Protein Hydrogen Exchange Measured at Single-Residue Resolution

E-mail [email protected], phone 1-617-373-4291, fax 1-617-373-2855 ... a popular method for measuring the solution hydrogen (1H/2H) exchange (HX) of ...
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Anal. Chem. 2009, 81, 5577–5584

Accelerated Articles Protein Hydrogen Exchange Measured at Single-Residue Resolution by Electron Transfer Dissociation Mass Spectrometry Kasper D. Rand,*,† Martin Zehl,‡ Ole N. Jensen, and Thomas J. D. Jørgensen* Department of Biochemistry and Molecular Biology, University of Southern Denmark, Campusvej 55, DK-5230 Odense M, Denmark Because of unparalleled sensitivity and tolerance to protein size, mass spectrometry (MS) has become a popular method for measuring the solution hydrogen (1H/2H) exchange (HX) of biologically relevant protein states. While incorporated deuterium can be localized to different regions by pepsin proteolysis of the labeled protein, the assignment of deuteriums to individual residues is typically not obtained, thereby limiting a detailed understanding of HX and the dynamics of protein structure. Here we use gas-phase fragmentation of peptic peptides by electron transfer dissociation (ETD) to measure the HX of individual amide linkages in the amyloidogenic protein β2-microglobulin. A comparison of the deuterium levels of 60 individual backbone amides of β2-microglobulin measured by HXETD-MS analysis to the corresponding values measured by NMR spectroscopy shows an excellent correlation. The deuterium labeling pattern of β2microglobulin is retained in the gaseous fragment ions by employing mild declustering conditions for electrospray ionization. A recently developed model peptide is used to arrive at such ion source declustering conditions that prevent the occurrence of intramolecular gas-phase hydrogen (1H/2H) migration (i.e., hydrogen scrambling). This article demonstrates that ETD can be implemented in a mass spectrometric method to monitor the conformational dynamics of proteins in solution at single-residue resolution. Protein function is intimately tied to the dynamics of protein structure and the hydrogen exchange (HX) of amides of the * To whom correspondence should be addressed. E-mail [email protected], phone 1-617-373-4291, fax 1-617-373-2855 (K.D.R.); e-mail [email protected], phone (+45) 6550 2414, fax (+45) 6550 2467 (T.J.D.J.). 10.1021/ac9008447 CCC: $40.75  2009 American Chemical Society Published on Web 06/16/2009

protein backbone provides a valuable probe of such conformational dynamics. Measurement of protein hydrogen (1H/2H) exchange, pioneered by K. Linderstrøm-Lang and subsequently S.W. Englander, has since its inception in the 1950s yielded valuable insight into the myriad of local and global conformational fluctuations of proteins that govern folding, function and interactions.1-3 Mass spectrometry (MS) is increasingly being utilized to monitor protein hydrogen exchange,4 as this technique presents some advantages over other traditional spectroscopic methods (i.e., NMR and FT-IR). Most importantly, mass spectrometry offers the ability to perform analysis with very small quantities of protein (a few nanomoles), the ability to study large (>40 kDa) proteins, the ability to analyze different protein components in a mixture, and the ability to resolve differentially labeled coexisting populations of a given protein.4,5 These capabilities have been exploited to study a range of important but elusive aspects of molecular biology, including aggregation prone proteins and proteins not amenable to crystallization,6-10 large protein † Present address: The Barnett Institute of Chemical and Biological Analysis, Northeastern University, Boston, MA. ‡ Present address: Department of Pharmacognosy, University of Vienna, Althanstrasse 14, A-1090 Vienna, Austria. (1) Hvidt, A.; Linderstrom-Lang, K. Biochim. Biophys. Acta 1954, 14, 574– 575. (2) Hvidt, A.; Nielsen, S. O. Adv. Protein Chem. 1966, 21, 287–386. (3) Englander, S. W.; Kallenbach, N. R. Q. Rev. Biophys. 1983, 16, 521–655. (4) Wales, T. E.; Engen, J. R. Mass Spectrom. Rev. 2006, 25, 158–170. (5) Hoofnagle, A. N.; Resing, K. A.; Ahn, N. G. Annu. Rev. Biophys. Biomol. Struct. 2003, 32, 1–25. (6) Hochrein, J. M.; Wales, T. E.; Lerner, E. C.; Schiavone, A. P.; Smithgall, T. E.; Engen, J. R. Biochemistry 2006, 45, 7733–7739. (7) Iacob, R. E.; Pene-Dumitrescu, T.; Zhang, J.; Gray, N. S.; Smithgall, T. E.; Engen, J. R. Proc. Natl. Acad. Sci. U.S.A. 2009, 106, 1386–1391. (8) Mitchell, J. l.; Trible, R. P.; Emert-Sedlak, L. A.; Weis, D. D.; Lerner, E. C.; Applen, J. J.; Sefton, B. M.; Smithgall, T. E.; Engen, J. R. J. Mol. Biol. 2007, 366, 1282–1293.

