Deuterium Exchange for Distinguishing

Mar 17, 2017 - Department of Medicinal Chemistry, University of Washington, Seattle, Washington 98195, United States. Anal. Chem. , 2017, 89 (8), pp 4...
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Gas-Phase Hydrogen/Deuterium Exchange for Distinguishing Isomeric Carbohydrate Ions Sanjit S. Uppal, Sarah E. Beasley, Michele Scian, and Miklos Guttman* Department of Medicinal Chemistry, University of Washington, Seattle, Washington 98195, United States S Supporting Information *

ABSTRACT: The structural diversity of carbohydrates presents a major challenge for glycobiology and the analysis of glycoconjugates. Mass spectrometry has become a primary tool for glycan analysis thanks to its speed and sensitivity, but the information content regarding the glycan structure of protonated glycoconjugates is hindered by the inability to differentiate linkage and stereoisomers. Here, we examine a variety of protonated carbohydrate structures by gas-phase hydrogen/ deuterium exchange (HDX) to discover that the exchange rates are distinct for isomeric carbohydrates with even subtle structural differences. By incorporating an internal exchange standard, HDX could effectively distinguish all linkage and stereoisomers that were examined and presents a mass spectrometry-based approach for glycan structural analysis with immense potential.

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Here, we investigate the potential of gas-phase hydrogen/ deuterium exchange (HDX) to differentiate isomeric carbohydrate structures. Gas-phase HDX has previously been used for determining the number of labile protons, distinguishing structural isomers of small molecules, and investigating the gas-phase conformations of peptides and proteins.17−23 Ions are mixed with a deuterated exchange reagent and the resulting incorporation of deuterium is monitored by the shift in the mass envelope.24 Because the exchange rate of each hydrogen depends on its position, basicity, and involvement in hydrogen bonds,25−28 HDX presents a potential method for discrimination of isomeric carbohydrate structures as they contain many hydroxyl groups with labile protons.

ver 50% of the human proteome is estimated to be glycosylated, and its glycosylation plays a role in nearly all aspects of biology including cell−cell communication, differentiation and development, modulating enzymatic activity, governing immune regulation, and host−pathogen interactions.1 Despite their importance to clinical diagnostics and therapeutic development, analysis of glycoconjugates has been hampered by the structural complexity of carbohydrates. All biologically relevant glycan chains are composed of relatively few types of monosaccharides, but the large number of possible branching patterns and linkage stereochemistry results in an enormous diversity of glycan structures.2,3 Unlike nucleic acids and proteins, for which technologies have enabled rapid sequencing, glycan structural elucidation remains a very involved and specialized task, often requiring several orthogonal methods. Mass spectrometry (MS) has become a widespread method for monitoring protein glycosylation that is capable of rapid glycan profiling on a site-specific level.4,5 However, the isobaric nature of most sugar subunits within glycan chains presents a major limitation for MS. Tandem MS/MS-based approaches cannot always effectively identify branching patterns and provide little information on stereochemistry. For this reason, glycans in MS/MS spectra are often depicted as Hex (for any hexose) and HexNAc (for any N-acetyl hexose) as the identity of the specific sugar from the MS data is not determined. Compositions are often inferred with the knowledge of biochemical pathways; however, recent studies have illustrated how this assumption can be misleading.6,7 Ion mobility mass spectrometry (IM-MS) has emerged as a promising MS-based tool for glycobiology.6,8−16 Ions are pushed through an inert gas to separate them by their collision cross section, which can adequately resolve many, but not all, types of carbohydrate isomers. © 2017 American Chemical Society

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RESULTS AND DISCUSSION To evaluate the utility of gas-phase HDX for distinguishing carbohydrate ions, we first compared the deuterium uptake kinetics of three isobaric monosaccharides: N-acetylglucosamine (GlcNAc), N-acetylgalactosamine (GalNAc), and Nacetylmannosamine (ManNAc) using a Waters Synapt HDMS mass spectrometer modified for gas-phase deuterium exchange in a Traveling Wave Ion Guide (TWIG)21 (see Figure S1). Each monosaccharide forms an abundant oxonium ion (m/z 204)29 that was monitored for deuterium uptake (Figure 1a). By altering the speed at which the ions pass through the region with the exchange reagent, in this case deuterated ammonia (ND3), a temporal range of 2 orders of magnitude was sampled. The HDX kinetics for the three monosaccharides are remarkably distinct (Figure 1b). All three undergo rapid Received: February 23, 2017 Accepted: March 17, 2017 Published: March 17, 2017 4737

