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Anal. Chem. 2006, 78, 1613-1619

Pulsed Hydrogen/Deuterium Exchange MS/MS for Studying the Relationship between Noncovalent Protein Complexes in Solution and in the Gas Phase after Electrospray Ionization Belal M. Hossain and Lars Konermann*

Department of Chemistry, The University of Western Ontario, London, Ontario N6A 5B7, Canada

Electrospray ionization mass spectrometry (ESI-MS) has become a standard method for monitoring noncovalent protein-protein interactions. Studies employing this approach tend to operate on the premise that the ionic species observed in the mass spectrum directly reflect the corresponding solution-phase protein quaternary structures. However, dissociation or clustering events taking place during ESI may lead to disparities between the ions observed in the mass spectrum and the protein binding state in bulk solution. Recognizing the occurrence of dissociation or clustering artifacts is not straightforward, leading to possible ambiguities in the interpretation of ESI-MS data. This work employs on-line pulsed hydrogendeuterium exchange (HDX) for probing the origin of various species in the ESI mass spectrum of hemoglobin. In addition to the canonical hemoglobin tetramer, ESIMS reveals the presence of monomers, dimers, hexamers, and octamers. Tandem mass spectrometry (MS/MS) is used for extracting HDX levels in a subunit-specific manner. Dimeric species exhibit exchange levels that are significantly above those of the tetramer. Monomeric hemoglobin subunits are labeled to an even greater extent. This HDX pattern implies that monomers and dimers do not represent dissociation artifacts generated during ESI. Instead, they are derived from preexisting solution-phase structures. In contrast, hexamers and octamers exhibit HDX levels that resemble those of the tetramer, thus identifying these larger species as nonspecific clustering artifacts. Overall, it appears that the pulsed HDX MS/MS approach introduced in this work represents a widely applicable tool for deciphering the relationship between ESI mass spectra and protein quaternary structures in solution. Numerous research efforts are directed toward mapping the noncovalent interactions of biological macromolecules with their binding partners. Of particular interest are protein-protein and protein-ligand interactions. A comprehensive knowledge of all molecular entities that can interact with any given protein is a key requirement for understanding biochemical regulation and * To whom correspondence should be addressed. E-mail: [email protected]; http://publish.uwo.ca/∼konerman/. 10.1021/ac051687e CCC: $33.50 Published on Web 01/26/2006

© 2006 American Chemical Society

signaling processes. Studies in this area can provide fundamental insights into disease mechanisms, resulting in new targets for drug discovery.1 The methods that are currently being used for detecting the interactions of proteins with their binding partners include frontal affinity chromatography,2 surface plasmon resonance assays,3 various diffusion-based techniques,4,5 hydrogendeuterium exchange approaches,6 and large-scale tandem-affinity purification strategies,7 as well as screening assays that use yeast two-hybrid methods8 and protein microarray chips.9 A major drawback of most of these techniques is that they are relatively labor intensive and time-consuming. Over the past decade, electrospray mass spectrometry (ESIMS) has emerged as an important tool for the characterization of protein-protein complexes. With this conceptually simple approach, noncovalent interactions can be studied in straightforward “mix-and-measure” experiments.10-17 The softness of the ESI process allows the transfer of intact molecular assemblies into the gas phase, such that protein complexes are directly observable as ions in the mass spectrum. The stoichiometry of a complex can be determined from its mass,18 and it is often possible to (1) Schermann, S. M.; Simmons, D. A.; Konermann, L. Exp. Rev. Proteomics 2005, 2, 475-485. (2) Schriemer, D. C. Anal. Chem. 2004, 76, 441A-448A. (3) McDonnell, J. M. Curr. Opin. Chem. Biol. 2001, 5, 572-577. (4) Gradl, G.; Guenther, R.; Sterrer, S. BioMethods 1999, 10, 331-351. (5) Clark, S. M.; Konermann, L. Anal. Chem. 2004, 76, 1257-1263. (6) Powell, K. D.; Ghaemmaghami, S.; Wang, M. Z.; Ma, L.; Oas, T. G.; Fitzgerald, M. C. J. Am. Chem. Soc. 2002, 124, 10256-10257. (7) Gavin, A.-C.; Bosche, M.; Krause, R.; et al. Nature 2002, 415, 141-147. (8) Titz, B.; Schlesner, M.; Uetz, P. Exp. Rev. Proteomics 2004, 1, 111-121. (9) Zhu, H.; Bilgin, M.; Bangham, R.; Hall, D.; Casamayor, A.; Bertone, P.; Lan, N.; Jansen, R.; Bidlingmaier, S.; Houfek, T.; Mitchell, T.; Miller, P.; Dean, R. A.; Gerstein, M.; Snyder, M. Science 2001, 293, 2101. (10) Katta, V.; Chait, B. T. J. Am. Chem. Soc. 1991, 113, 8534-8535. (11) Ganem, B.; Henion, J. D. Bioorg. Med. Chem. 2003, 11, 311-314. (12) Loo, J. A. Int. J. Mass Spectrom. 2000, 200, 175-186. (13) Daniel, J. M.; Friess, S. D.; Rajagopalan, S.; Wendt, S.; Zenobi, R. Int. J. Mass Spectrom. 2002, 216, 1-27. (14) Rostom, A. A.; Fucini, P.; Benjamin, D. R.; Juenemann, R.; Nierhaus, K. H.; Hartl, F. U.; Dobson, C. M.; Robinson, C. V. Proc. Natl. Acad. Sci. U.S.A. 2000, 97, 5185-5190. (15) Heck, A. J. R.; Van den Heuvel, R. H. H. Mass Spectrom. Rev. 2004, 23, 368-389. (16) Kaltashov, I. A.; Eyles, S. J. Mass Spectrometry in Biophysics; John Wiley and Sons: Hoboken, NJ, 2005. (17) Wigger, M.; Eyler, J. R.; Benner, S. A.; Li, W.; Marshall, A. G. J. Am. Soc. Mass Spectrom. 2002, 13, 1162-1169. (18) Zhang, Z.; Krutchinsky, A.; Endicott, S.; Realini, C.; Rechsteiner, M.; Standing, K. G. Biochemistry 1999, 38, 5651-5658.

