Signal Response of Coexisting Protein Conformers in Electrospray

activity should also enhance the ionization efficiency in cases where the CRM applies. .... Cone and desolvation gas flow rates were 50 and 500 L ...
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Anal. Chem. 2007, 79, 2499-2506

Signal Response of Coexisting Protein Conformers in Electrospray Mass Spectrometry Mark C. Kuprowski and Lars Konermann*

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

Electrospray ionization mass spectrometry (ESI-MS) is a commonly used tool for characterizing conformational changes of proteins in solution. Different conformations can be distinguished on the basis of their ESI charge state distributions. ESI-MS studies carried out under semidenaturing conditions result in bi- or multimodal distributions that reflect the presence of coexisting conformers. This study explores whether the concentration ratios of these species in solution are reflected in the measured ion intensities. Experiments on two model proteins, lysozyme and myoglobin, reveal that non-native polypeptide chains tend to result in a much stronger signal response than natively folded species. The measured ion intensity ratios can differ from the actual concentration ratios by as much as 2 orders of magnitude. It is proposed that the higher ionization efficiency of unfolded proteins is due to their partially hydrophobic character, which results in a larger surface activity and facilitates protein transfer into ion-producing progeny droplets. Conversely, natively folded proteins have a lower affinity for the air/ liquid interface, such that ionization of these conformers is suppressed. The extent of ion suppression is strongly dependent on the experimental conditions such as flow rate and protein concentration, which determine if ESI occurs in a charge deficient or a charge surplus regime. These aspects should be taken into account for the design of ESI-MS-based protein folding experiments and for studies that use ion intensity ratios for the determination of protein-ligand binding affinities. Electrospray ionization mass spectrometry (ESI-MS) has become an important tool for studying a wide range of analytes, including proteins and noncovalently bound ligand-protein complexes.1 In positive ion mode, droplets of analyte solution carrying excess positive charge are emitted from a Taylor cone. Solvent evaporation increases the charge density on the droplet surface until Coulombic repulsion overcomes the surface tension of the liquid, such that smaller offspring droplets are formed by jet fission.2 Due to the uneven nature of these fission events, the progeny droplets carry away only a small fraction of mass but a * To whom correspondence should be addressed. Phone: 519-661-2111 ext. 86313. Fax: 519-661-3022. E-mail: [email protected]. Web: http:// publish.uwo.ca/∼konerman. (1) Fenn, J. B. Angew. Chem., Int. Ed. 2003, 42, 3871-3894. (2) Gomez, A.; Tang, K. Phys. Fluids 1994, 6, 404-414. 10.1021/ac0620056 CCC: $37.00 Published on Web 02/09/2007

© 2007 American Chemical Society

disproportionately high amount of charge.3 Subsequent evaporation and fission events ultimately lead to the formation of nanometer-sized droplets from which multiply protonated analyte ions are produced.4 According to the charged residue model (CRM), solvent evaporation to dryness releases the analyte which retains some of the droplet’s charge.5 In contrast, the ion evaporation model (IEM) stipulates that charged analytes can be ejected from the droplet surface.6 The protonation states of ions generated from proteins and other large analytes are close to the Rayleigh limit, thereby providing support for the notion that these species are ionized via the CRM mechanism.7-11 The appearance of protein ESI charge state distributions depends on a number of experimental parameters.10,12-15 By far the most important factor, however, is the protein solution-phase conformation. ESI mass spectra of unfolded polypeptide chains generally show wide distributions centered around relatively high protonation states. In contrast, ions derived from tightly folded proteins are less extensively charged and show more narrow distributions.4,16 This empirical relationship provides the basis for the widespread use of ESI-MS as a tool for monitoring protein conformational changes in solution.17 Several factors have been proposed to account for the higher charge states seen for unfolded proteins. These include the accessibility of protonation sites,18,19 their spacing,20 the surface area of the protein,13 the occurrence of partial charge compensation,9,21,22 and the increased conforma(3) Kebarle, P.; Tang, L. Anal. Chem. 1993, 65, 972A-986A. (4) Kebarle, P.; Peschke, M. Anal. Chim. Acta 2000, 406, 11-35. (5) Dole, M.; Mack, L. L.; Hines, R. L.; Mobley, R. C.; Ferguson, L. D.; Alice, M. B. J. Chem. Phys. 1968, 49, 2240-2249. (6) Iribarne, J. V.; Thomson, B. A. J. Chem. Phys. 1975, 64, 2287-2294. (7) de la Mora, F. J. Anal. Chim. Acta 2000, 406, 93-104. (8) Felitsyn, N.; Peschke, M.; Kebarle, P. Int. J. Mass Spectrom. Ion Processes 2002, 219, 39-62. (9) Nesatyy, V. J.; Suter, M. J.-F. J. Mass Spectrom. 2004, 39, 93-97. (10) Iavarone, A. T.; Williams, E. R. J. Am. Chem. Soc. 2003, 125, 2319-2327. (11) Kaltashov, I. A.; Mohimen, A. Anal. Chem. 2005, 77, 5370-5379. (12) Pan, P.; McLuckey, S. A. Anal. Chem. 2003, 75, 1491-1499. (13) Fenn, J. B. J. Am. Soc. Mass Spectrom. 1993, 4, 524-535. (14) Samalikova, M.; Grandori, R. J. Am. Chem. Soc. 2003, 125, 13352-13353. (15) Wang, G.; Cole, R. B. In Electrospray Ionization Mass Spectroscopy; Cole, R. B., Ed.; John Wiley & Sons, Inc.: New York, 1997; pp 137-174. (16) Dobo, A.; Kaltashov, I. A. Anal. Chem. 2001, 73, 4763-4773. (17) Kaltashov, I. A.; Eyles, S. J. Mass Spectrom. Rev. 2002, 21, 37-71. (18) Chowdhury, S. K.; Katta, V.; Chait, B. T. J. Am. Chem. Soc. 1990, 112, 9012-9013. (19) Katta, V.; Chait, B. T. J. Am. Chem. Soc. 1991, 113, 8534-8535. (20) Downard, K. M.; Biemann, K. Int. J. Mass Spectrom. Ion Processes 1995, 148, 191-202. (21) Samalikova, M.; Grandori, R. J. Mass Spectrom. 2003, 38, 941-947. (22) Prakash, H.; Mazumdar, S. J. Am. Soc. Mass Spectrom. 2005, 16, 14091421.