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to differentiate even very closely related carbohydrates. Examination of the P1 HDX kinetics collected on different days over the course of several months also showed excellent consistency demonstrating the general reproducibility of gasphase HDX (Figure S2). We examined whether the collision energy (CE) used to generate the oxonium ions impacted the observed kinetics of HDX. The data at higher CE revealed a second isotopic mass envelope starting from m/z 207 with an accurate mass consistent with the loss of H2O and the addition of ND3 (Figure S3). This is not unwarranted as water loss is frequently observed during CID29 and noncovalent complexing with ND3 is a necessary step in the HDX exchange mechanism.25,27,28 This highlights the need for high resolving power to circumvent potential misinterpretations of deuterium uptake that can occur with lower resolution instruments that cannot sufficiently resolve these types of nearly overlapping isotopic mass envelopes. Examination of the deuterium uptake at the first time point (0.1 ms) as a function of CE shows a clear trend in which higher CE leads to less exchange (Figure 2a). Similar results were obtained for ManNAc and GalNAc (Figure S4a,b). To test whether the CE setting has systematic effects on the observed exchange rates, we examined the HDX profiles of two

Figure 1. (a) HDX spectra for GlcNAc oxonium ion undeuterated and after 0.1, 1.2, 2.5, and 9.9 ms of deuterium exchange. Purple arrows indicate the centroid of the isotopic peaks. (b) Deuterium uptake plots for the oxonium ions of GlcNAc (blue dashed line), GalNAc (red solid line), and ManNAc (green dotted line). The centroids at each time point are plotted with error bars showing the standard deviation from triplicate measurements. (c) Plots of the corresponding exchange control data (peptide P1) from each monosaccharide data set.

exchange of one of the four labile protons by 0.1 ms. After 0.4 ms, the remaining protons begin to exchange and the longer time points reveal significant differences. At the last time point (9.9 ms), the deuterium uptake for GlcNAc, GalNAc, and ManNAc is 3.91 ± 0.04, 3.44 ± 0.02, and 3.78 ± 0.03, respectively. Therefore, a change in stereochemistry at just a single position is enough to alter the HDX kinetics of monosaccharide oxonium ions. To account for any potential differences in instrument conditions that might offset the apparent kinetics of HDX, we included an internal “exchange” standard. Peptide P1 (HHHHHHIIKIIK), commonly used to monitor deuterium migration during HDX analysis,30 served as a suitable exchange control. The P1 HDX data was cosampled with each monosaccharide and showed impressive consistency (Figure 1c). Therefore, the differences in the observed HDX rates for the monosaccharides were not due to instrumental variations but rather structural differences in the carbohydrate ions that alter the exchangeability of their labile protons. This demonstrates that the various carbohydrate oxonium ions retain their distinct structures in the gas phase and HDX can effectively be used

Figure 2. (a) Deuterium uptake plots for the GlcNAc oxonium ion obtained with CE values ranging from 2 (purple) to 15 eV (red). Error bars represent standard deviation from triplicate measurements. The inset shows the deuterium uptake of the earliest time point as a function of CE. (b) Drift time of the GlcNAc oxonium ion generated with the same CE values as in (a). The inset shows the drift time as a function of CE. 4738

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galactose (Gal) or mannose (Man) were examined. The differences in deuterium uptake among the structures were dramatic (Figure 4a). Galβ1−4GlcNAc showed very little