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determine binding affinities from the ion abundance ratio of free to bound protein.13,19-23 Although ESI-MS represents one of the fastest and simplest approaches for monitoring protein interactions, there are a number of considerations that have to be taken into account when this technique is employed. Despite the softness of the overall process, it has to be recognized that ESI results in dramatic changes in the physical environment of the analyte. Potentially, this can lead to situations where the ions observed in the mass spectrum no longer reflect the solution-phase binding state of a protein.24 Charge-balancing redox reactions inside the ESI emitter can induce pH changes that may cause the disruption of protein complexes.21,25,26 Collision-induced or thermal dissociation of noncovalent interactions can occur in the declustering region of the mass spectrometer. Another important aspect is that the overall charge of gas-phase proteins after ESI can be considerably different from what it was in solution.27 Moreover, electrostatic interactions will undergo drastic alterations due to the 80-fold change in the dielectric constant of the surroundings.28 These and other environmental changes may cause a destabilization of noncovalent assemblies to a point where they become almost undetectable.17,29-31 Conversely, ESI can also give rise to clustering events within the rapidly shrinking solvent droplets that are produced at the emitter tip.32,33 Nonspecific adducts generated in this way represent artificial ionic complexes that do not correspond to preexisting solution-phase assemblies.34,35 Taken together, the above considerations imply that the binding state of a protein in solution cannot always be unambiguously deduced from its mass spectrum. ESI-MS studies on noncovalent protein complexes frequently report the observation of quaternary structures that are consistent with solution-phase or crystallographic measurements. However, the detection of additional species that are either larger or smaller than those anticipated is a very common occurrence.15,36,37 These unexpected complexes could represent genuine assemblies that are derived from solution-phase complexes, or they could be artifacts of the ESI process. Regrettably, (19) Jorgensen, T. J. D.; Roepstorff, P.; Heck, A. J. R. Anal. Chem. 1998, 70, 4427-4432. (20) Hagan, N.; Fabris, D. Biochemistry 2003, 42, 10736-10745. (21) Wang, W.; Kitova, E. N.; Klassen, J. S. Anal. Chem. 2003, 75, 4945-4955. (22) Wendt, S.; McCombie, G.; Daniel, J.; Kienhofer, A.; Hilvert, D.; Zenobi, R. J. Am. Soc. Mass Spectrom. 2003, 14, 1470-1476. (23) Wortmann, A.; Rossi, F.; Lelais, G.; Zenobi, R. J. Mass Spectrom. 2005, 40, 777-784. (24) Peschke, M.; Verkerk, U. H.; Kebarle, P. J. Am. Soc. Mass Spectrom. 2004, 15, 1424-1434. (25) Van Berkel, G. J.; Zhou, F.; Aronson, J. T. Int. J. Mass Spectrom. Ion Processes 1997, 162, 55-67. (26) Konermann, L.; Silva, E. A.; Sogbein, O. F. Anal. Chem. 2001, 73, 48364844. (27) Wang, G.; Cole, R. B. In Electrospray Ionization Mass Spectroscopy; Cole, R. B., Ed.; John Wiley & Sons: New York, 1997; pp 137-174. (28) Schmidt, A.; Karas, M. J. Am. Soc. Mass Spectrom. 2001, 12, 1092-1098. (29) Robinson, C. V.; Chung, E. W.; Kragelund, B. B.; Knudsen, J.; Aplin, R. T.; Poulsen, F. M.; Dobson, C. M. J. Am. Chem. Soc. 1996, 118, 8646-8653. (30) Clark, S. M.; Konermann, L. Anal. Chem. 2004, 76, 7077-7083. (31) Mauk, M. R.; Mauk, A. G.; Chen, Y.-L.; Douglas, D. J. J. Am. Soc. Mass Spectrom. 2002, 13, 59-71. (32) Juraschek, R.; Dulcks, T.; Karas, M. J. Am. Soc. Mass Spectrom. 1999, 10, 300-308. (33) Bakhoum, S. F. W.; Agnes, G. R. Anal. Chem. 2005, 77, 3189-3197. (34) Cunniff, J. B.; Vouros, P. J. Am. Soc. Mass Spectrom. 1995, 6, 437-447. (35) Wang, W.; Kitova, E. N.; Klassen, J. S. Anal. Chem. 2005, 77, 3060-3071. (36) van Berkel, W. J. H.; van den Heuvel, R. H. H.; Versluis, C.; Heck, A. J. R. Protein Sci. 2000, 9, 435-439. (37) Kaltashov, I. A.; Mohimen, A. Anal. Chem. 2005, 77, 5370-5379.

