reduced oxytocin (m/z 1009) was observed in the LC chromatogram. However, no signal for the intact form was obtained at the same time, meaning that no re-oxidation during ESI ionization occurred. In further experiments, the peptide hormone insulin isolated from bovine pancreas, which regulates the carbohydrate and fat metabolism in the organism, was analyzed [41]. The structure of insulin is schematically shown in Fig. 5. It consists of the A- and the B-chain, and contains in total 51 amino acid residues. The two chains are connected via two intermolecular disulfide bonds. Furthermore, one additional intramolecular disulfide bond is present in the A-chain [42].

L. Büter et al. / J. Chromatogr. A 1479 (2017) 153–160

157

Fig. 4. LC/MS chromatograms of a) intact and b) reduced oxytocin. Shown are the extracted ion traces of oxytocin (m/z 1007) and reduced oxytocin (m/z 1009). LC/MS chromatograms of c) intact and d) reduced insulin. Shown are the extracted ion traces of the most abundant charge states of insulin (m/z 1147, charge state +5), the A chain (m/z 1170, charge state +2) and the B chain (m/z 850, charge state +4).

3.2. Combination of electrochemical reduction and oxidation for the subsequent labelling of generated free cysteines

Fig. 5. Scheme of the amino acid sequence of insulin. A- and B-chain are connected via two intermolecular disulfide bonds. Additionally, the A-chain contains one intramolecular disulfide bond.

Analogous to the experiments carried out with oxytocin, insulin was electrochemically reduced in a preparative EC cell (−3.0 V vs. Pd/H2 ) and an online LC separation was performed. In Fig. 4, the observed LC/MS chromatograms of intact (Fig. 4c) and reduced (Fig. 4d) insulin are shown. The m/z of the shown XICs as well as the corresponding charge states of the molecules are summarized in Table 2. The native form of insulin elutes at tR = 4.65 min, as can be seen in the corresponding chromatogram, in which the extracted ion trace of the most abundant charge state (m/z 1147, charge state +5) is plotted. After electrochemical reduction, two additional signals were detected. In both cases, the extracted ion chromatograms of the most abundant charge states show an increasing retention time (A chain: m/z 1170, charge state +2, tR = 5.85 min and B chain: m/z 850, charge state +4, tR = 5.55 min) compared to native insulin. This can be explained by an unfolding of the chains due to breakdown of the tertiary structure. To conclude, by coupling electrochemistry online to LC/MS, it was possible to reduce disulfide bonds of the more complex biomolecule insulin and identify the obtained free thiol groups by means of HPLC/ESI-MS. Therefore, it can be concluded that EC offers the potential for the reduction of disulfide bonds also in larger biomolecules. However, the efficiency of the reduction will have to be further optimized to routinely use the method in proteomics studies.

Since electrochemical reduction of GSSG, oxytocin and insulin has been successfully performed, within the next series of experiments, the possibility of subsequent labelling of the obtained free cysteines with electrochemically generated reactive benzoquinone intermediates was investigated. All analytes and the obtained reduced and labelled peptides are summarized in Table 2, where the observed m/z with their corresponding charge state are presented. Analogous to the previous part of this chapter, the first compound studied was GSSG. For this purpose, GSSG was again electrochemically reduced in an analytical thin-layer cell (titanium working electrode, applied potential: −3.0 V vs. Pd/H2 ) and to the effluent of this cell, an effluent of a second analytical thin-layer cell was added via a T-piece (Fig. 1). In the second cell, phenol was electrochemically oxidized at 2.5 V (vs. Pd/H2 ) using a boron-doped diamond (BDD) working electrode in order to generate reactive benzoquinone species. The oxidation pathway of phenol and the reaction between its product benzoquinone and GSH are schematically shown in Fig. 6. In a first step, the transfer of two electrons and one proton leads to the formation of an intermediary cation, which directly undergoes hydrolysis resulting in the formation of hydroquinone (H2 Q). Since hydroquinone has a lower oxidation potential than phenol, dehydrogenation can immediately take place, leading to the generation of reactive benzoquinone (Q) intermediates. Fig. 6 solely illustrates the formation of para-hydroquinone and parabenzoquinone, but in principle also ortho-hydroquinone formation as well its corresponding quinoid structure is possible due to the positive mesomeric effect of the hydroxyl group of phenol. Due to its electrophilic character, benzoquinone is capable of specifically reacting in a Michael-type addition with the nucleophilic thiol group of cysteine in GSH. Rearomatization then leads to the formation of a stable product (Q-GSH adduct). Labelling of cysteines by means of electrochemically generated reactive species,