noncarbohydrate compounds, Tris and Bis-tris propane. Within the same range of CEs, there was no significant difference in the HDX profiles for either compound (Figure S4c,d). Therefore, the CE used to generate the oxonium ions is likely influencing their gas-phase structure, which in turn changes the exchangeability of certain labile protons. IM-MS was used as an orthogonal technique to interrogate the effect of CE on the gas-phase conformation of the GlcNAc oxonium ion. The drift times of GlcNAc using the same range of CEs showed a slight but gradual shift (Figure 2b). The shift in drift times did not result from global drift time offsets as the drift time of all other ions, including Leucine Enkephalin used as a reference ion for mass calibration, were not affected by the altered CE. The difference in drift time for the GlcNAc oxonium ion as a function of CE resembles the trend observed by HDX (Figure 2b inset). The specific effect of the CE on the oxonium ion structure remains to be determined, but this highlights the need to consider carbohydrate ions ensembles of structures that can vary based on the CE used to generate them. It is important to note that for the longer HDX time points the CE energy had no significant effect on the observed level of deuteration. Thus, the HDX at the longest time point (9.9 ms) provides information for distinguishing carbohydrate structures that circumvents the caveat of CE induced effects. A variety of larger oligosaccharides were examined to assess the robustness of MS/MS-HDX for distinguishing compositional isomers. Each carbohydrate was selected and fragmented to provide a strong signal for the GlcNAc/GalNAc oxonium ions (m/z 204), and the deuterium uptake at 9.9 ms was measured (Figure 3). The comparisons demonstrate that various N-acetyl

Figure 4. Deuterium uptake plots for oxonium ions of various disaccharides: GlcNAcβ1−3Gal (green), GlcNAcβ1−2Man (orange), Galβ1−3GalNAc (teal), Galβ1−3GlcNAc (purple), and Galβ1− 4GlcNAc (red) (a); sialylated trisaccharides: Neu5Acα2−3Galβ1− 4GlcNAc (gray), Galβ1−3(Neu5Acα2−6)GlcNAc (red), Neu5Acα2− 6Galβ1−4GlcNAc (blue), and Neu5Acα2−3Galβ1−3GlcNAc (purple) (b). Structures are indicated next to each curve with symbol nomenclature as shown in Figure 3.

deuterium uptake at the longest time point compared to Galβ1−3GlcNAc (1.85 ± 0.01 vs 3.05 ± 0.03 Da). The GlcNAcβ1−2Man, GlcNAcβ1−3Gal, and Galβ1−3GalNAc ions also had very distinct HDX profiles. As with the monosaccharides, the CE used to generate the oxonium ions had a minor influence on the observed HDX profile but did not significantly affect the exchange at the longest time point (Figure S5). Various sialylated structures with different linkages of Nacetylneuraminic acid (Neu5Ac) were also examined. Again, drastic differences in deuterium uptake kinetics were apparent that could readily distinguish the various structures (Figure 4b). The diversity of the exchange profiles shows the importance of sampling multiple time points as some carbohydrate structures have zones of similar exchange behavior. For example, Neu5Acα2−6Galβ1−4GlcNAc and Galβ1−3(Neu5Acα2−6)GlcNAc have identical exchange behavior at short time points and only deviate at the later time points. To effectively discern all 4 sialylated structures examined here, sampling at least two different time points is required. As a comparison, we tested IM-MS on the Waters Synapt G2 platform for distinguishing the various mono-, di-, and trisaccharide oxonium ions. While many of the structures are resolved to different extents, some are not (Figure 5). For example, the β1−3 and β1−4 linkage isomers of Gal−GlcNAc have nearly identical drift times. The same is true for the Neu5Acα2−3Galβ1−3GlcNAc and Neu5Acα2−6Galβ1− 4GlcNAc oxonium ions. In contrast, the HDX kinetics of all the structures examined were remarkably distinct, illustrating

Figure 3. Deuterium uptake at 9.9 ms is shown for the HexNAc oxonium ion (m/z 204) fragmented from various glycan precursors including a glycopeptide and a glycolipid (diamond and square, respectively). Symbol and linkage nomenclature for the glycan structures are shown on the left. Error bars show the standard deviation from triplicate measurements. The deuterium uptake of the 204 m/z oxonium ion can be used to distinguish whether the precursor glycan contained GlcNAc, GalNAc, or a mixture as shown by blood group type A antigen pentaose type 1, which contains both (star).

hexosamine (HexNAc) fragment ions can be distinguished and identified by their deuterium uptake regardless of the starting structure of the carbohydrate, including glycopeptides and glycolipids. We next sought to test whether oligosaccharides with identical composition but different linkages also had distinct HDX kinetics. A series of disaccharide oxonium ions (m/z 366) containing GlcNAc or GalNAc with various attachments to 4739