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assemblies such as viral capsids and molecular motors,11,12 protein folding intermediates,13 and cooperative unfolding/folding events of natively folded proteins that are important for function or amyloidogenicity.14-17 Hydrogen exchange mass spectrometry (HX-MS) enables sensitive measurement of hydrogen/deuterium exchange by monitoring the resulting change in the mass of a protein.4,5 Incorporated deuterium is localized to different regions by mass analysis of peptides generated by pepsin proteolysis of the labeled protein. Single-residue resolution is, however, typically not obtained by this approach, and localization of deuterium labeling is confined to peptide segments typically 6-10 residues in length. Thus, while hydrogen exchange imprints information on local conformational fluctuations into each individual amide group of the protein backbone, such information is only resolved to intermediate resolution in a classical HX-MS experiment. Considerable effort has been devoted to identify and apply a gas-phase fragmentation method that allows the deuterium labeling pattern from solution to be deduced from the masses of fragment ions and thus providing residue-specific information. The occurrence of gas-phase hydrogen (1H/2H) scrambling is, however, the main obstacle for achieving this goal. Hydrogen scrambling results from an interchange of protons and deuterons among all exchangeable sites, and this process occurs upon excessive vibrational excitation of protonated peptides in the gas phase. This phenomenon precludes the use of collision-induced dissociation, the most widely used gas-phase fragmentation technique, as a general MS method to obtain information about protein solution deuteration patterns from peptic peptides.18-26 (9) Rand, K. D.; Jorgensen, T. J. D.; Olsen, O. H.; Persson, E.; Jensen, O. N.; Stennicke, H. R.; Andersen, M. D. J. Biol. Chem. 2006, 281, 23018–23024. (10) Rist, W.; Graf, C.; Bukau, B.; Mayer, M. P. J. Biol. Chem. 2006, 281, 16493– 16501. (11) Lanman, J.; Lam, T. T.; Emmett, M. R.; Marshall, A. G.; Sakalian, M.; Prevelige, P. E., Jr. Nat. Struct. Mol. Biol. 2004, 11, 676–677. (12) Lisal, J.; Lam, T. T.; Kainov, D. E.; Emmett, M. R.; Marshall, A. G.; Tuma, R. Nat. Struct. Mol. Biol. 2005, 12, 460–466. (13) Miranker, A.; Robinson, C. V.; Radford, S. E.; Aplin, R. T.; Dobson, C. M. Science 1993, 262, 896–900. (14) Dumoulin, M.; Last, A. M.; Desmyter, A.; Decanniere, K.; Canet, D.; Larsson, G.; Spencer, A.; Archer, D. B.; Sasse, J.; Muyldermans, S.; Wyns, L.; Redfield, C.; Matagne, A.; Robinson, C. V.; Dobson, C. M. Nature 2003, 424, 783–788. (15) Jorgensen, T. J. D.; Cheng, L.; Heegaard, N. H. H. Int. J. Mass Spectrom. 2007, 268, 207–216. (16) Truhlar, S. M. E.; Torpey, J. W.; Komives, E. A. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 18951–18956. (17) Tsutsui, Y.; Kuri, B.; Sengupta, T.; Wintrode, P. L. J. Biol. Chem. 2008, 283, 30804–30811. (18) Demmers, J. A. A.; Rijkers, D. T. S.; Haverkamp, J.; Killian, J. A.; Heck, A. J. R. J. Am. Chem. Soc. 2002, 124, 11191–11198. (19) Ferguson, P. L.; Pan, J.; Wilson, D. J.; Dempsey, B.; Lajoie, G.; Shilton, B.; Konermann, L. Anal. Chem. 2007, 79, 153–160. (20) Harrison, A. G.; Yalcin, T. Int. J. Mass Spectrom. 1997, 165, 339–347. (21) Johnson, R. S.; Krylov, D.; Walsh, K. A. J. Mass Spectrom. 1995, 30, 386– 387. (22) Jorgensen, T. J. D.; Bache, N.; Roepstorff, P.; Gardsvoll, H.; Ploug, M. Mol. Cell. Proteomics 2005, 4, 1910–1919. (23) Jorgensen, T. J. D.; Gardsvoll, H.; Ploug, M.; Roepstorff, P. J. Am. Chem. Soc. 2005, 127, 2785–2793. (24) McLafferty, F. W.; Guan, Z.; Haupts, U.; Wood, T. D.; Kelleher, N. L. J. Am. Chem. Soc. 1998, 120, 4732–4740. (25) Mueller, D. R.; Eckersley, M.; Richter, W. J. Org. Mass Spectrom. 1988, 23, 217–222. (26) Rand, K. D.; Jorgensen, T. J. D. Anal. Chem. 2007, 79, 8686–8693.