158

L. Büter et al. / J. Chromatogr. A 1479 (2017) 153–160

Fig. 6. a) Pathway of the electrochemical oxidation of phenol. The formed hydroquinone (H2 Q) can directly undergo further oxidation yielding a reactive benzoquinone (Q). b) Reaction scheme of the Michael-type addition of the free thiol group of glutathione (GSH) with Q.

such as benzoquinone, has the advantage of a simple instrumental setup without the need of purification steps or the presence of interferences during MS analysis resulting from excessive labelling reagents used in conventional approaches. For MS analysis of the subsequent labelling of electrochemically reduced GSSG with generated benzoquinone, three different experiments were carried out. The recorded mass spectra of each experiment are shown in Fig. 7. First, a mass spectrum of GSSG and phenol was recorded, where both oxidation and reduction were switched off (Fig. 7a). Two signals for GSSG (protonated form [GSSG + H]+ : 613 and sodium adduct [GSSG + Na]+ : 635) were detected. However, no signal for phenol was detected since it is not ionized during electrospray ionization in the positive ion mode. When applying only a reductive potential, additional signals for GSH arise (Fig. 7b). Again, protonated GSH ([GSH + H]+ : 308) as well as the respective sodium adduct [GSH + Na]+ : 330] were found. Afterwards, the second cell was switched on and phenol was oxidized. In the corresponding mass spectrum in Fig. 7c, next to the signals for GSSG and GSH, signals for an adduct of GSH with benzoquinone ([Q + GSH + H]+ : 416 and [Q + GSH + Na]+ : 438) were identified. Thus, by reducing the intermolecular disulfide bond of GSSG in a first cell and oxidizing phenol in a second cell, it was possible to obtain benzoquinone tagged with GSH in an on-line flow through system. In the next experiment, oxytocin was electrochemically reduced and allowed to react with electrochemically generated benzoquinone. Analogous to the reduction experiments of oxytocin presented in the first part, a preparative EC cell was used. Oxidation of phenol was again performed in an analytical cell. After reduction of the intramolecular disulfide bond of oxytocin, the resulting free cysteines can subsequently be labelled with benzoquinone. In Fig. 8, the extracted ion traces of intact (m/z 1007), reduced (m/z 1009), singly (m/z 1117) and doubly tagged oxytocin (m/z 1225) are shown. After switching on both potentials, the signal of the intact form decreases, while the signal intensities for the reduced and labelled forms increase. The presence of a signal for reduced oxytocin during the entire experiment can again be explained by an overlap of the main isotope signal of reduced oxytocin with an isotope peak of the intact form. Furthermore, the slopes of the traces of reduced, singly and doubly tagged oxytocin indicate that tagging of the two thiol groups takes place successively. While the traces of reduced and singly tagged oxytocin directly increase when the potentials were switched on, the trace of the doubly tagged form increases more steeply after around 2 min.

Fig. 7. Mass spectra of a) phenol and dimerized glutathione (GSSG) without applied potential, b) only reduction and c) reduction and oxidation. After reduction, signals for reduced glutathione (GSH) were observed (b). Electrochemical generation of benzoquinone (Q) allowed the subsequent labelling of GSH (c).

Fig. 8. Extracted ion traces of native (m/z 1007), reduced (m/z 1009), singly (m/z 1117) and doubly (m/z 1225) labelled oxytocin. After 1.9 min, electrochemical oxidation and reduction potentials were switched on.