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tional flexibility of unfolded proteins that may facilitate intramolecular charge solvation.23 Assuming CRM conditions, it has been suggested that noncompact protein conformations will increase the size of the final nanodroplets, possibly by inducing nonspherical shapes, such that these droplets can accommodate a larger number of charges at the Rayleigh limit.7 Whereas the relationship between solution-phase conformation and charge state distribution continues to be an active area of research, surprisingly little work has been done to explore how protein conformation affects signal intensities in ESI-MS. Of particular interest is the behavior of water-soluble proteins which represent the most commonly encountered class of polypeptide chains. The native state of these proteins is characterized by a hydrophobic core, consisting of tightly packed nonpolar side chains that tend to avoid contact with the solvent. Most polar and charged residues are located on the exterior, where they can favorably interact with the surrounding water. Unfolding exposes nonpolar residues to the solvent and increases the hydrodynamic radius of the polypeptide chain.24 These and other factors might be expected to affect the ion yield during ESI. A better understanding of the relationship between conformation and ESI-MS signal response is essential for the proper interpretation of data obtained for samples containing coexisting solution-phase conformers. Spectra of proteins obtained under semidenaturing conditions usually show multimodal charge state distributions, reflecting the presence of different polypeptide conformations that are in equilibrium with each other in solution. Unfortunately, these data do not provide direct information on the concentrations of the various species.16 A related issue is the determination of protein-ligand binding affinities by ESI-MS. Studies of this kind often rely on the assumption that the signal intensity ratio of free to bound receptor in ESI-MS matches the corresponding concentration ratio in solution.25,26 This supposition may not always be justified, especially in cases where binding is accompanied by major conformational changes.27-29 Studies comparing the signal response of various analytes in ESI-MS typically focus on the ionization efficiency, presuming that the subsequent ion sampling, transmission, and detection are not strongly analyte-dependent.12,30 Nano-ESI can have ionization efficiencies approaching 100%, whereas regular ESI sources that operate in the µL min-1 flow rate range have efficiencies that are orders of magnitude lower.31 Expanding on work by Kebarle and co-workers,3 Enke et al. developed a model on the basis of the presence of two different phases in ESI droplets, namely the electrically neutral interior and the surface layer that carries excess charge. The model assumes that analytes undergo partitioning between these two phases and that the ionization efficiency is (23) Konermann, L.; Simmons, D. A. Mass Spectrom. Rev. 2003, 22, 1-26. (24) Fersht, A. R. Structure and Mechanism in Protein Science; W. H. Freeman & Co.: New York, 1999. (25) Wang, W.; Kitova, E. N.; Klassen, J. S. Anal. Chem. 2003, 75, 4945-4955. (26) Peschke, M.; Verkerk, U. H.; Kebarle, P. J. Am. Soc. Mass Spectrom. 2004, 15, 1424-1434. (27) Gunasekaran, K.; Tsai, C.-J.; Kumar, S.; Zanuy, D.; Nussinov, R. Trends Biochem. Sci. 2003, 28, 81-85. (28) Verkhivker, G. M.; Bouzida, D.; Gehlhaar, D. K.; Rejto, P. A.; Freer, S. T.; Rose, P. W. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 5148-5153. (29) Wright, P. E.; Dyson, H. J. J. Mol. Biol. 1999, 293, 321-331. (30) Pan, P.; McLuckey, S. A. Anal. Chem. 2003, 75, 5468-5474. (31) Smith, R. D.; Shen, Y.; Tang, K. Acc. Chem. Res. 2004, 37, 269-278.