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To address this limitation, we have investigated the use of electron transfer dissociation (ETD)27 to accurately measure the hydrogen exchange of individual backbone amide hydrogens in a protein, i.e., an HX-ETD-MS experiment (Figure 1). In ETD, the backbone N-CR bond is cleaved with limited vibrational excitation when a multiply protonated peptide abstracts an electron from a radical molecular anion.27 With the use of a selectively labeled model peptide described recently,26 we carefully optimize the electrospray ion source conditions to obtain sufficient sensitivity, while maintaining negligible levels of gas-phase (1H/2H) scrambling before ETD.28 To demonstrate the general utility of the HX-ETD-MS experiment, we have subjected the amyloidogenic protein β2-microglobulin to isotopic exchange followed by peptic digestion and gas-phase fragmentation by ETD. β2-microglobulin is a 99 residue protein that has received considerable attention in recent years as this normally well-folded serum protein is deposited in vivo as amyloid fibrils in dialysis patients with chronic renal failure yielding the disorder dialysis-related amyloidosis (DRA).29-31 We evaluate the HX-ETD-MS approach by comparing the deuterium levels of individual residues of human β2-microglobulin calculated from mass analysis of ETD fragment ions of peptic peptides to previously published results from nuclear magnetic resonance (NMR) spectroscopy,32 which is currently the only established method to provide site-specific hydrogen-exchange data. EXPERIMENTAL SECTION Human β2-microglobulin was purchased from Sigma-Aldrich (St. Louis, MO). D2O (99.9 atom % D) was obtained from SigmaAldrich (St. Louis, MO). Model peptide P1 with the sequence HHHHHHIIKIIK was obtained from Genscript Corp. (Piscataway, NJ). The selective deuterium labeling of peptide P1 is described in detail elsewhere.26,33 All other chemicals were of the highest grade commercially available. Hydrogen Exchange of β2-Microglobulin. Equilibrium deuteration and proper folding of β2-microglobulin (0.45 or 0.9 mM) was achieved by incubation in deuterated buffer (20 mM phosphate pH 7.4, 100 mM NaCl, 99% D2O) for 6 h at 37 °C. β2-Microglobulin undergoes a global transient unfolding with a half-life of ∼70 min at 37 °C,34 which ensures that more than 97% of the protein molecules are fully deuterated and properly folded after 6 h. D/H exchange (exchange-out) was initiated by a 50-fold dilution of fully deuterated β2-microglobulin into (27) Syka, J. E. P.; Coon, J. J.; Schroeder, M. J.; Shabanowitz, J.; Hunt, D. F. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 9528–9533. (28) Zehl, M.; Rand, K. D.; Jensen, O. N.; Jorgensen, T. J. D. J. Am. Chem. Soc. 2008, 130, 17453–17459. (29) Eakin, C. M.; Berman, A. J.; Miranker, A. D. Nat. Struct. Mol. Biol. 2006, 13, 202–208. (30) Gejyo, F.; Yamada, T.; Odani, S.; Nakagawa, Y.; Arakawa, M.; Kunitomo, T.; Kataoka, H.; Suzuki, M.; Hirasawa, Y.; Shirahama, T.; Cohen, A. S.; Schmid, K. Biochem. Biophys. Res. Commun. 1985, 129, 701–706. (31) Jahn, T. R.; Parker, M. J.; Homans, S. W.; Radford, S. E. Nat. Struct. Mol. Biol. 2006, 13, 195–201. (32) Villanueva, J.; Hoshino, M.; Katou, H.; Kardos, J.; Hasegawa, K.; Naiki, H.; Goto, Y. Protein Sci. 2004, 13, 797–809. (33) Rand, K. D.; Adams, C. M.; Zubarev, R. A.; Jorgensen, T. J. D. J. Am. Chem. Soc. 2008, 130, 1341–1349. (34) Heegaard, N. H. H.; Jorgensen, T. J. D.; Rozlosnik, N.; Corlin, D. B.; Pedersen, J. S.; Tempesta, A. G.; Roepstorff, P.; Bauer, R.; Nissen, M. H. Biochemistry 2005, 44, 4397–4407.

