Structural Interrogation of Electrosprayed Peptide Ions by Gas-Phase

Article pubs.acs.org/ac

Structural Interrogation of Electrosprayed Peptide Ions by Gas-Phase H/D Exchange and Electron Capture Dissociation Mass Spectrometry Jingxi Pan,† Brittany L. Heath,‡ Rebecca A. Jockusch,*,‡ and Lars Konermann*,† †

Department of Chemistry, The University of Western Ontario, London, Ontario, N6A 5B7, Canada Department of Chemistry, University of Toronto, Toronto, Ontario M5S 3H6, Canada

‡

ABSTRACT: The structural characterization of gaseous biomolecular ions remains a challenging task. Here, we employ a combination of gas-phase hydrogen−deuterium exchange (HDX) and electron capture dissociation (ECD) mass spectrometry for gaining insights into the properties of two electrosprayed peptides: RA9K and RG9K. Mass analysis of ECD fragments provides spatially resolved labeling information. ND3-mediated HDX at peptide termini and amino acid side chains goes to completion within 1 s. Backbone amide labeling occurs more slowly, and proceeds in a structurally sensitive fashion. HDX is more extensive for RG9K than for RA9K, suggesting a more “open” conformation for the former. Residues 7−10 in RA9K are strongly protected, which indicates the presence of stable backbone hydrogen bonds at these sites. Our findings are consistent with the results of previous ion mobility measurements and computational investigations. Overall, it appears that the combination of gas-phase HDX and ECD represents a viable approach for uncovering structural features of biomolecular ions in the gas phase.

E

digestion step into the experimental workflow. Deuteration levels of individual protein regions can then be determined from the mass shifts of proteolytic peptides.18,21−24 In parallel to these solution-phase investigations,18−24 attempts have been made to employ gas-phase HDX with the goal of probing structural features of biomolecules in vacuo.2,25−32 Various labeling agents, such as ND3, D2O, or deuterated alcohols, have been employed for these experiments. The mechanisms of gas-phase HDX are dependent on the nature of the deuteration agent and on the type of labeling site.33−35 Analogous to solution-phase studies, the interpretation of gas-phase HDX data is usually guided by the tenet that “open” conformers undergo more-extensive exchange than compact structures.36 Support for this view comes from experiments on shaperesolved protein ions.37,38 However, the correlation between gas-phase conformation and HDX behavior may not always be as straightforward, especially when using D2O as a labeling agent.5,39−42 Nonetheless, it is undisputed that HDX approaches have tremendous potential in aiding the elucidation of biomolecular structures in the gas phase. A key problem that has hampered the development of gasphase HDX as a structural tool is the difficulty in determining the spatial distribution of deuterium atoms in gaseous biomolecular ions after labeling.43 For very small systems (e.g., single amino acids), this information is obtainable via IR methods,44 but this approach quickly becomes unfeasible as the analyte size increases. Hence, previous gas-phase HDX studies were largely restricted to analyses of overall exchange levels and rates.

lectrospray ionization (ESI) allows the transfer of intact biological molecules from solution into the gas phase, thereby making these species amenable to analysis by mass spectrometry (MS).1 In addition, there is considerable interest in exploring the behavior of biomolecular ions in a solvent-free environment. Comparative studies of solution-phase and gas-phase structures can reveal “intrinsic” properties of biomolecular analytes. Also, investigations of this type can provide fundamental insights into the nature of biomolecule-solvent interactions.2,3 Techniques that have been applied for structural investigations on gaseous biomolecules include computational modeling,4 ion mobility spectrometry (IMS),5−9 proton transfer measurements,10 infrared/ultraviolet (IR/UV) double-resonance spectroscopy,11 IR multiple photon dissociation,12 and other fragmentation approaches,13−15 as well as fluorescence techniques.16,17 Unfortunately, obtaining detailed structural information for biomolecules in the gas phase remains a challenging task, particularly for large biopolymers. This is in stark contrast to the condensed phase, where atomically resolved data are available from X-ray crystallography and NMR spectroscopy. Hydrogen−deuterium exchange (HDX) methods represent a well-established tool for probing the structure and dynamics of proteins in solution.18−20 Hydrogen atoms in O−H, N−H, and S−H bonds can be replaced with deuterium from a D2Ocontaining solvent. Most studies focus on the HDX behavior of backbone amide hydrogens. These sites are strongly affected by the presence of secondary structure elements. Backbone −CO−NH− groups located in disordered and highly dynamic regions undergo rapid exchange. In contrast, HDX is much slower in segments that are tightly folded and hydrogenbonded. The readout of solution-phase HDX experiments can be performed by NMR spectroscopy19 or by MS. Spatially resolved data are obtainable by HDX/MS when incorporating a © 2011 American Chemical Society