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the origin of these species often remains unresolved. Concentration-dependent measurements are of limited use for addressing this issue. For example, the formation of complexes that result from clustering artifacts will be favored at high protein concentrations, just as elevated concentrations will favor the formation of large complexes in a solution-phase binding equilibrium. Our group has recently proposed a method for delineating the relationship between ESI mass spectrum and protein binding state in solution using an on-line pulsed hydrogen-deuterium exchange (HDX) approach.38 The protein is exposed to D2O under rapid exchange conditions for a short amount of time immediately preceding ESI. HDX can occur for hydrogens in N-H, O-H, and S-H bonds, including those in amide groups, in side chains, and on the chain termini. However, extensive HDX will only affect sites that are solvent-exposed and not involved in hydrogen bonds. Binding to another moiety will decrease the HDX level of a protein as a result of several factors, namely, the steric protection of exchangeable sites, the formation of intermolecular hydrogen bonds, and the fact that binding usually causes proteins to adopt more tightly folded solution-phase conformations.6,39-42 Consequently, in a case where ESI-MS directly reflects the solutionphase binding state of a protein, each individual quaternary structure in the spectrum should show a unique HDX signature. In contrast, proteins that adopt a single binding state in the aqueous phase will all experience the same level of labeling. Therefore, artifactual dissociation or clustering products will show labeling characteristics that are indistinguishable. In a preliminary report, we demonstrated the feasibility of this approach for studying heme binding to a simple monomeric protein, myoglobin.38 The current study demonstrates how the described pulselabeling approach can be combined with tandem mass spectrometry (MS/MS) for studies on larger protein complexes. Bovine hemoglobin represents an interesting system for investigating the relationship between quaternary structure in solution and in the gas phase. The canonical tetrameric X-ray structure of this protein comprises two pairs of heme-containing R and β subunits in a tetrahedral arrangement.43 This complex will be denoted as (Rhβh)2, where the superscript indicates the presence of a heme group on the respective subunits. Previous studies on hemoglobin, employing suitably gentle interface conditions and a nativelike solvent environment, have confirmed that (Rhβh)2 tetramers are easily detectable by ESI-MS. In addition, however, the hemoglobin spectra show a range of ionic species that correspond to other quaternary structures.44,45 Dimers of the composition Rhβh are observed along with monomeric Rh ions. Monomeric β subunits are observed exclusively in their apo form. Close examination of the latter reveals a mass increase of 32 Da, which has been attributed to the oxidation of two methionine side chains.45 This species will be denoted as βoxa, with the superscript (a ) apo), indicating the lack of heme. Oxidized apo-β subunits also appear (38) Hossain, B. M.; Simmons, D. A.; Konermann, L. Can. J. Chem. In press. (39) Zhu, M. M.; Rempel, D. L.; Du, Z.; Gross, M. L. J. Am. Chem. Soc. 2003, 125, 5252-5253. (40) Kaltashov, I. A.; Eyles, S. J. Mass Spectrom. Rev. 2002, 21, 37-71. (41) Konermann, L.; Simmons, D. A. Mass Spectrom. Rev. 2003, 22, 1-26. (42) Krishna, M. M. G.; Hoang, L.; Lin, Y.; Englander, S. W. Methods 2004, 34, 51-64. (43) Zubay, G. Biochemistry, 4 ed.; Wm. C. Brown: Dubuque, IA, 1998. (44) Griffith, W. P.; Kaltashov, I. A. Biochemistry 2003, 42, 10024-10033. (45) Simmons, D. A.; Wilson, D. J.; Lajoie, G. A.; Doherty-Kirby, A.; Konermann, L. Biochemistry 2004, 43, 14792-14801.