This shows that reduction and labelling of intramolecular disulfide bonds was successfully carried out using oxytocin as model compound. In order to investigate whether the method is also able to reduce and tag larger biomolecules, insulin was studied. The corresponding reduction and labelling pathway is shown in Fig. 9a. Insulin contains three disulfides, resulting in the formation of in total six free cysteines after reduction, which may be modified with benzoquinone. Again, insulin was reduced using a preparative

L. Büter et al. / J. Chromatogr. A 1479 (2017) 153–160

159

Fig. 9. a) Schematic overview on the reduction of insulin and subsequent tagging of the generated free thiol groups with electrochemically formed benzoquinone. Extracted ion traces of b) insulin (m/z 1147, intensity divided by 10), A-chain (m/z 1169), singly (m/z 1223) and doubly (m/z 1277) labelled A-chain. c) Ion traces of insulin (m/z 1147, intensity divided by 10), B-chain (m/z 850), singly (m/z 877) and doubly (m/z 904) labelled B-chain. *: Two cysteine residues are labelled with Q.

cell, while phenol was oxidized in an analytical cell. The extracted ion traces obtained from the experiment are shown in Fig. 9b, c. On the left, the traces for intact insulin (m/z 1147, charge state +5), the A-chain (m/z 1169, charge state +2) as well as for singly (m/z 1223, charge state +2) and doubly (m/z 1277, charge state +2) labelled A-chain are shown. When the potentials are applied to the working electrodes, the intensity of insulin decreases. At the same time, signals for the free, singly and doubly tagged A-chain appear with increasing intensities. Nevertheless, no three and four times tagged A chain could be detected. This leads to the assumption that the intramolecular disulfide bond of the A chain is not reduced. This assumption needs to be confirmed by further experiments like enzymatic digestion or LC/MS/MS, since it would also be possible that a mixture of doubly tagged A-chain at different positions was formed. In addition, as shown in Fig. 9c, increasing signal intensities for the free (m/z 850, charge state +4), singly (m/z 877, charge state +4) and doubly (m/z 904, charge state +4) labelled B-chain were obtained after switching on the potentials. Hence, partial and complete labelling of the electrochemically generated free cysteine moieties in the B-chain was achieved. To conclude, in the second part of the work, a method was developed, which allows the direct tagging of electrochemically generated free cysteines from peptides and proteins by electrochemically formed reactive intermediates. Labelling of reduced cysteines was successful not only for smaller peptides such as glutathione, but also for more complex biomolecules like insulin. While the proof of principle has been demonstrated clearly, further work requires the optimization of the reduction and oxidation efficiencies to allow automated protein tagging in the future. 4. Conclusion Within this study, the first on-line hyphenated technique consisting of two electrochemical cells (one operated under oxidative, one under reductive conditions), a reversed phase liquid chromatographic separation and high resolution electrospray mass spectrometry has been developed. It was demonstrated that reduction of a disulfide to a thiol, simultaneous oxidation of phenol to

benzoquinone, the subsequent reaction of thiol and benzoquinone as well as their LC separation and ESI-MS detection is possible using a continuous flow reaction system, which discharges into the injection valve of an LC/MS system. Peptide and protein labelling were demonstrated using this approach, but significant further optimization, particularly regarding the efficiency of the electrochemical reactions, is required, before an application in the field of routine proteomics analysis can be considered. Acknowledgement The authors would like to thank the NRW Graduate School of Chemistry for financial support in form of a Ph.D. scholarship for L.B. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.chroma.2016.12. 008. References [1] J. Alegre-Cebollada, P. Kosuri, J.A. Rivas-Pardo, J.M. Fernandez, Direct observation of disulfide isomerization in a single protein, Nat. Chem. 3 (2011) 882–887. [2] J.M. Thornton, Disulphide bridges in globular proteins, J. Mol. Biol. 151 (1981) 261–287. [3] W.J. Wedemeyer, E. Welker, M. Narayan, H.A. Scheraga, Disulfide bonds and protein folding, Biochemistry 39 (2000) 4207–4216. [4] Y.Y. Zhang, B.R. Fonslow, B. Shan, M.C. Baek, J.R. Yates, Protein analysis by shotgun/bottom-up proteomics, Chem. Rev. 113 (2013) 2343–2394. [5] J.A. Loo, C.G. Edmonds, R.D. Smith, Primary sequence information from intact proteins by electrospray ionization tandem mass spectrometry, Science 248 (1990) 201–204. [6] R.A. Zubarev, N.A. Kruger, E.K. Fridriksson, M.A. Lewis, D.M. Horn, B.K. Carpenter, F.W. McLafferty, Electron capture dissociation of gaseous multiply-charged proteins is favoured at disulfide bonds and other sides of high hydrogen atom affinity, J. Am. Chem. Soc. 121 (1999) 2857–2862. [7] H.P. Gunawardena, L. Gorenstein, D.E. Erickson, Y. Xia, S.A. McLuckey, Electron transfer dissociation of multiply protonated and fixed charge disulfide linked polypeptides, Int. J. Mass Spectrom. 265 (2007) 130–138.