Received: October 14, 2011 Accepted: November 30, 2011 Published: November 30, 2011 373

dx.doi.org/10.1021/ac202730d | Anal. Chem. 2012, 84, 373−378

Analytical Chemistry

Article

(1.0 mm o.d., 0.78 mm i.d.), which had been pulled to a 5−10 μm opening at one end using a micropipet puller (Model P97, Sutter Instruments, Novato, CA). A platinum wire inserted into the glass tip was held at ground, while a voltage of −800 V was applied to the entrance of the mass spectrometer to generate the electrospray. Gas-Phase HDX and ECD. Experiments were performed using a modified 7 T Apex Qe Fourier transform ion cyclotron resonance (FTICR) mass spectrometer (Bruker Daltonics, Billerica, MA) equipped with an Apollo II source. The Apex Qe features a quadrupole mass filter and a hexapole collision cell for mass selection and storage of ions prior to their injection into the ICR cell. Gaseous peptide ions of the desired m/z were mass selected with the quadrupole and then accumulated in the hexapole collision cell. All instrument parameters, excluding quadrupole mass selection, were kept the same for experiments on both peptides. The collision voltage, which helps to control the kinetic energy of the ions, was kept low (0 V), to achieve maximum exchange. Under normal operating conditions, argon gas is used for ion trapping and CID within the hexapole cell. For gas-phase HDX experiments, the argon gas flow was stopped and a second port was used to introduce deuterated ammonia gas (ND3) into the hexapole as the HDX agent. The background pressure in the hexapole collision cell (with no added gas flow) was 2.5 × 10−6 mbar. For HDX experiments, a needle valve was used to adjust the ND3 flow rate to obtain a steady pressure reading of 1.4 × 10−5 mbar, as measured by an ionization gauge mounted externally to the collision cell. The actual pressure of ND3 in the collision cell is significantly higher than this value (estimated to be ∼10−3 mbar), because a conductance limit is located between the collision cell and the ion gauge. The ions were accumulated in the ND3-filled collision cell for times ranging from 0.5 s to 6 s, whereupon they were passed into the ICR cell for ECD and mass analysis. Attempts to use longer HDX times were hampered by fluctuations in the measured deuteration levels. The origin of these fluctuations could not be determined with certainty. Importantly, for accumulation times up to 6 s, the data were found to be highly reproducible (see below). A future extension of the labeling time window might be possible by accumulating protein ions in an external radiofrequency trap, prior to injection into the HDX cell.66 Electrons for ECD were generated by passing a current of 1.7 A through a hollow cathode heater. The electrons were emitted with a pulse duration of 0.2 s. A 2.5 V electron beam bias and a 35 V lens potential were used to guide and focus the electrons into the ICR cell. Each ECD spectrum represents the average of twenty 0.1180 s transients with 256k data points. Data Analysis. The average number of deuterium atoms acquired by intact, doubly charged peptide ions (Figure 1) was determined as 2 × (RD − R0), where RD and R0 represent centroid m/z values after and before HDX, respectively. RD values were determined from the measured mass spectra using HX-Express,67 whereas R0 values were obtained from ProteinProspector (http://prospector.ucsf.edu). In an analogous fashion, the average number of deuterium atoms incorporated into singly charged c ions was calculated as (RD − R0). The deuteration level D for individual backbone amide groups is defined as D = 1 for sites that are completely deuterated (no protection). Conversely, D = 0 corresponds to a total lack of deuteration (full protection). Fractional D values represent partial protection. Similar to earlier experiments,45 c ion signals in the ECD spectra were found to be very intense, whereas z•