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oxide and deuterated methanol may be more effective for HDX of carbohydrate anions.20,35 Certain types of protonated glycan ions have been shown to undergo gas-phase linkage rearrangements during CID (e.g., fucose migration), which can complicate the interpretation of tandem mass spectra.5,36 While no such linkage rearrangements were apparent for the ions examined here, this caveat may potentially complicate the analysis of other types of glycan ions. HDX could be used as a tool to help elucidate the mechanism of glycan linkage rearrangements, which if understood, may provide more insight into the precursor glycan structure. Permethylation is a common approach with MS to prevent any gas-phase rearrangements and obtain more detailed structural information.5,37 However, permethylation is incompatible with HDX as all of the labile protons are lost.

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CONCLUSIONS This study demonstrates the utility of gas-phase HDX for distinguishing compositional and linkage isomers of carbohydrates and glycoconjugates along with the caveats (e.g., influence of the collision energy) that need to be appreciated. Incorporation of an internal exchange control, as presented here, should enable reproducible exchange profiles as it can correct for any offsets with instrument conditions and normalize HDX data obtained on different platforms. For the current study, such corrections were not necessary as the exchange conditions were remarkably consistent over the course of months. HDX could distinguish both mono- and oligosaccharides, several of which could not be effectively resolved by ion mobility. Gas-phase HDX is compatible with a LC-MS time scale23 and can be performed in tandem with orthogonal MS techniques including ion mobility.21,22 MSbased platforms have enabled rapid, site-specific, glycosylation analysis, and the integration of HDX can provide a higher level of information for identifying specific carbohydrate structures.

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EXPERIMENTAL METHODS Reagents. ND3 (99%) and D2O (99.99%) were purchased from Cambridge isotope laboratories (Tewksbury, MA, USA). Carbohydrate standards, GlcNAc, ManNAc, GalNAc, LacNAc, and bovine Ribonuclease B (RNaseB), were purchased from Sigma-Aldrich (St. Louis, MO, USA); GlcNAcβ1-2Man, Fucα1-6GlcNAc, Galβ1-3GalNAc, Gaβ1-3GlcNAc, 3′- and 6′sialyl N-acetyllactosamine, lacto-N-biose, lacto-N-triose, and ganglioside GM1 were from Carbosynth (Compton, UK). Sialylated tetraose types 1 and 2, globo-H hexaose, blood group A antigen pentaose type 1, and blood group H antigen pentaose type 1 were from Elicityl (Crolles, France). For glycopeptide preparation, 10 mg of bovine RNaseB (Sigma-Aldrich) was reduced with 10 mM dithiothreitol (DTT) at 60 °C for 1 h and alkylated with 20 mM iodoacetamide (IAM) for 1 h at room temperature in the dark. IAM was quenched with another addition of 10 mM DTT, and the sample was digested with 0.25 μg of chymotrypsin (Roche) in 50 mM Tris, pH 8.0, at 37 °C for 8 h. The glycopeptide spanning residues 56−72 was purified by HPLC (DELTA-PAK column, 300 mm × 19 mm, 15 μm, 300 Å C18, Waters, Milford MA) using a linear gradient of 5% to 50% acetonitrile in 0.1% trifluoroacetic acid over 40 min. The glycopeptide was lyophilized and stored at −80 °C. For infusions, RNaseB glycopeptide 56−72 was resuspended in

Figure 5. IM-MS drift time distributions of monosaccharide (a), disaccharide (b), and sialylated trisaccharide (c) oxonium ions. For some larger precursor glycans, the fragment ion monitored is highlighted in blue. Glycan structures are shown next to each trace as outlined in Figure 3. Dashed lines are added for a reference to clarify slight shifts in the drift times. The IM traveling wave velocity was set to 1000 m/s for (a) and (b) and 650 m/s for (c).