Figure 1. Experimental approach used to delineate the relationship between solution-phase hemoglobin quaternary structure and protein binding states observed in ESI-MS. In this simplistic diagram, hemoglobin subunits are indicated as squares; heme groups have been omitted for clarity. Bold lines indicate regions that have undergone hydrogen-deuterium exchange (HDX). (A) Hypothetical scenario where ionic species generated by ESI directly reflect the corresponding solution-phase quaternary structures. (B) Hypothetical scenario where the protein in solution exists exclusively as tetramer; monomers, dimers, octamers, etc., are artifacts of the ESI process. At the bottom of both panels it is illustrated how MS/MS analysis reveals the HDX levels of individual subunits. For (A), all R subunits show distinct labeling levels that depend on the type of precursor ion used for the analysis. The same is true for the β subunits. In (B), all R subunits show the same labeling level, and all β subunits show the same labeling level. Note that scenarios (A) and (B) would be indistinguishable without isotope labeling.

in heme-deficient dimers of the composition Rhβoxa. Additionally, ESI-MS reveals the presence of protein assemblies that are larger than the expected tetramer, namely, hexamers (Rhβh)3, octamers (Rhβh)4, and possibly even larger complexes that are not easily assigned.44,45 There appear to be no previous studies to support the existence of these larger quaternary structures in solution. What is the origin of the nontetrameric species in the hemoglobin mass spectrum? It will be helpful to consider two extreme cases: In scenario A, all the observed ions directly reflect the presence of the corresponding protein binding states in solution. Partial methionine oxidation of β subunits occurs during isolation or storage, as it does for other proteins.46-49 In scenario B, the protein in solution exists exclusively as tetramer. All other ionic species are formed as artifacts of the ESI process; monomers as well as dimers represent dissociation products, and hexamers/ octamers are the result of nonspecific clustering. The oxidation of β subunits is caused by hydroxyl radicals generated in the ion source.50,51 Based on the appearance of a hemoglobin ESI mass spectrum alone, it is impossible to differentiate these two scenarios or other possibilities that might conceivably exist. However, when incorporating a pulsed HDX step into the experimental protocol, the origin of the protein ions should, in principle, be apparent from their isotope exchange levels. One problem with this idea is the (46) Hoshi, T.; Heinemann, S. H. J. Physiol. 2001, 531, 1-11. (47) Kim, Y. H.; Berry, A. H.; Spencer, D. S.; Stites, W. E. Protein Eng. 2001, 14, 343-347. (48) Vogt, W. Free Radical Biol. Med. 1995, 18, 93-105. (49) Creighton, T. E. Proteins; W. H. Freeman & Co: New York, 1993. (50) Wong, J. W. H.; Maleknia, S. D.; Downard, K. M. Anal. Chem. 2003, 75, 1557-1563. (51) Wong, J. W. H.; Maleknia, S. D.; Downard, K. M. J. Am. Soc. Mass Spectrom. 2005, 16, 225-233.

commonly observed incomplete desolvation of multisubunit protein ions,52,53 which causes considerable mass shifts and peak broadening that will obscure any HDX patterns. To circumvent this problem, the current work employs a MS/MS approach. In addition to providing greatly improved peak shapes, MS/MS allows the readout of HDX levels in a subunit-specific manner. The experimental strategy is summarized in Figure 1. For scenario A, the protein subunits should exhibit exchange levels that are significantly different for the various precursor ions. The highest HDX levels will be observed for monomeric species and the lowest levels for large protein complexes (Figure 1A). In contrast, for scenario B, the HDX levels of the subunits will be uniform for all the quaternary structures observed in the spectrum (Figure 1B). The data obtained by using this approach strongly suggest that monomers, dimers, and tetramers in the hemoglobin ESI mass spectrum are derived from the corresponding solution-phase species, whereas larger complexes are clustering artifacts. EXPERIMENTAL SECTION Chemicals. Ammonium-d4 deuterioxide, acetic acid-OD (Isotec, Inc., Miamisburg, OH), deuterium oxide (Cambridge Isotope Laboratories, Andover, MA), ammonium hydroxide (Fisher Scientific, Nepean, ON, Canada), and ammonium acetate (Fluka, Buchs, Switzerland) were used without further purification. Bovine ferrihemoglobin (Sigma, St. Louis, MO) was dialyzed over a period of 5 days at 0 °C against decreasing concentrations (1 M, 250 mM, and 10 mM) of ammonium acetate in water prior to ESIMS. This procedure results in significantly improved peak shapes, (52) Green, B. N.; Vinogradov, S. N. J. Am. Soc. Mass Spectrom. 2004, 15, 2227. (53) Loo, J. A. In The Encyclopedia of Mass Spectrometry; Gross, M. L., Caprioli, R. M., Eds.; Elsevier: Amsterdam, 2005; Vol. 2, pp 289-299.