Gas-phase fragmentation techniques represent an obvious strategy for spatially resolved HDX studies on desolvated biomolecular ions. Surprisingly, this avenue remains largely unexplored for experiments that involve isotope exchange in the gas phase.27 In contrast, the use of gas-phase dissociation techniques following HDX in solution has become a widely accepted strategy.45−50 Collision-induced dissociation (CID) is unsuitable for experiments of this type, because it tends to induce H/D scrambling.27,51−55 In addition, there is some evidence for the occurrence of intermolecular isotope migration during CID.56 Electron capture dissociation (ECD)57 and electron transfer dissociation (ETD),58 on the other hand, allow the fragmentation of polypeptide ions in a scramblingfree fashion. The viability of using ECD and ETD for spatially resolved HDX measurements has been empirically demonstrated in numerous studies on peptides48−50,59,60 and proteins.45,46,61 The exact mechanism of electron-induced N−Cα backbone cleavage, however, is still under investigation.57,58,62−64 The current work employs a combination of gas-phase HDX and ECD-MS, with the objective of probing the structural features of biomolecules in the gas phase. These experiments involve the exposure of electrosprayed analyte ions to a deuterating agent, followed by gas-phase fragmentation. We focus on the behavior of gaseous [M+2H]2+ ions of RA9K and RG9K (Figure 1). The choice of these peptides is based on the

Figure 1. Structure of doubly protonated peptides used in this study.65 Exchangeable hydrogens are highlighted in bold/red. X = CH3 for RA9K; X = H for RG9K.

fact that their fragmentation behavior65 and their gas-phase structures4 have previously been studied in detail. IMS and molecular dynamics simulations suggest that residues 5−10 in RA9K form an α-helix, whereas the first four residues appear to be disordered. In the case of RG9K, the entire backbone adopts a more open conformation without any discernible secondary structure.4 Our experiments reveal significant differences in the gas-phase HDX characteristics of the two peptides, consistent with their structural properties. Overall, this work demonstrates that HDX/ECD-MS represents a viable approach for structural investigations of biomolecular analytes in the gas phase.

■

EXPERIMENTAL SECTION Materials and Sample Preparation. RA9K and RG9K peptides were obtained as lyophilized powder from GenScript USA, Inc. (Piscataway, NJ). The peptides were transferred into the gas phase using nanoESI. As in previous investigations on the gas-phase behavior of these species,4 the solvent used consisted of 50:50 methanol/water (v/v) and 0.1% acetic acid. The peptide concentration was 5 μM. NanoESI was performed in positive ion mode using borosilicate glass capillaries 374