the advantages of HDX over existing MS methods for glycan structural analysis. We examined the HDX behavior of sodium adducts of the various glycans analyzed. In all cases, the sodium adduct had very low deuterium uptake showing little promise for distinguishing isomeric glycans (Figure S6b). This is consistent with the slow HDX kinetics of sodium adducted ions of other classes of compounds.31−34 We also examined the HDX profile of several glycans as deprotonated anions and found absolutely no deuterium incorporation (Figure S6c). ND3 has previously been shown to be a poor reagent for HDX with anions due to its relatively low acidity, and other reagents such as deuterium 4740

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spectrometer. Ten μM glycans in 0.1% formic acid were directly infused and sampled with the same source conditions as used for HDX data collection. The drift gas (N2) was set to 90 mL/min, and the IM TW velocity and height was 1000 m/s and 40 V, except for the sialylated trisaccharide comparisons for which the TW velocity was 650 m/s. Protonated precursor ions were isolated and fragmented in the trap TWIG. Drift data was fit to Gaussian distributions using custom scripts in Excel (Microsoft, Redmond WA).

0.1% formic acid for a final concentration of approximately 1 μM. Mass Spectrometry. All HDX experiments were performed on a Waters Synapt G1 quadrupole-time-of-flight (QTOF) mass spectrometer. Instrument modifications to enable infusion of ND3 have been described previously.21 Briefly, the gas in the transfer traveling wave ion guide (TWIG) is replaced with deuterated ammonia (ND3) to enable deuterium exchange (Figure S1). Additional instrument modifications included a thermal mass flow controller (Aalborg, Orangeburg NY) for more precise infusion of ND3. Steel lines, valves, and unions were from purchased from Swagelok (Solon, OH, USA). The ND3 lecture bottle was fitted with a stainless steel regulator (Matheson Gas, Basking Ridge, New Jersey, USA). All gas plumbing for ND3 infusion was stainless steel with corrosion resistant components using either polytetrafluoroethylene (PTFE) or nylon seals. All lines were washed in 50% isopropanol and flushed thoroughly with N2 prior to use. Carbohydrates were resuspended in LC-MS grade optima water and diluted to a working concentration of 50−200 μM in 0.1% formic acid, except for ganglioside GM1 which was resuspended in 50% acetonitrile, 0.1% formic acid. Samples were infused at a flow rate of 10 μL/min with a syringe pump. With all samples, peptide P1 (1.5 μM in 99% isopropanol, 0.1% formic acid) was sampled through the secondary (lock mass) inlet at 5 μL/min. Tris and Bis-tris propane were infused at a concentration of 200 μM in 0.1% formic acid. For each infusion, the resolving quadruple (resolution of 4) was used to isolate the protonated precursor ion, and scans were collected using several collision energy (CE) settings in the trap TWIG. Each CE setting was collected in an interleaved manner with 4 s/scan. The exchange standard (“lock channel”) was sampled with 4 s scans in each acquisition. We initially noticed that, without IM gas flow, the IM TWIG wave velocity settings influenced the level of deuterium exchange, probably due to ND3 diffusing into the IM TWIG. This was completely alleviated by applying a moderate IM gas flow of N2 at 5 mL/ min. IM trap height and velocities were set to 5 V and 100 m/s. Transfer TWIG lens/aperture voltages were set to 5 and 30 V to help reduce the abundance of ammonia adducts entering the TOF region. A TW height of 8 V in the transfer TWIG was found to be optimal for efficient ion transmission and to ensure that no ions fall behind the wave leading to increased transit time (“roll-over”).38 Capillary, sample cone, and extraction cone voltages were 2.5 kV, 30 V, and 3.3 V, respectively. Source and desolvation temperatures were 100 and 250 °C. All data was collected with “W” TOF optics in positive mode and “V” optics in negative mode. A series of acquisitions were collected using a transfer CE of 5 eV, a wave height of 8 V, and wave velocities of 512, 362, 256, 181, 128, 90, 64, 45, 32, 23, 16, 11, and 8 m/s to sample a temporal range of ∼0.1 to 10 ms.21 The ND3 mass flow controller was adjusted to precisely flow 1 mL/min into the transfer TWIG for all HDX measurements. Undeuterated samples were recorded with the same settings using a wave velocity of 512 m/s with Ar flowing into the transfer TWIG. As an additional control, fully deuterated samples were prepared in 95% D2O and infused as described above after rinsing the lines and the ESI source with D2O for 1 min. Data was batch converted using scripts in UniDec39 and analyzed using HXExpress v2.40 Ion Mobility Spectrometry. Ion mobility measurements were performed on a Waters Synapt G2-Si Q-TOF mass