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Figure 2. ESI mass spectrum of 100 µM hemoglobin recorded after 53 ms of isotope labeling in 80% D2O at pD 8.5. The subunit composition of the observed ions is denoted as explained in the text; e.g., (Rhβh)2 corresponds to intact hemoglobin heterodimers with one heme per subunit. Numbers indicate positive ion charge states.

which allowed unambiguous peak assignments for hexameric and octameric protein assemblies. In addition, it facilitated the precursor ion selection in MS/MS. When compared to spectra that had been recorded on less extensively dialyzed samples, the data in this work showed somewhat elevated relative ion intensities of monomeric and dimeric species (compare Figure 2 of this work with Figure 1 of ref 45). The origin of this effect was not investigated, as it was deemed irrelevant for the current work. pH and pD values were adjusted through the addition of ammonium hydroxide and ammonium-d4 deuterioxide, respectively, and measured with an AB15 pH meter (Fisher). Reported pD values were corrected for isotope effects based on the relation pD ) pH meter reading + 0.4.54 Pulsed Isotope Labeling. On-line pulsed HDX was carried out by employing a two-syringe on-line mixing setup similar to that used for earlier studies from our laboratory.55 Syringe 1 contained aqueous hemoglobin solution at a concentration of 500 µM in 2 mM ammonium acetate (pH 8.5), and syringe 2 contained D2O at pD 8.5. They both were advanced simultaneously by syringe pumps (Harvard Apparatus, South Natick, MA) at flow rates of 10 and 40 µL/min, respectively. The solutions were combined at a mixing tee, which empties into a 1-cm-long fusedsilica capillary (TSP075150, 75-µm i.d., Polymicro Technologies, Phoenix, AZ) for isotope labeling to take place in 80% D2O at a final protein concentration of 100 µM. The total flow rate of 50 µL/min corresponds to a protein residence time within the labeling capillary of 53 ms. The limited duration of this time interval ensures that the labeling event is short compared to the interconversion time scale of any putative binding equilibria. Longer labeling times might compromise the viability of the experimental strategy illustrated in Figure 1. Slightly basic solution conditions were used to ensure a sufficiently high degree of HDX during the labeling pulse. At pD 8.5, the exchange of unprotected (54) Glasoe, P. K.; Long, F. A. J. Am. Chem. Soc. 1960, 64, 188-190. (55) Simmons, D. A.; Dunn, S. D.; Konermann, L. Biochemistry 2003, 42, 58965905.

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amide hydrogens occurs on a time scale of 10 ms.41,56 Previous work indicates that these solution conditions are not detrimental to the stability of hemoglobin in solution.44 ESI-MS and MS/MS. Data were recorded on a Q-TOF Ultima API mass spectrometer (Waters/Micromass) that had been modified by the manufacturer to allow precursor ion selection in MS/MS mode up to m/z 8000. A custom-built ESI source was used to accommodate the 1-cm-long labeling capillary. The sprayer voltage was set at +4 kV. The mass spectrometer was operated by employing elevated pressure conditions in the first pumping stage to facilitate the transmission and preservation of large noncovalent complexes.57,58 For this purpose, the first stage roughing pump was throttled by a speedy valve to a backing Pirani gauge reading of 5 mbar. Setting the source temperature to 80 °C at a cone voltage of 30 V was found to give optimal signal intensities for the hemoglobin tetramer. MS/MS measurements were carried out using argon as collision gas. The collision voltages used for fragmenting hemoglobin monomers, dimers, tetramers, hexamers, and octamers were 68, 78, 103, 123, and 135 V, respectively. Cs(n+1)In cluster ions were used for mass calibration. In agreement with previous studies,59 isotope back exchange in the ion sampling region was found to be negligible for the Z-spray system used. Prior to analysis, minimal smoothing was applied to the experimental data, using the MassLynx software package supplied by the instrument manufacturer. HDX mass shifts ∆M of the apo-R and apo-β MS/MS product ions were calculated from the peak maximums using the relationship ∆M ) (m/z × i) - (0.8 × MD × i) - (0.2 × MH × i) - M0. Here, M0 is the mass of the unlabeled subunit, 15 054 Da for Ra, 15 954 Da for βa, and (15 954 + 32) Da for βoxa; i is the ion charge state, MD and MH are the masses of deuterium (2.014 Da) and protium (1.008 Da), respectively. The total number of exchangeable hydrogens in apo-R globin is 227, whereas the slightly larger β subunit has 244 exchangeable sites. These numbers include contributions from the amide backbone, the amino acid side chains, and the termini. RESULTS AND DISCUSSION A typical hemoglobin ESI mass spectrum acquired after online pulsed HDX is depicted in Figure 2. In agreement with earlier studies,44,45 these data reveal the presence of ions corresponding to the intact tetrameric quaternary structure ((Rhβh)2), as well as dimeric (Rhβh, Rhβoxa), monomeric (Rh, βoxa), hexameric ((Rhβh)3), and octameric ((Rhβh)4) species. As expected from the discussion above, the signals of the multimeric protein ions are substantially broadened and shifted to higher mass due to incomplete desolvation.52,53 MS/MS dissociation of these species results in wellresolved peaks corresponding to the monomeric subunits in their apo form (Figure 3). This tandem mass spectrometry approach not only facilitates the precise readout of HDX levels but it also allows the labeling behavior of the R and β subunits to be monitored separately. As an interesting side aspect, it is noted that the most abundant product species appear in disproportionately high charge states. (56) Bai, Y.; Milne, J. S.; Mayne, L.; Englander, S. W. Proteins: Struct. Funct. Genet. 1993, 17, 75-86. (57) Tahallah, N.; Pinkse, M.; Maier, C. S.; Heck, A. J. R. Rapid. Commun. Mass Spectrom. 2001, 15, 596-601. (58) Chernushevich, I. V.; Thomson, B. A. Anal. Chem. 2004, 76, 1754-1760. (59) Clemens, M.; Wayne, K.; Winger, B. E. Proceedings of the 52nd ASMS Conference of the American Conference on Mass Spectrometry, Nashville, TN 2004; MPH 099.