dx.doi.org/10.1021/ac202730d | Anal. Chem. 2012, 84, 373−378

Analytical Chemistry

Article

considered, because of S/N limitations. Representative ECD fragment ion signals are displayed in Figure 3. The number of exchangeable hydrogens in each ECD fragment can be determined from the known c ion composition (Figure 4A). In this way, a reference profile is obtained against which experimental data can be compared. Mass analysis of ECD fragments after gas-phase HDX yields the data shown in Figures 4B and 4C. To simplify the following discussion, only two HDX time points (1 s and 6 s) will be considered. As expected, the deuteration levels of all ECD fragments increase with labeling time. This increase is less pronounced for RA9K than for RG9K (Figures 4B and 4C), consistent with the behavior observed at the intact peptide level (Figure 2). The experimental profiles of Figure 4 allow the determination of spatial peptide deuteration patterns with almost single-residue resolution. We will first explore if short-term HDX primarily affects side chains and termini, or whether backbone amides are labeled first. Focusing on the t = 1 s data for RG9K (Figure 4C), it is seen that c1 carries seven deuterium atoms, consistent with complete labeling of the N-terminal amine group (2 sites) and the protonated R side chain (five sites). Figure 4A also shows that complete labeling of the K side chain and the −COOH terminus should provide a deuteration difference of 3 between c10 and intact RG9K. This is consistent with the measured difference of 3.1 ± 0.2 (Figure 4C). Analogous considerations apply for RA9K (Figure 4B). Thus, it can be concluded that short-term (t = 1 s) ND3 exposure of RA9K and RG9K induces complete deuteration at the R/K side chains and at the H2N− and −COOH backbone termini. In contrast, backbone deuteration is quite low for t = 1 s, with average deuteration levels on the order of 0.1 for each −CO− NH− site. This implies that the backbone amide HDX rates are considerably lower than those of side chains and termini. Backbone Deuteration Behavior. To discuss the backbone HDX behavior in more detail, it is beneficial to focus on the t = 6 s results, because the overall level of deuteration is higher under these conditions. From the fragment ion data of Figures 4B and 4C, it is possible to determine the deuteration level D of individual −CO−NH− sites (see the Experimental Section for the definition of D). The deuteration level of several backbone sites in RA9K is ∼0.3. Strikingly, however, residues 7−10 are completely protected (D ≈ 0, Figure 5A). We ascribe this protection to the presence of stable hydrogen bonds for these residues, most likely due to helical secondary structure. This interpretation is consistent with the findings of Albrieux et al.,4 although the authors of that work suggested the participation of two additional residues (5 and 6) in an α-helix formation. Perhaps the relatively low protection seen in Figure 5A for residues 5 and 6 can be attributed to helix fraying, i.e., dynamic transitions between a helical conformation and more disordered orientations for these two residues. Fraying phenomena of this type are a common occurrence for polypeptide helices in solution.69,70 A very different backbone deuteration pattern is seen for RG9K (Figure 5B), with D values that are generally higher than for RA9K. The C-terminal K amide exhibits the lowest level of protection, with a D value close to unity. This general behavior is consistent with the known fact that RG9K adopts a moreopen structure.4 Interestingly, residue 4 in RG9K exhibits strong protection, which is a phenomenon that might reflect an involvement of this residue in charge solvation.4 Overall, our data reveal that the differences in global exchange profiles of the two peptides are caused by dissimilarities in their backbone HDX behavior.

ion signals exhibited much lower signal-to-noise (S/N) ratios. For this reason, the data analysis of this work focuses exclusively on the c ion behavior. It has been reported that ECD can also result in the formation of low-abundance Nterminal fragments that have lost a hydrogen (c•), and Cterminal fragments that have gained a hydrogen (z).68 Under the conditions of the current work, however, no such c• or z ions were observed, as confirmed in control experiments on nondeuterated peptides. To determine the deuteration level of individual amide linkages, it has to be considered that the fragment ion cn encompasses the backbone amide NH sites of residues 2 to n + 1. Furthermore, only the first and the last residue in the peptides studied here contain exchangeable side chain sites. Hence, the backbone amide deuteration level D of residue n+1 can be determined as (average number of deuterium atoms in cn) − (average number of deuterium atoms in cn−1).59 Residue 1 does not have a backbone amide but carries the free Nterminus. In other words, the first peptide backbone −NH− is part of residue 2. All measurements were conducted in triplicate; error bars represent standard deviations.

■

RESULTS AND DISCUSSION Intact Peptide HDX. Gaseous [M+2H]2+ ions of RA9K and RG9K (Figure 1) were generated by nanoESI. Both species possess 21 exchangeable hydrogens (10 backbone amides, 6 side-chain sites, 3 on the termini, and 2 charge carriers on the side chains of R and K).65 ND3 was chosen as gas-phase labeling agent, because it has been shown to provide more-extensive HDX and higher deuteration rates than other deutero-compounds.33 Mass-selected peptide ions were trapped for times ranging from 0.5 s to 6 s in the presence of ∼10−3 mbar ND3. The ions were then passed into the ICR cell of a Fourier transform mass spectrometer for ECD and subsequent mass analysis. HDX measurements conducted at the intact peptide level reveal different isotope exchange properties for the two peptides (Figure 2). At the earliest time point studied, both