Figure 1. Workflow of the HX-ETD-MS experiment. The amyloidogenic protein β2-microglobulin was subjected to isotopic exchange followed by peptic digestion and gas-phase fragmentation by ETD tandem mass spectrometry. Mass analysis of ETD fragment ions is performed at conditions of minimal vibrational excitation thus preventing gas-phase positional randomization (i.e., hydrogen scrambling) of the deuterium labeling pattern from solution.

protiated buffer at 25 °C (20 mM phosphate pH 7.4, 100 mM NaCl). Sample aliquots (50 µL) of the exchange-out reaction mixture (450 or 900 pmol) were removed at regular timeintervals (tex ) 0, 10 s, 30 s, 2 min, 8 min, 16 min, 40 min, and 4 h) and quenched by a 1:1 dilution in ice-cold quenching buffer (1.25 M tris(2-carboxyethyl)phosphine hydrochloride adjusted to pH 2.2 using NaOH) resulting in a final pH of 2.5 (uncorrected value). Quenched samples were snap frozen in liquid N2 and stored at -80 °C. Liquid Chromatography and ETD Tandem Mass Spectrometry. The experimental setup for HX-ETD-MS analyses consisted of a cooled high-pressure liquid chromatography (HPLC) system for online pepsin digestion and rapid desalting of protein samples connected to an ion trap mass spectrometer. The cooled HPLC system (Figure S1 in the Supporting Information) was configured largely as described previously35 and as detailed in the Supporting Information text. The HPLC system was coupled to a 3D quadrupole ion trap instrument equipped with an orthogonal ESI source and a separate chemical ionization source to produce the fluoranthene radical anions required for ETD (Agilent 6340 Ion Trap, Agilent Technologies, Santa Clara, CA). The ion trap was operated in positive ion mode (standard/ enhanced) with a scan range from 360 to 800 m/z in MS mode and 200-2000 m/z in MS/MS mode. Analyte ions were selected for ETD in the ion trap with an isolation width of 10 Th to prevent sideband excitation during precursor ion selection.28 Ion-ion (35) Jorgensen, T. J. D.; Gårdsvoll, H.; Danø, K.; Roepstorff, P.; Ploug, M. Biochemistry 2004, 43, 15044–15057.

reactions between analyte cations and fluoranthene radical anions were performed for 100 ms. The ion source and ion transfer optics of the ion trap were maintained at settings ensuring minimal collisional activation of doubly and triply charged peptide precursors prior to ETD in the trap: capillary exit voltage, 50 V; dry gas temperature, 100 °C; dc voltage on the first octopole, 6 V. Tuning of the declustering parameters of the mass spectrometer to conditions of negligible gas-phase hydrogen scrambling was performed as described previously employing a selectively labeled model peptide.26,28,33 Briefly, the selectively labeled model peptide P1 (sequence, HHHHHHIIKIIK) was continuously infused into the mass spectrometer for ETD via a dry ice cooled syringe mounted on an external sample pump. By measurement of the presence of deuterium in N-terminal ETD fragment ions (c2-c5) of the model peptide as a gauge of the occurrence of gas-phase hydrogen scrambling,28 declustering conditions were defined that allowed maximal intensity of ETD fragment ions while maintaining negligible levels of scrambling. To verify that these declustering conditions provided negligible hydrogen scrambling in protonated peptides during HX-ETD-MS analyses, the experiment was repeated with infusion of the selectively labeled model peptide into the mass spectrometer via the cooled HPLC system (Figure S2 in the Supporting Information). A complete list of ion trap instrument settings used during HX-ETD-MS analyses is provided in Table S2 in the Supporting Information. It should be noted that we have previously shown that the level of hydrogen scrambling is negligible for six different model peptides using similar instrumental settings.28 Experiments conducted at Analytical Chemistry, Vol. 81, No. 14, July 15, 2009