160

L. Büter et al. / J. Chromatogr. A 1479 (2017) 153–160

[8] W.W. Cleland, Dithiotreitol, a new protective agent for SH groups, Biochem. US 3 (1964) 480–482. [9] W.R. Gray, Disulfide structures of highly bridged peptides A new strategy for analysis, Protein Sci. 2 (1993) 1732–1748. [10] T.Y. Yen, H. Yan, B.A. Macher, Characterizing closely spaced, complex disulfide bond patterns in peptides and proteins by liquid chromatography/electrospray ionization tandem mass spectrometry, J. Mass Spectrom. 37 (2002) 15–30. [11] A. Kraj, H.J. Brouwer, N. Reinhoud, J.P. Chervet, A novel electrochemical method for effcient reduction of disulfide bonds in peptides and proteins prior to MS detection, Anal. Bioanal. Chem. 405 (2013) 9311–9320. [12] S. Nicolardi, M. Giera, P. Kooijman, A. Kraj, J.P. Chervet, A.M. Deelder, Y.E.M. van der Burgt, On-line electrochemical reduction of disulfide bonds Improved FTICR-CID and −ETD coverage of oxytocin and hepcidin, J. Am. Soc. Mass Spectr. 24 (2013) 1980–1987. [13] Y. Zhang, Z.Q. Yuan, H.D. Dewald, H. Chen, Coupling of liquid chromatography with mass spectrometry by desorption electrospray ionization (DESI), Chem. Commun. 47 (2011) 4171–4173. [14] J.W. Li, H.D. Dewald, H. Chen, Online coupling of electrochemical reactions with liquid sample desorption electrospray ionization-mass spectrometry, Anal. Chem. 81 (2009) 9716–9722. [15] Y. Zhang, H.D. Dewald, H. Chen, Online mass spectrometric analysis of proteins/peptides following electrolytic cleavage of disulfide bonds, J. Proteome Res. 10 (2011) 1293–1304. [16] M. Lu, C. Wolff, W.D. Cui, H. Chen, Investigation of some biologically relevant redox reactions using electrochemical mass spectrometry interfaced by desorption electrospray ionization, Anal. Bioanal. Chem. 403 (2012) 355–365. [17] Q. Zheng, H. Zhang, H. Chen, Integration of online digestion and electrolytic reduction with mass spectrometry for rapid disulfide-containing protein structural analysis, Int. J. Mass Spectrom. 353 (2013) 84–92. [18] Q. Zheng, H. Zhang, L. Tong, S. Wu, H. Chen, Cross-linking electrochemical mass spectrometry for probing protein three-dimensional structures, Anal. Chem. 86 (2014) 8983–8991. [19] Y. Zhang, W.D. Cui, H. Zhang, H.D. Dewald, H. Chen, Electrochemistry-assisted top-down characterization of disulfide-containing proteins, Anal. Chem. 84 (2012) 3838–3842. [20] G.J. Van Berkel, V. Kertesz, Expanded electrochemical capabilities of the electrospray ion source using porous flow-through electrodes as the upstream ground and emitter high-voltage contact, Anal. Chem. 77 (2005) 8041–8049. [21] J.J. Gorman, T.P. Wallis, J.J. Pitt, Protein disulfide determination by mass spectrometry, Mass Spectrom. Rev. 21 (2002) 183–216. [22] T. Peleg-Shulman, Y. Najajreh, D. Gibson, Interaction of cisplatin and transplatin with proteins comparison of binding kinetics, binding sites and reactivity of the Pt-protein adducts of cisplatin and transplatin towards biological nucleophiles, J. Inorg. Biochem. 91 (2002) 306–311. [23] C. Brauckmann, H. Faber, C. Lanvers-Kaminsky, M. Sperling, U. Karst, Influence of cimetidine and ist metabolites on cisplatin – investigation of adduct formation by means of electrochemistry/liquid chromatography/electrospray mass spectrometry, J. Chromatogr. A 1279 (2013) 49–57. [24] F.T. van den Brink, L. Büter, M. Odijk, W. Olthuis, U. Karst, A. van den Berg, Mass spectrometric detection of short-lived drug metabolites generated in an electrochemical microfluidic chip, Anal. Chem. 87 (2015) 1527–1535. [25] W. Lohmann, H. Hayen, U. Karst, Covalent protein modification by reactive drug metabolites using online electrochemistry/liquid chromatography/mass spectrometry, Anal. Chem. 80 (2008) 9714–9719.