Figure 2. Global HDX kinetics of doubly protonated peptides RA9K and RG9K after exposure to ND3 in the gas phase.

species have acquired 11 deuterium atoms. In the case of RA9K, the average deuteration level only increases to ∼12 within the experimental time window. In contrast, RG9K undergoes more extensive HDX and incorporates a total of ∼14 deuteriums at t = 6 s (Figure 2). The lower protection seen for RG9K is consistent with its more-open structure.4 Spatial H/D DistributionTermini and Side Chains. ECD of the [M+2H]2+ ions produces spectra that are dominated by singly charged c fragments, providing virtually complete sequence coverage. Only the c1 ion of RA9K could not be 375

dx.doi.org/10.1021/ac202730d | Anal. Chem. 2012, 84, 373−378

Analytical Chemistry

Article

Figure 3. Representative ECD fragment ion MS signals of (A−F) RA9K and (G−L) RG9K. For both peptides, the data shown represent c6+ and c9+ ions, as indicated along the top. The first row of spectra depicts data obtained for undeuterated peptide ions. Data obtained after gas-phase HDX with ND3 at labeling times of 1 and 6 s, are shown in the second and third row, respectively.

Figure 5. Amide deuteration level (D) of individual residues after 6 s of gas-phase HDX for (A) RA9K and (B) RG9K. These data were obtained from Figure 4 by considering the difference of consecutive c ion deuteration values, as outlined in the Experimental Section.

Backbone isotope exchange is slow and proceeds in a structurally sensitive fashion, whereas rapid HDX at side chains and termini does not yield structural insights under the conditions used here.

■

Figure 4. (A) Total number of exchangeable sites in ECD fragments of RA9K and RG9K. The last data point refers to intact [M+2H]2+. (B) Average number of deuterium atoms contained in ECD fragments of RA9K after 1 s (open squares) and 6 s (closed circles) of gas-phase HDX. (C) Same is in panel B, but for RG9K. Note that the ordinate in panel (A) is scaled differently from panels (B) and (C).

CONCLUSIONS Our work demonstrates that ECD is capable of providing spatially resolved information on the deuteration pattern of gaseous biomolecular ions following gas-phase HDX. This 376