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harsh declustering conditions optimized for maximum precursor ion transmission while causing high levels of hydrogen scrambling (Figure S2 in the Supporting Information) were performed by increasing the following parameters: capillary exit voltage, 250 V; dc voltage on the first octopole, 15 V. On average, the sensitivity of the HX-ETD-MS analyses on β2-microglobulin dropped about 2-3-fold upon changing the declustering parameters from conditions of maximal ETD fragment ion signal to the defined conditions for negligible gas-phase hydrogen scrambling used throughout this study (data not shown). Optimizing the Sequence Coverage Provided by ETD Fragment Ions. The ion trap mass spectrometer was operated in automatic MS/MS (data-dependent acquisition) mode, and optimized exclusion mass ranges were set to prevent selection of highly abundant cluster ions (see also Table S2 in the Supporting Information). In this manner, the deuterium levels of 42 out of 93 amide groups (45%) of β2-microglobulin could be measured in a single data-acquisition run (we are only considering 93 backbone amide bonds because proline residues do not have an amide hydrogen). To further optimize the sequence coverage of fragment ions of β2microglobulin, analyses of replicate samples were performed in manual MS/MS mode enabling selection of particular lower abundant peptic peptides. In separate experiments, analyses of replicate samples were performed with the addition of m-nitrobenzyl alcohol (m-NBA) to the HPLC solvents. The presence of m-NBA significantly increases the charge state of peptic peptides upon electrospray ionization. This effect was significantly more pronounced than observed previously for tryptic peptides,36 likely due to the less specific cleavage pattern of pepsin. The presence of 0.05% (w/v) m-NBA significantly reduced the abundance of doubly protonated peptic peptides while dramatically increasing the abundance of triply protonated peptides. The latter typically produced more abundant fragment ions and a more complete ion series upon ETD in the ion trap, thus facilitating improved localization of the incorporated deuterium (Figure S3 in the Supporting Information). Notably, while higher concentrations of m-NBA in mobile HPLC phases further depleted the population of doubly charged peptic peptides, they favored the formation of peptides with charge states greater than +3 (data not shown). These ions often yield good ETD spectra, but their high charge state gives them an elevated kinetic energy during the declustering process in the ion source and they are thus potentially more prone to undergo hydrogen scrambling compared to their triply charged counterparts (due to increased vibrational excitation upon collisions with residual gas molecules along the ion path).28,33 Only peptic peptide precursor ions of +2 and +3 charge states were selected for ETD in the present study. With the employment of the abovementioned procedures for optimizing sequence coverage at single-residue resolution, the deuterium content of 60 out of 93 individual amide groups (65%) of β2-microglobulin was monitored in the combined HX-ETD-MS analyses. (36) Kjeldsen, F.; Giessing, A. M. B.; Ingrell, C. R.; Jensen, O. N. Anal. Chem. 2007, 79, 9243–9252. (37) Bai, Y.; Milne, J. S.; Mayne, L.; Englander, S. W. Proteins 1993, 17, 75– 86.

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RESULTS AND DISCUSSION Measuring the Solution Isotopic Exchange of β2-Microglobulin by ETD. Hydrogen exchange of β2-microglobulin was initiated by diluting fully deuterated β2-microglobulin into 1H2O phosphate buffer. In such an “exchange-out” experiment, deuteriums initially bound to heteroatoms in β2-microglobulin exchange with hydrogens from surrounding solvent such that highly exposed sites exchange rapidly while deuterium is retained at slow-exchanging sites of the protein that are shielded from solvent or engaged in hydrogen bonding (see ref 1 for a review). Notably, while the described HX-ETD-MS method is equally compatible with an “exchange-in” experiment (i.e., dilution of protiated protein into deuterated buffer), data from exchange-out experiments on β2-microglobulin enabled straightforward comparison with NMR reference values which were available only at longer exchange times (>40 min). Isotopic exchange was quenched after various periods of incubation by acidification (pH 2.5) and cooling (0 °C). The labeled protein was digested online by immobilized pepsin and the resulting peptic peptides were separated by reversed-phase chromatography and analyzed by ETD tandem mass spectrometry (see also the Supporting Information text and Figure S1 in the Supporting Information). Deuterium located in side-chains, the amino/carboxy termini, and the N-terminal amide of peptic peptides are completely lost due to fast exchange37 with the 1H2O HPLC solvents during the chromatographic separation. This enables measurement of deuterium exclusively at amide positions of the peptic peptides. The mass spectrometer continuously acquired mass spectra of the eluting peptides and automatically selected the most abundant doubly or triply charged ions for ETD fragmentation (Figure 2a). In contrast to standard HPLC-MS/MS conditions used, e.g., for proteomicstype experiments, instrumental parameters were optimized to ensure a low degree of vibrational excitation of the peptide ions (see Experimental Section and Figure S2 in the Supporting Information), and to yield sufficiently good ion statistics to allow determination of the average mass of the fragment ions (i.e., 10-15 MS/MS scans were accumulated per spectrum). Pepsin proteolysis of β2-microglobulin produced 20 peptides covering 97% of the amino acid sequence. These peptides enabled the localization of retained amide deuterons in β2-microglobulin to short segments typically 6-10 residues in length. To increase the resolution further, peptide ions were fragmented in the gasphase by ETD to yield c- and z-type fragment ions corresponding to cleavage of the backbone N-CR bond. The deuterium content of a single backbone amide was then determined by measuring the difference in the average mass of sequential fragment ions (e.g., cn/zn and cn+1/zn+1) from a labeled peptic peptide and comparing it to the corresponding mass difference obtained from the unlabeled peptide. Note that, since ETD fragment ions originate from backbone N-CR bond cleavage, the mass difference of the cn and the cn+1 ion actually reports the deuterium content of the amide bond linking residues n + 1 and n + 2 in the peptide. In this way, the residue-specific deuterium content of 65% of the backbone amides of β2microglobulin was obtained (Figure 2b). Deuterium labeling of a further 20% of the backbone amides of β2-microglobulin were resolved to short segments of 2-6 residues in length by