[26] K.G. Madsen, J. Olsen, C. Skonberg, S.H. Hansen, U. Jurva, Development and evaluation of an electrochemical method for studying reactive phase-I-metabolites correlation to in vitro drug metabolism, Chem. Res. Toxicol. 20 (2007) 821–831. [27] H.P. Permentier, A.P. Bruins, R. Bischoff, Electrochemistry-mass spectrometry in drug metabolism and protein research, Mini Rev. Med. Chem. 8 (2008) 46–56. [28] L. Büter, M. Vogel, U. Karst, Adduct formation of electrochemically generated reactive intermediates with biomolecules, TrAC Trends Anal. Chem. 70 (2015) 74–91. [29] T.C. Rohner, J.S. Rossier, H.H. Girault, On-line electrochemical tagging of cysteines in proteines during nanospray, Electrochem. Commun. 4 (2002) 695–700. [30] C. Roussel, T.C. Rohner, H. Jensen, H.H. Girault, Mechanistic aspects of on-line electrochemical tagging of free L-cysteine residues during electrospray ionization for mass spectrometry in protein analysis, ChemPhysChem 4 (2003) 200–206. [31] T.C. Rohner, J. Josserand, H. Jensen, H.H. Girault, Numerical investigations of an electrochemically induced tagging in a nanospray for protein analysis, Anal. Chem. 75 (2003) 2065–2074. [32] C. Roussel, L. Dayon, H. Jensen, H.H. Girault, On-line cysteine modification for protein analysis new probes for electrochemical tagging nanospray mass spectrometry, J. Electroanal. Chem. 570 (2004) 187–199. [33] C. Roussel, L. Dayon, N. Lion, T.C. Rohner, J. Josserand, J.S. Rossier, H. Jensen, H.H. Girault, Generation of mass tags by the inherent electrochemistry of electrospray for protein mass spectrometry, J. Am. Soc. Mass Spectr. 15 (2004) 1767–1779. [34] L. Dayon, C. Roussel, H.H. Girault, On-line electrochemical tagging of free cysteines in peptides during nanospray ionization mass spectrometry an overview, Chimia 58 (2004) 204–207. [35] L. Dayon, C. Roussel, M. Prudent, N. Lion, H.H. Girault, On-line counting of cysteines during electrospray ionization by electrogenerated tags and their application to protein identification, Electrophoresis 26 (2005) 238–247. [36] L. Dayon, C. Roussel, H.H. Girault, Probing cysteine reactivity in proteins by mass spectrometric EC-tagging, J. Proteome Res. 5 (2006) 793–800. [37] M. Abonnenc, L. Dayon, B. Perruche, N. Lion, H.H. Girault, Electrospray micromixer chip for on-line derivatization and kinetic studies, Anal. Chem. 80 (2008) 3372–3378. [38] G.J. Van Berkel, V. Kertesz, Electrochemically initiated tagging of thiols using an electrospray ionization based liquid microjunction surface sampling probe two-electrode cell, Rapid Commun. Mass Spectrom. 23 (2009) 1380–1386. [39] A. Pastore, G. Federici, E. Bertini, F. Piemonte, Analysis of glutathione implication in redox and detoxification, Clin. Chim. Acta 333 (2003) 19–39. [40] A. Argiolas, G.L. Gessa, Central functions of oxytocin, Neurosci. Biobehav. Rev. 15 (1991) 217–231. [41] P. Sonksen, J. Sonksen, Insulin understanding its action in health and disease, Br. J. Anaesth. 85 (2000) 69–79. [42] A.P. Ryle, F. Sanger, L.F. Smith, R. Kitai, The disulphide bonds of insulin, Biochem. J. 60 (1955) 541–556.