dx.doi.org/10.1021/ac202730d | Anal. Chem. 2012, 84, 373−378

Analytical Chemistry

Article

(14) Kitova, E. N.; Wang, W.; Bundle, D. R.; Klassen, J. S. J. Am. Chem. Soc. 2002, 124, 13980−13981. (15) Enyenihi, A. A.; Yang, H.; Ytterberg, A. J.; Lyutvinskiy, Y.; Zubarev, A. R. J. Am. Soc. Mass Spectrom. 2011, 22, 1763−1770. (16) Iavarone, A. T.; Patriksson, A.; van der Spoel, D.; Parks, J. H. J. Am. Chem. Soc. 2007, 129, 6726−6735. (17) Talbot, F. O.; Rullo, A.; Yao, H.; Jockusch, R. A. J. Am. Chem. Soc. 2010, 132, 16156−16164. (18) Wales, T. E.; Engen, J. R. Mass Spectrom. Rev. 2006, 25, 158− 170. (19) Krishna, M. M. G.; Hoang, L.; Lin, Y.; Englander, S. W. Methods 2004, 34, 51−64. (20) Uzawa, T.; Nishimura, C.; Akiyama, S.; Ishimori, K.; Takahashi, S.; Dyson, H. J.; Wright, P. E. Proc. Natl. Acad. Sci., U.S.A. 2008, 105, 13859−13864. (21) Englander, S. W. J. Am. Soc. Mass Spectrom. 2006, 17, 1481− 1489. (22) Kaltashov, I. A.; Engen, J. R.; Gross, M. L. J. Am. Soc. Mass Spectrom. 2006, 17, i−ii. (23) Konermann, L.; Pan, J.; Liu, Y. Chem. Soc. Rev. 2011, 40, 1224− 1234. (24) Smith, D. L.; Deng, Y.; Zhang, Z. J. Mass Spectrom. 1997, 32, 135−146. (25) Winger, B. E.; Light-Wahl, K. J.; Rockwood, A. L.; Smith, R. D. J. Am. Chem. Soc. 1992, 114, 5897−5898. (26) Cheng, X.; Fenselau, C. Int. J. Mass Spectrom. Ion Process. 1992, 122, 109−119. (27) McLafferty, F. W.; Guan, Z.; Haupts, U.; Wood, T. D.; Kelleher, N. L. J. Am. Chem. Soc. 1998, 120, 4732−4740. (28) Weinmann, D. P.; Winkler, H. D. F.; Falenski, J. A.; Koksch, B.; Schalley, C. A. Nat. Chem. 2009, 1, 573−577. (29) Heck, A. J. R.; Jørgensen, T. J. D.; O’Sullivan, M.; von Raumer, M.; Derrick, P. J. J. Am. Soc. Mass Spectrom. 1998, 9, 1255−1266. (30) Herrmann, K. A.; Kuppannan, K.; Wysocki, V. H. Int. J. Mass Spectrom. 2006, 249−250, 93−105. (31) Kang, Y.; Douglas, D. J. J. Am. Soc. Mass Spectrom. 2011, 22, 1187−1196. (32) Nagy, K.; Redeuil, K.; Rezzi, S. Anal. Chem. 2009, 81, 9365− 9371. (33) Campbell, S.; T., R. M.; Marzluff, E. M.; Beauchamp, J. L. J. Am. Chem. Soc. 1995, 117, 12840−12854. (34) Ziegler, B. E.; McMahon, T. B. J. Phys. Chem. A 2010, 114, 11953−11963. (35) Wyttenbach, T.; Bowers, M. T. J. Am. Soc. Mass Spectrom. 1999, 10, 9−14. (36) Geller, O.; Lifshitz, C. J. Phys. Chem. A 2005, 109, 2217−2222. (37) Valentine, S. J.; Clemmer, D. E. J. Am. Chem. Soc. 1997, 119, 3558−3566. (38) Rand, K. D.; Pringle, S. D.; Murphy, J. P.; Fadgen, K. E.; Brown, J.; Engen, J. R. Anal. Chem. 2009, 81, 10019−10028. (39) Cox, H. A.; Julian, R. R.; Lee, S. W.; Beauchamp, J. L. J. Am. Chem. Soc. 2004, 126, 6485−6490. (40) Wyttenbach, T.; Paizs, B.; Barran, P.; Breci, L.; Liu, D.; Suhai, S.; Wysocki, V. H.; Bowers, M. T. J. Am. Chem. Soc. 2003, 125, 13768− 13775. (41) Huang, Y.; Marini, J. A.; McLean, J. A.; Tichy, S. E.; Russell, D. H. J. Am. Soc. Mass Spectrom. 2009, 20, 2049−2057. (42) Jurchen, J. C.; Cooper, R. E.; Williams, E. R. J. Am. Soc. Mass Spectrom. 2003, 14, 1477−1487. (43) He, F.; Marshall, A. G. J. Phys. Chem. A 2000, 104, 562−567. (44) Contreras, A. M.; Polfer, N. C.; Chung, A. C.; Oomens, J.; Eyler, J. R. Int. J. Mass Spectrom. 2011, 297, 162−169. (45) Pan, J.; Han, J.; Borchers, C. H.; Konermann, L. Anal. Chem. 2011, 83, 5386−5393. (46) Abzalimov, R. R.; Kaplan, D. A.; Easterling, M. L.; Kaltashov, I. A. J. Am. Soc. Mass Spectrom. 2009, 20, 1514−1517. (47) Kaltashov, I. A.; Bobst, C. E.; Abzalimov, R. R. Anal. Chem. 2009, 81, 7892−7899.