Figure 2. HX-ETD-MS analysis of the amyloidogenic protein β2-microglobulin. (a) Chromatographic separation and MS analysis of deuterium labeled peptic peptides from β2-microglobulin. The eluting peptides were automatically selected for ETD (MS/MS). ETD fragment ions resulting from backbone cleavage at the N-CR bond (c- and z-ions) are shown in red. (b) Sequence coverage of β2-microglobulin obtained from peptic peptides and their ETD fragment ions. Peptides are shown as horizontal bars where double lines denote ETD cleavage sites producing abundant fragment ions suitable for measuring the deuterium levels of individual residues (bounded by red squares). The charge states of peptide precursor ions are indicated by plus signs, and secondary structure elements of β2-microglobulin are shown above the sequence.

combined mass analysis of both ETD fragment ions and intact peptic peptides. An example of HX-ETD-MS data is shown in Figure 3, where the exact location of retained deuteriums after 40 min exchange-out in a section of β2-microglobulin (residues 40-54) is readily apparent from the masses of sequential fragment ions of the peptic peptide. In a recent NMR study,32 the amide hydrogens of E44 and I46 of β2-microglobulin were identified as slow-exchanging. The fragment ions of peptide 40-54 clearly show that the deuterium retained in this peptide is localized exclusively to E44 and I46 in excellent agreement with the NMR data (see also Supporting Information text). Comparison of MS and NMR Measurements on β2Microglobulin. All ETD fragment ions of peptic peptides were analyzed to extract site-specific deuterium levels of individual residues of β2-microglobulin. The complete list of measured deuterium contents of individual residues of β2-microglobulin after 40 min exchange-out (DETD) are given in Table S1 in the Supporting Information. In order to be able to evaluate our results against the data obtained from NMR spectroscopy, the measured site-specific deuterium levels were corrected for the unintentional but inherent loss of deuterium during HX-ETD-MS analysis by normalizing them to the measured deuterium levels of fully deuterated β2-microglobulin (back-exchange correction,38 Supporting Information text). A comparison of backbone amide deuterium levels of all residues of β2-microglobulin measured by ETD and NMR spectroscopy32 is shown in Figure 4. The (38) Zhang, Z.; Smith, D. L. Protein Sci. 1993, 2, 522–531.

back-exchange corrected deuterium levels as measured by ETD (DETD*) correlate very well with expected deuterium levels obtained from NMR measurements (DNMR). Minor differences exist between DETD* and DNMR for a few residues, and a subpopulation of these residues display small negative deuterium contents (Table S1 in the Supporting Information). Such deviations are most likely due to inaccurate measurement of the relative abundances of the individual isotopes or overlap from interfering ions in the mass spectrum. Regardless, the standard error of the mean (sem) from replicate HX-ETD-MS experiments never exceeded 0.2 even for deviant residues (with an average sem of 0.04 for the uncorrected DETD and 0.07 for the backexchange corrected DETD* values, Table S1 in the Supporting Information). Thus, a clear correlation exists between our MS result and NMR reference data. This further demonstrates that the level of hydrogen scrambling for all β2-microglobulin peptides is negligible using the present instrumental settings. If scrambling was prevalent then it would have erased the specific labeling obtained in solution. For certain residues, ETD and NMR data proved to be complementary. For instance, quantitative data for the slow exchanging residue Q8 could not be obtained in NMR due to signal overlap with Y66, whereas the deuterium content of this residue could be accurately measured with ETD. On the other hand, we were not able to perform correction for back-exchange of the slow exchanging residue L87, which was fully characterized by NMR, due to poor ion statistics in the fully labeled control measurements. Analytical Chemistry, Vol. 81, No. 14, July 15, 2009