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.analchem.7b00683. Schematic of the Synapt HDMS for gas-phase HDX; reproducibility of gas-phase HDX; second isotopic species of GlcNAc at higher CE; CE dependence on HDX for ManNAc, GalNAc, Tris, and Bis-tris propane; CE dependence on HDX for disaccharide ions; HDX spectra of sialyl N-acetyllactosamine cations and anions (PDF)

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Miklos Guttman: 0000-0003-2419-1334 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors wish to thank Jeff Brown, Dale Whittington, and J. Scott Edgar for assistance with instrument modifications. We are grateful to Michael Marty and Tim Allison for assistance with data conversion scripts. We also thank Elizabeth Komives for helpful discussions and assistance with glycopeptide purification.

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REFERENCES

(1) Varki, A.; Cummings, R.; Esko, J.; Freeze, H.; Stanley, P.; Bertozzi, C.; Hart, G. W.; Etzler, M. Essentials of Glycobiology, 2nd ed.; Cold Spring Harbor Laboratory Press: Plainview, NY, 2009. (2) Marino, K.; Bones, J.; Kattla, J. J.; Rudd, P. M. Nat. Chem. Biol. 2010, 6 (10), 713−23. (3) Cummings, R. D.; Pierce, J. M. Chem. Biol. 2014, 21 (1), 1−15. (4) Medzihradszky, K. F. Methods Enzymol. 2005, 405, 116−38. (5) Leymarie, N.; Zaia, J. Anal. Chem. 2012, 84 (7), 3040−3048. (6) Both, P.; Green, A. P.; Gray, C. J.; Sardzik, R.; Voglmeir, J.; Fontana, C.; Austeri, M.; Rejzek, M.; Richardson, D.; Field, R. A.; Widmalm, G.; Flitsch, S. L.; Eyers, C. E. Nat. Chem. 2014, 6 (1), 65− 74. (7) Marino, F.; Bern, M.; Mommen, G. P.; Leney, A. C.; van Gaansvan den Brink, J. A.; Bonvin, A. M.; Becker, C.; van Els, C. A.; Heck, A. J. J. Am. Chem. Soc. 2015, 137 (34), 10922−5. (8) Dwivedi, P.; Bendiak, B.; Clowers, B. H.; Hill, H. H., Jr. J. Am. Soc. Mass Spectrom. 2007, 18 (7), 1163−75. (9) Fenn, L. S.; McLean, J. A. Phys. Chem. Chem. Phys. 2011, 13 (6), 2196−205. (10) Li, H.; Bendiak, B.; Siems, W. F.; Gang, D. R.; Hill, H. H., Jr. Anal. Chem. 2013, 85 (5), 2760−9. (11) Huang, Y.; Dodds, E. D. Anal. Chem. 2013, 85 (20), 9728−35. (12) Hofmann, J.; Hahm, H. S.; Seeberger, P. H.; Pagel, K. Nature 2015, 526 (7572), 241−4. 4741