Figure 4. MS/MS product ion spectrum of unlabeled(Rhβh)2 in the 17+ charge state. Monomeric apo-R and apo-β subunits (denoted as Ra and βa, respectively) represent the most abundant dissociation products. In addition, the spectrum shows a weak residual precursor ion signal, as well as unresolved products at higher m/z. The intensity scale of the high m/z region has been expanded for better visualization.

Figure 3. (A) (Rhβh)2 17+ signal of pulse-labeled hemoglobin, taken from the spectrum in Figure 2. The data are plotted on a mass shift scale; i.e., the peak maximum of a completely desolvated and unlabeled ion would be located at zero. Partial MS/MS product ion scans, obtained by dissociation of (Rhβh)2 17+, are shown for apoR-globin (B) and for apo-β-globin (C) in the 7+ charge state. To allow a direct comparison of the mass shifts for the tetrameric and monomeric species the x-axis in (A) has been expanded by a factor of 4. Note that (i) the distributions in (B, C) are much more narrow than that in (A); (ii) the peak maximums in (B, C) are shifted to the left. Both of these effects are attributed to the shedding of low molecular weight adducts (likely residual solvent molecules) during collision-induced dissociation.52,53 The resulting greatly improved peak shape of the product ions (B, C) facilitates the readout of the mass shifts resulting from HDX.

For example, charge-symmetric cleavage of a (Rhβh)217+ precursor ion should result in a charge of (17+)/4 ≈ 4+ for the monomeric product species. Heme may be lost as a neutral species, but also as singly charged ion. The latter would be expected to lower the charge state of the product ions even further. Yet, the experimentally observed charge-state distributions of the R and β subunits exhibit maximums around 8+ (Figure 4). Unresolved signals in the m/z range above that of the precursor are attributed to fragment ions in lower charge states, as required for the overall charge balance of the dissociation process. The asymmetric charge partitioning seen here for hemoglobin is consistent with data previously reported for other protein complexes, although the exact mechanisms underlying this phenomenon are still a matter of debate.60-62 The most relevant aspect of this dissociation (60) Felitsyn, N.; Kitova, E. N.; Klassen, J. S. Anal. Chem. 2001, 73, 46474661. (61) Jurchen, J. C.; Williams, E. R. J. Am. Chem. Soc. 2003, 125, 2817-2826. (62) Jurchen, J. C.; Garcia, D. E.; Williams, E. R. J. Am. Soc. Mass Spectrom. 2004, 15, 1408-1415.