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Figure 3. Measuring the deuterium content of individual backbone amides of β2-microglobulin by ETD: (a) ETD mass spectra of the triply charged β2-microglobulin peptide LKNGERIEKVEHSDL (residues 40-54) from an unlabeled (upper panels) and a labeled sample obtained after 40 min isotopic exchange (lower panels). The isotopic patterns of the fragment ions c3-c6 are displayed. (b) Bar chart showing the deuterium level of all c ions. The deuterium level of the intact peptide precursor ion is indicated by a dotted line. The protected residues E44 and I46 are colored in red in the sequence. Note that the amide hydrogen of residue n is contained in the cn-1 fragment ion. (c) Crystal structure of β2microglobulin (PDB: 1LDS) with the sequence LKNGERIEKVEHSDL highlighted in blue. Residues E44 and I46, which were identified as slowexchanging by recent NMR32 and present ETD data, are shown as red sticks.

Figure 4. Deuterium levels of individual backbone amides in β2-microglobulin measured by NMR (open black circles) and ETD (filled red circles). The deuterium levels obtained from ETD experiments were determined by the difference in deuterium content of adjacent c- or z-type ETD fragment ions from peptic peptides of labeled β2-microglobulin after 40 min exchange-out and corrected for back-exchange. Reference deuterium levels were obtained from exchange-rate constants determined by NMR.32 The standard error of the mean (sem) of ETD-measured deuterium levels from at least two independent replicate experiments is indicated. Residues with small negative deuterium levels due to the error of mass measurement were not corrected for back-exchange. 5582

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CONCLUSIONS Our results show that gas-phase dissociation by ETD of proteolytically generated peptides can yield reliable information on the labeling of individual residues of a protein in solution. This is achieved because gas-phase hydrogen scrambling is negligible during the present liquid chromatography-tandem mass spectrometric method. The potentials of the ion source and the ion transfer optics of the mass spectrometer used in this study (Table S2 in the Supporting Information) were specifically tuned to ensure minimal vibrational excitation of deuterium labeled peptide ions thus minimizing gas-phase hydrogen scrambling. Instrumental parameters can, however, not be transferred directly from one instrument type to another. Hence, for other ETD mass spectrometers, the settings should be optimized individually to avoid gas-phase randomization of peptide deuterium labeling patterns. As described in the Experimental Section and in greater detail elsewhere,26,28 this can be conveniently done by use of a selectively labeled model peptide as a gauge of gas-phase hydrogen scrambling during ETD (Figure S2 in the Supporting Information). Using pepsin proteolysis and ETD tandem mass spectrometry, we deconvoluted the deuterium labeling of β2-microglobulin and localized deuteriums to 65% of individual backbone amides in the protein while resolving the labeling pattern of an additional 20% to segments of 2-6 backbone amides. Thus for smaller proteins such as β2-microglobulin, the hydrogen exchange data provided by an HX-ETD-MS experiment can complement and extend NMR measurements that typically offer detection of exchange of some 60-80% of individual backbone amides (depending on spectral overlap). Unlike NMR, however, the use of mass spectrometry to monitor protein hydrogen exchange allows the study of large proteins and protein complexes (>40 kDa), low sample concentration and consumption, versatile labeling conditions (buffer, pH, temperature, presence of cofactors/ligands), and the ability to monitor individual conformational states of a protein.4,5,13 The present HX-ETDMS approach should therefore enable a delineation of protein dynamics and function in the many areas of molecular biology hitherto not accessible to high-resolution hydrogen exchange studies. In addition, the HX-ETD-MS experiment readily offers the possibility of performing short labeling times. This enables the characterization of amide hydrogens that exchange on a time scale not routinely resolvable by standard NMR experiments (illustrated for V9 and A15 in Figure S4 in the Supporting Information). Of note, a recent combination of rapid-mixing quench-flow techniques and NMR detection has provided measurements on fast exchanging amide hydrogens.39 With the use of high-resolution mass spectrometers, electronbased fragmentation methods should also be applicable for direct fragmentation of labeled proteins (i.e., without preceding proteolytic cleavage) to obtain single-residue deuteration levels. In a recent study, direct fragmentation by electron capture dissociation (ECD) enabled the assignment of incorporated deuteriums to individual residues of ubiquitin.40 Such “top-down” approaches require, however, relatively long spectral acquisition times of (39) 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. (40) Pan, J.; Han, J.; Borchers, C. H.; Konermann, L. J. Am. Chem. Soc. 2008, 130, 11574–11575.