approach should be beneficial for probing the properties of these desolvated species, especially when considering that previous investigations were largely limited to global exchange data. Intriguingly, we find a clear link between labeling pattern and gas-phase structure. Our data suggest the presence of a short helical segment in RA9K, whereas no such feature appears in RG9K. A direct correlation between structure and labeling behavior has previously been noted in ND3-based (global) HDX studies,36,38 whereas this relationship may not be as straightforward with other labeling agents.5,39,40,42 Our findings suggest qualitative parallels between gas-phase and solutionphase HDX; side chains and backbone termini exchange (or back-exchange)18−20 rapidly, and structural information comes mostly from slow backbone labeling. Nonetheless, the HDX mechanisms in the gas phase33−35 and in solution18−20 are clearly very different. Although gas-phase HDX cannot provide high-resolution structural information per se, the use of deuterium protection patterns, in combination with IMS data, as constraints in computational modeling studies, should prove to be very useful. In future work, it will be interesting to extend the approach employed here to larger biomolecular systems. Attempts in this direction are currently underway in our laboratories, as well as elsewhere.71

■

AUTHOR INFORMATION Corresponding Author *Tel.: (519) 661-2111, ext. 86313 (L.K.), (416) 946-7198 (R.A.J.). E-mail: [email protected] (L.K.), [email protected] (R.A.J.).

■

ACKNOWLEDGMENTS This work was supported by the Natural Sciences and Engineering Research Council of Canada and by the Canada Research Chairs Program.

■

REFERENCES

(1) Fenn, J. B. Angew. Chem., Int. Ed. 2003, 42, 3871−3894. (2) Jarrold, M. F. Annu. Rev. Phys. Chem. 2000, 51, 179−207. (3) Barran, P. E.; Polfer, N. C.; Campopiano, D. J.; Clarke, D. J.; Langridge-Smith, P. R. R.; Langley, R. J.; Govan, J. R. W.; Maxwell, A.; Dorin, J. R.; Millar, R. P.; Bowers, M. T. Int. J. Mass Spectrom. 2005, 240, 273−284. (4) Albrieux, F.; Calvo, F.; Chirot, F.; Vorobyev, A.; Tsybin, Y. O.; Lepère, V.; Antoine, R.; Lemoine, J.; Dugourd, P. J. Phys. Chem. A 2010, 114, 6888−6896. (5) Bohrer, B. C.; Atlasevich, N.; Clemmer, D. E. J. Phys. Chem. B 2011, 115, 4509−4515. (6) Wyttenbach, T.; Grabenauer, M.; Thalassinos, K.; Scrivens, J. H.; Bowers, M. T. J. Phys. Chem. B 2010, 114, 437−447. (7) Ruotolo, B. T.; Giles, K.; Campuzano, I.; Sandercock, A. M.; Bateman, R. H.; Robinson, C. V. Science 2005, 310, 1658−1661. (8) Kohtani, M.; Jones, T. C.; Sudha, R.; Jarrold, M. F. J. Am. Chem. Soc. 2006, 128, 7193−7197. (9) Smith, D. P.; Radford, S. E.; Ashcroft, A. E. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 6794−6798. (10) Gross, D. S.; Schnier, P. D.; Rodriguez-Cruz, S. E.; Fagerquist, S. K.; Williams, E. R. Proc. Natl. Acad. Sci., U.S.A. 1996, 93, 3143−3148. (11) Nagornova, N. S.; Guglielmi, M.; Doemer, M.; Tavernelli, I.; Rothlisberger, U.; Rizzo, T. R.; Boyarkin, O. V. Angew. Chem., Int. Ed. 2011, 50, 5383−5386. (12) Polfer, N. C.; Oomens, J. Phys. Chem. Chem. Phys. 2007, 9, 3804− 3817. (13) Breuker, K.; Oh, H.; Horn, D. M.; Cerda, B. A.; McLafferty, F. W. J. Am. Chem. Soc. 2002, 124, 6407−6419. 377