behavior for the present study is that it allows a precise measurement of HDX mass shifts in the R and β subunits of the various protein assemblies. Without this MS/MS approach, the determination of HDX levels for the poorly desolvated ions in the hemoglobin spectrum (Figures 2 and 3A) would be difficult. Selected data obtained in MS/MS experiments on precursor ions representing different quaternary structures are depicted in Figure 5. The subunits generated from tetrameric hemoglobin (Figure 5C-F) show mass shifts that are lower than those observed for dimeric complexes (Figure 5G, H). These, in turn, exhibit lower HDX levels than monomeric species (Figure 5I, J). Hexamers (Figure 5K, L) and octamers (Figure 5M, N) cover a mass shift range similar to that observed for the tetramer. A comprehensive summary of the HDX characteristics observed for all the major species in the hemoglobin mass spectrum is given in Figure 6. It is convenient to discuss the individual types of protein structures separately. Tetrameric Hemoglobin (rhβh)2. Protein tetramers appear in charge states 18+ to 15+. Interestingly, the charge state of these ions is correlated with the extent of isotope exchange. The dissociation products of the 15+ charge state show mass shifts of ∼70 Da, which is the lowest of any of the species in Figure 6. For more highly charged tetrameric ions, the extent of HDX steadily increases, up to a value of ∼77 Da for 18+. This behavior reveals that the different tetramer peaks in the spectrum represent different structures of the (Rhβh)2 complex in solution. Increasing charge states correspond to an increasing degree of structural perturbation. The conformational heterogeneity uncovered by pulsed HDX is consistent with recent work from our laboratory that addressed the kinetic mechanism of acid-induced hemoglobin unfolding.45 It was found that the rate of tetramer decay increases with increasing charge state. This was taken as evidence that higher charge states of the tetramer represent solution-phase conformations that are less resilient to a change in pH. The current study implies that the susceptibility to undergo acid-induced unfolding is correlated with a tendency to undergo more extensive HDX. These results are in line with the widely accepted notion Analytical Chemistry, Vol. 78, No. 5, March 1, 2006

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Figure 5. Partial MS/MS product ion scans, obtained by dissociation of hemoglobin complexes in various subunit compositions after pulse labeling. The precursor and product ion used are indicated in each panel. Data are plotted on a mass shift scale; i.e., the peak maximums of the unlabeled subunits would be located at zero. Panels on the left-hand side refer to the R subunit, panels on the right-hand side reflect the corresponding labeling behavior of the β subunit. The data in panel H for monomeric βoxa were derived directly from the hemoglobin spectrum in Figure 2, without employing MS/MS. Dashed vertical lines indicate the locations of peak maxima. Panels (A) and (B) show the results of control experiments obtained on nonlabeled hemoglobin.

that proteins can populate a range of different coexisting structures, even under nondenaturing conditions.63,64 It is interesting that the pulse-labeling approach used here is capable of directly probing this structural heterogeneity. Dimers rhβh and rhβoxa. For the hypothetical case where hemoglobin in solution exists exclusively as a tetramer (scenario B, see introduction), ions of the composition Rhβh and Rhβoxa would represent dissociation artifacts generated during ESI. Under such conditions, the mass shifts observed for the R and β subunits in the dimeric ions should fall within the range of those observed for the tetramer (Figure 1B). Inspection of the data in Figure 6 (63) Frauenfelder, H.; Sligar, S. G.; Wolynes, P. G. Science 1991, 254, 15981603. (64) Nienhaus, G. U.; Mu ¨ ller, J. D.; McMahon, B. H.; Frauenfelder, H. Physica D 1997, 107, 297-311.

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reveals that this is not the case. Rather, the β subunits of the dimeric species exhibit labeling levels that are significantly higher than the corresponding data measured for the tetramer. Higher mass shifts are also observed for the R subunits, although the differences relative to the tetramer are somewhat less pronounced. These observations strongly suggest that the dimeric ions observed in the hemoglobin ESI mass spectrum are derived from dimeric proteins in solution; that is, they do not represent dissociation products of the ESI process. This behavior is consistent with scenario A (Figure 1A). The occurrence of somewhat larger mass shifts for the β subunits may be partly due to the slightly higher number of exchangeable hydrogens, when compared to R. In addition, it likely reflects a higher degree of unfolding of the β subunits, which is in agreement with the “asymmetric” behavior of R and β reported in a previous study.44 Both Rhβh and Rhβoxa exhibit larger HDX levels in the 12+ than in the 11+ charge state. This behavior is reminiscent of the correlation between charge state and labeling behavior seen for the tetramer, thus once again indicating the presence of heterogeneous solution-phase conformations. Monomers βoxa and rh. Very large mass shifts close to 100 Da are observed for βoxa. This extensive labeling strongly suggests that βoxa ions originate from monomeric proteins in solution. The high HDX levels are indicative of a significant degree of unfolding, which is consistent with the fact that the βoxa ions appear in high charge states, ranging up to 22+. Also the mass shifts observed for monomeric Rh are higher than those seen for the tetramer. These observations provide evidence that the monomeric species in the hemoglobin mass spectrum are not ESI dissociation artifacts generated from tetrameric hemoglobin. The data obtained in this work do not provide direct information regarding the origin of methionine oxidation in βoxa. However, it seems most likely that this modification is the result of solutionphase processes, similar to those observed for other proteins.46-49 In principle, oxidation could be a hydroxyl radical-mediated process occurring in the ion source of the mass spectrometer. However, such a mechanism requires excessive sprayer voltages that lead to corona discharge conditions.50,51 In the experiments described here, there was no indication for the presence of a discharge. It is not surprising that the R subunit is unaffected by oxidation, because it possesses only a single methionyl residue that is buried deep within the tertiary structure. In contrast, the β subunit contains three methionines, all of which are solventexposed in the X-ray structure of the protein.65 Hexamers (rhβh)3 and Octamers (rhβh)4. If hemoglobin hexamers and octamers were solution-phase species, these structures should be highly protected against HDX, resulting in lower mass shifts than those observed for (Rhβh)2 (Figure 1A). Interestingly, Figure 6 reveals that this is not the case. The mass shifts of (Rhβh)3 and (Rhβh)4 fall within the range of HDX levels covered by the tetramer. It might be argued that the relatively high HDX levels of hexamer and octamer could be due to partial unfolding. However, the low charge states of these complexes (∼3+ per subunit, compared to ∼4+ for the tetramer) makes this possibility appear highly unlikely. Thus, it is concluded that hexameric and octameric ions in the hemoglobin mass spectrum are not derived from the corresponding quaternary structures in solution. Instead, they represent clustering artifacts of the ESI (65) Mueser, T. C.; Rogers, P. H.; Arnone, A. Biochemistry 2000, 39, 1535315364.