continuously infused protein samples. Therefore, these approaches do not appear to be compatible with a HPLC-based workflow where eluting chromatographic peaks limit the acquisition time. Furthermore, top-down ECD/ETD is likely to only yield information on site-specific deuterium levels of small proteins. Larger proteins typically fail to yield sufficient fragment ions upon ETD or ECD without concomitant vibrational activation, which in turn causes hydrogen scrambling.41,42 The “bottom-up” approach used in the present study does not have similar limitations regarding protein mass. However, as in the classical HX-MS experiment, the chromatographic separation of complex peptic peptide mixtures during HX-ETD-MS analysis of very large proteins would be limited by the need for rapid chromatography to minimize loss of isotopic labeling (back-exchange). Specifically, the ability to record ETD spectra of individual peptides that coelute into the mass spectrometer depends on the times required during each acquisition to accumulate sufficient ion statistics to accurately determine the average masses of ETD fragment ions (10-15 MS/ MS scans per ETD spectrum were acquired in this study). Thus, obtaining a similar sequence coverage of individual sites from ETD fragment ions as demonstrated presently for β2-microglobulin would be a considerable challenge for much larger (>60-70 kDa) proteins. However, of particular utility for such large proteins, the HX-ETD-MS experiment provides the option of manually selecting only key peptic peptides for optimized ETD fragmentation, thus extracting detailed hydrogen exchange information for a specific protein segment of particular functional relevance at single-residue resolution, while resolving the remaining exchanging amides at the level of peptic peptides as in a classical HX-MS experiment. Improvements of the chromatographic separation, e.g., by applying ultraperformance liquid chromatography (UPLC)-type chromatographic systems as described recently,43 and mass spectrometric resolution and sensitivity, e.g., by ETD-capable linear ion trap/orbitrap hybrid instruments,44 will further improve the HX-ETD-MS analysis of complex peptic peptide mixtures from larger proteins and protein complexes. In conclusion, we propose that the presented methodology will be a valuable and widely applicable tool for high-resolution monitoring of the conformational dynamics of proteins and protein complexes in solution. ACKNOWLEDGMENT We gratefully acknowledge financial support by The Danish Natural Science Research Council (FNU Grant No. 272-07-0276 and Grant No. 272-06-0493, K.D.R and T.J.D.J., respectively), the EU (M.Z.; Grant FP6, MRTN-CT-2005-019566), The Lundbeck Foundation (O.N.J), and The Carlsberg Foundation (T.J.D.J). We thank Dr. Anders Giessing and Dr. John Engen for valuable discussions and Dr. Hoshino and Dr. Goto for providing the NMR reference data. NOTE ADDED IN PROOF Very recently Abzalimov et al. have reported a low degree of gas phase hydrogen scrambling upon fragmentation of a multiply (41) Han, X.; Jin, M.; Breuker, K.; McLafferty, F. W. Science 2006, 314, 109– 112. (42) Horn, D. M.; Ge, Y.; McLafferty, F. W. Anal. Chem. 2000, 72, 4778–4784. (43) Wales, T. E.; Fadgen, K. E.; Gerhardt, G. C.; Engen, J. R. Anal. Chem. 2008, 80, 6815–6820. (44) McAlister, G. C.; Phanstiel, D.; Good, D. M.; Berggren, W. T.; Coon, J. J. Anal. Chem. 2007, 79, 3525–3534.

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protonated protein by ETD. Single-residue resolution was, however, not obtained with this top-down fragmentation approach.45 SUPPORTING INFORMATION AVAILABLE Tables S1 and S2, Figures S1–S4, and descriptions of the HPLC setup for pepsin digestion, rapid desalting, and chromatographic (45) Abzalimov, R. R.; Kaplan, D. A.; Easterling, M. L.; Kaltashov, I. A. J. Am. Soc. Mass Spectrom. 2009, in press, DOI: 10.1016/j.jasms.2009.04.006.

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separation and the procedure for analysis of HX-ETD-MS data. This material is available free of charge via the Internet at http://pubs.acs.org.

Received for review April 20, 2009. Accepted May 22, 2009. AC9008447