DOI: 10.1021/acs.analchem.7b00683 Anal. Chem. 2017, 89, 4737−4742

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Analytical Chemistry (13) Plasencia, M. D.; Isailovic, D.; Merenbloom, S. I.; Mechref, Y.; Clemmer, D. E. J. Am. Soc. Mass Spectrom. 2008, 19 (11), 1706−1715. (14) Pu, Y.; Ridgeway, M. E.; Glaskin, R. S.; Park, M. A.; Costello, C. E.; Lin, C. Anal. Chem. 2016, 88 (7), 3440−3. (15) Guttman, M.; Lee, K. K. Anal. Chem. 2016, 88 (10), 5212−7. (16) Hinneburg, H.; Hofmann, J.; Struwe, W. B.; Thader, A.; Altmann, F.; Varon Silva, D.; Seeberger, P. H.; Pagel, K.; Kolarich, D. Chem. Commun. (Cambridge, U. K.) 2016, 52 (23), 4381−4. (17) Cheng, X.; Fenselau, C. Int. J. Mass Spectrom. Ion Processes 1992, 122, 109−119. (18) Suckau, D.; Shi, Y.; Beu, S. C.; Senko, M. W.; Quinn, J. P.; Wampler, F. M., 3rd; McLafferty, F. W. Proc. Natl. Acad. Sci. U. S. A. 1993, 90 (3), 790−3. (19) Hemling, M. E.; Conboy, J. J.; Bean, M. F.; Mentzer, M.; Carr, S. A. J. Am. Soc. Mass Spectrom. 1994, 5 (5), 434−42. (20) Zhang, J.; Brodbelt, J. S. J. Am. Chem. Soc. 2004, 126 (18), 5906−19. (21) Rand, K. D.; Pringle, S. D.; Murphy, J. P., 3rd; Fadgen, K. E.; Brown, J.; Engen, J. R. Anal. Chem. 2009, 81 (24), 10019−28. (22) Donohoe, G. C.; Khakinejad, M.; Valentine, S. J. J. Am. Soc. Mass Spectrom. 2015, 26 (4), 564−76. (23) Mistarz, U. H.; Brown, J. M.; Haselmann, K. F.; Rand, K. D. Anal. Chem. 2014, 86 (23), 11868−76. (24) Campbell, S.; Rodgers, M. T.; Marzluff, E. M.; Beauchamp, J. L. J. Am. Chem. Soc. 1995, 117 (51), 12840−12854. (25) Ausloos, P.; Lias, S. G. J. Am. Chem. Soc. 1981, 103, 3641−3647. (26) Gard, E.; Green, M. K.; Bregar, J.; Lebrilla, C. B. J. Am. Soc. Mass Spectrom. 1994, 5 (7), 623−31. (27) Wyttenbach, T.; Bowers, M. T. J. Am. Soc. Mass Spectrom. 1999, 10, 9−14. (28) Evans, S. E.; Lueck, N.; Marzluff, E. M. Int. J. Mass Spectrom. 2003, 222, 175−187. (29) Halim, A.; Westerlind, U.; Pett, C.; Schorlemer, M.; Rüetschi, U.; Brinkmalm, G.; Sihlbom, C.; Lengqvist, J.; Larson, G.; Nilsson, J. J. Proteome Res. 2014, 13 (12), 6024−6032. (30) Rand, K. D.; Jorgensen, T. J. Anal. Chem. 2007, 79 (22), 8686− 93. (31) Kaltashov, I. A.; Doroshenko, V. M.; Cotter, R. J. Proteins: Struct., Funct., Genet. 1997, 28 (1), 53−58. (32) Reyzer, M. L.; Brodbelt, J. S. J. Am. Soc. Mass Spectrom. 2000, 11 (8), 711−21. (33) Jurchen, J. C.; Cooper, R. E.; Williams, E. R. J. Am. Soc. Mass Spectrom. 2003, 14 (12), 1477−87. (34) Chen, Y.; Yue, L.; Li, Z.; Ding, X.; Wang, L.; Dai, X.; Fang, X.; Pan, Y.; Ding, C.-F. Anal. Methods 2015, 7 (13), 5551−5556. (35) Mo, J.; Todd, G. C.; Hakansson, K. Biopolymers 2009, 91 (4), 256−64. (36) Harvey, D. J.; Mattu, T. S.; Wormald, M. R.; Royle, L.; Dwek, R. A.; Rudd, P. M. Anal. Chem. 2002, 74 (4), 734−40. (37) Reinhold, V.; Zhang, H.; Hanneman, A.; Ashline, D. Mol. Cell. Proteomics 2013, 12 (4), 866−873. (38) Giles, K.; Pringle, S. D.; Worthington, K. R.; Little, D.; Wildgoose, J. L.; Bateman, R. H. Rapid Commun. Mass Spectrom. 2004, 18 (20), 2401−14. (39) Marty, M. T.; Baldwin, A. J.; Marklund, E. G.; Hochberg, G. K.; Benesch, J. L.; Robinson, C. V. Anal. Chem. 2015, 87 (8), 4370−6. (40) Guttman, M.; Weis, D. D.; Engen, J. R.; Lee, K. K. J. Am. Soc. Mass Spectrom. 2013, 24 (12), 1906−12.

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