Figure 6. Mass shifts of hemoglobin R (black) and β (gray) subunits after pulse labeling as determined by MS/MS dissociation of various ions in the spectrum. The precursor ions used, together with their charge states, are indicated along the horizontal axis. Data displayed for dimers, tetramers, hexamers, and octamers represent the average values obtained from mass shifts of the differently charged product ions (see Figure 4). Standard deviations of these data sets are indicated by error bars. Mass shifts of βoxa were determined directly from the spectrum in Figure 2, i.e., without employing MS/MS.

process. Octameric clusters likely originate from pairs of tetramers that interact with each other during ESI. Loss of a dimeric unit from such an assembly could result in hexamer formation. Alternatively, hexameric clustering artifacts could result from the interaction of tetramers and dimers, both of which are solutionphase species. The latter mechanism might result in a high-mass shoulder in the hexamer profiles depicted in Figures 4I, J. Unfortunately, the relatively broad peak shapes and the persistence of residual adduction make it impossible to verify the presence of such a feature. CONCLUSIONS This study employed a combination of pulsed isotope labeling and MS/MS for probing the origin of several protein quaternary structures in the hemoglobin ESI mass spectrum. Adopting a very simplistic point of view, we had originally proposed two possible limiting cases. In scenario A, all of the ions observed in the spectrum were derived directly from the corresponding solutionphase species. In scenario B, proteins in binding states other than tetramers were artifacts of the ESI process. The labeling data obtained in this work reveal a situation that involves elements of both scenarios. Monomeric species exhibit HDX levels that are significantly higher than those seen for the tetramer, while dimeric structures cover an intermediate range. This behavior reveals that dimers and monomers are not dissociation artifacts of tetrameric hemoglobin but that they represent preexisting solution-phase species. This result is supported by previous ESI-MS studies on the unfolding of hemoglobin under equilibrium44 and nonequilibrium45 conditions. The HDX levels seen for hexameric and octameric ions fall within the range of those observed for the tetramer. This observation strongly suggests that hexamers and octamers represent nonspecific clustering artifacts that do not correspond to solution-phase quaternary structures. It appears that the pulsed HDX MS/MS technique introduced here represents a viable approach for deciphering the origin of

the various quaternary structures that are commonly observed in the ESI mass spectra of protein complexes. Dissociation products and clustering artifacts can be readily distinguished from “real” species, i.e., those that faithfully reflect the binding state of the protein in solution. The approach used represents a valuable tool that can help defining the conditions under which ESI-MS can be used as an alternative to more traditional techniques for studying protein-ligand and protein-protein interactions. Additional information on the structure of the various complexes could be obtained in gas-phase fragmentation experiments that can provide spatially resolved HDX data.40 Ideally, these studies would employ an MS/MS/MS strategy, where precursor protein complexes are first selected, then dissociated to generate individual subunits, and subsequently fragmented by cleavage of the peptide backbone. The feasibility of such an approach will be explored in future studies. It will also be interesting to see whether the technique reported in this work is applicable to more complicated systems, such as larger multisubunit complexes or glycoproteins. ACKNOWLEDGMENT We thank Douglas A. Simmons and Derek J. Wilson for helpful discussions and critical reading of the manuscript. Lisa Heydorn provided expert technical support with the newly installed instrument used for this study. Financial support was provided by the Natural Sciences and Engineering Research Council of Canada (NSERC), the Canada Foundation for Innovation (CFI), the Provincial Government of Ontario, The University of Western Ontario, and the Canada Research Chairs Program.

Received for review September 21, 2005. Accepted December 28, 2005. AC051687E

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