dx.doi.org/10.1021/ac202730d | Anal. Chem. 2012, 84, 373−378

Analytical Chemistry

Article

(48) Rand, K. D.; Adams, C. M.; Zubarev, R. A.; Jørgensen, T. J. D. J. Am. Chem. Soc. 2008, 130, 1341−1349. (49) Rand, K. D.; Pringle, S. D.; Morris, M.; Engen, J. R.; Brown, J. M. J. Am. Soc. Mass Spectrom. 2011, 22, 1784−1793. (50) Huang, R. Y.; Garai, K.; Frieden, C.; Gross, M. L. Biochemistry 2011, 50, 9273−9282. (51) Jørgensen, T. J. D.; Gårdsvoll, H.; Ploug, M.; Roepstorff, P. J. Am. Chem. Soc. 2005, 127, 2785−2793. (52) Percy, A. J.; Slysz, G. W.; Schriemer, D. C. Anal. Chem. 2009, 81, 7900−7907. (53) Ferguson, P. L.; Pan, J.; Wilson, D. J.; Dempsey, B.; Lajoie, G.; Shilton, B.; Konermann, L. Anal. Chem. 2007, 79, 153−160. (54) Johnson, R. S.; Krylov, D.; Walsh, K. A. J. Mass. Spectrom. 1995, 30, 386−387. (55) Hamuro, Y.; Tomasso, J. C.; Coales, S. J. Anal. Chem. 2008, 80, 6785−6790. (56) Hagman, C.; Hakansson, P.; Buijs, J.; Hakansson, K. J. Am. Soc. Mass Spectrom. 2004, 15, 639−646. (57) Zubarev, R. A.; Zubarev, A. R.; Savitski, M. M. J. Am. Soc. Mass Spectrom. 2008, 19, 753−761. (58) Coon, J. J. Anal. Chem. 2009, 81, 3208−3215. (59) Hagman, C.; Tsybin, Y. O.; Hakansson, P. Rapid Commun. Mass Spectrom. 2006, 20, 661−665. (60) Rand, K. D.; Zehl, M.; Jensen, O. N.; Jørgensen, T. J. D. Anal. Chem. 2009, 81, 5577−5584. (61) Sterling, H. J.; Williams, E. R. Anal. Chem. 2010, 82, 9050− 9057. (62) Sohn, C. H.; Chung, C. K.; Yin, S.; Ramachandran, P.; Loo, J. A.; Beauchamp, J. L. J. Am. Chem. Soc. 2009, 131, 5444−5459. (63) Syrstad, E. A.; Turecek, F. J. Am. Soc. Mass Spectrom. 2005, 16, 208−224. (64) Breuker, K.; Oh, H.; Lin, C.; Carpenter, B. K.; McLafferty, F. W. Proc. Natl. Acad. Sci., U.S.A. 2004, 101, 14011−14016. (65) Vorobyev, A.; Ben Hamidane, H.; Tsybin, Y. O. J. Am. Soc. Mass Spectrom. 2009, 20, 2273−2283. (66) Hofstadler, S. A.; Sannes-Lowery, K. A.; Griffey, R. H. Rapid Commun. Mass Spectrom. 1999, 13, 1971−1979. (67) Weis, D. D.; Engen, J. R.; Kass, I. J. J. Am. Soc. Mass Spectrom. 2006, 17, 1700−1703. (68) Kruger, N. A.; Zubarev, R. A.; Horn, D. M.; McLafferty, F. W. Int. J. Mass Spectrom. 1999, 185−187, 787−793. (69) Anderegg, R. J.; Wagner, D. S.; Stevenson, C. L.; Borchardt, R. T. J. Am. Soc. Mass Spectrom. 1994, 5, 425−433. (70) Fesinmeyer, R. M.; Peterson, E. S.; Dyer, R. B.; Andersen, N. H. Protein Sci. 2000, 14, 2324−2332. (71) Castro, S.; Breuker, K.; Saa, S. O.; Kong, X.; McLafferty, F. W. Presented at the ASMS Conference, Denver, CO, 2011; Poster No. TP499.

378

dx.doi.org/10.1021/ac202730d | Anal. Chem. 2012, 84, 373−378