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proportional to the analyte concentration in the surface layer.32,33 Analytes carrying nonpolar moieties have a high surface activity, resulting in a propensity to reside near the air/liquid interface. Assuming that these species also possess ionizable groups, it is predicted that they will compete more effectively for excess charge on the droplet surface than polar molecules, which preferentially reside in the droplet interior. In the case of analyte mixtures, therefore, most of the available excess charge will be taken up by easily ionizable species with high surface activities, such that signals originating from other molecules are suppressed. Additionally, the low desolvation energies of nonpolar analytes may promote their transition into a solvent-free environment.34 These effects have been verified experimentally for small peptides and many other low molecular weight compounds.13,33-38 Although Enke’s equilibrium partitioning model had originally been developed assuming IEM conditions, it seems that a high surface activity should also enhance the ionization efficiency in cases where the CRM applies. This notion is based on the fact that small offspring droplets are generated from the outermost layers of the parent droplets and it is these offspring droplets that eventually lead to the formation of gas-phase ions.3,39,40 Thus, the increased hydrophobicity and surface affinity of unfolded proteins would be expected to translate into more intense signals when compared to natively folded polypeptide chains.16 Surprisingly, there appear to be no previous studies that have directly tested this prediction. The goal of the current work is to explore in how far the concentration ratio of coexisting protein conformers can be deduced from the abundance ratio of the corresponding ionic species in ESI-MS. The cases studied here reveal a general trend for unfolded proteins to result in higher signal intensities. These observations are consistent with the view that the increased hydrophobicity of unfolded polypeptide chains allows them to compete more effectively for excess charge, thereby suppressing the signals originating from natively folded proteins. The extent of ion suppression is strongly dependent on the experimental conditions. EXPERIMENTAL SECTION Materials. Horse skeletal muscle myoglobin, chicken egg white lysozyme, 1,4-dithiothreitol (DTT), deuterated acetic acid, and HPLC grade glacial acetic acid were purchased from Sigma, St. Louis, MO. Ammonium acetate and formic acid were bought from Fluka, Buchs, Switzerland. Reagent grade ammonium hydroxide and glass distilled acetonitrile were from Caledon Laboratories, Georgetown, ON, Canada. Deuterium oxide was obtained from Cambridge Isotope Laboratories, Andover, MA. Ammonium-d4 deuteroxide was purchased from Isotec, Miamisburg, OH. The water used was subjected to reverse osmosis and subsequently glass distilled. pH and pD values were measured (32) Enke, C. G. Anal. Chem. 1997, 69, 4885-4893. (33) Cech, N. B.; Enke, C. G. Mass Spectrom. Rev. 2001, 20, 362-387. (34) Null, A. P.; Nepomuceno, A. I.; Muddiman, D. C. Anal. Chem. 2003, 75, 1331-1339. (35) Cech, N. B.; Enke, C. G. Anal. Chem. 2001, 73, 4632-4639. (36) Cech, N. B.; Krone, J. R.; Enke, C. G. Anal. Chem. 2001, 73, 208-213. (37) Zhou, S.; Cook, K. D. J. Am. Soc. Mass Spectrom. 2001, 12, 206-214. (38) Mirzaei, H.; Regnier, F. Anal. Chem. 2006, 78, 4175-4183. (39) Schmidt, A.; Karas, M.; Du ¨ lcks, T. J. Am. Soc. Mass Spectrom. 2003, 14, 492-500. (40) Tang, K.; Smith, R. D. J. Am. Soc. Mass Spectrom. 2001, 12, 343-347.

with an AB15 pH meter (Fisher Scientific, Nepean, ON). Reported values for D2O-containing solutions were corrected for isotope effects by using the relation pD ) pH meter reading + 0.4.41 Lysozyme Experiments. The 500 mM lysozyme (pI 9.3)42 was extensively dialyzed against water using Slide-A-Lyzer cassettes (Pierce, Rockford, IL) with a nominal 7 kDa molecular weight cutoff. Disulfide reduction was performed by incubating 100 µM protein in 10 mM DTT at 75 °C for 1 h. DTT was not removed for mass spectrometric analysis or UV-vis absorption measurements. Samples containing both disulfide-intact and reduced lysozyme were analyzed within less than 15 min of mixing. Reduction of the disulfide-intact protein at room temperature under these conditions is negligible. Prior to ESI-MS the lysozyme concentrations were adjusted on the basis of UV-vis absorption measurements at a wavelength of 280 nm, using a Cary 100 spectrophotometer (Varian, Mississauga, ON, Canada). DTT does not absorb at 280 nm to any significant extent. The absorption coefficients of disulfide-intact and reduced lysozyme are 37 470 and 37 970 M-1 cm-1, respectively (calculated using the ExPASy ProtParam routine). Before mass spectrometric analysis, 0.5% (v/v) glacial acetic acid was added to each solution, resulting in pH 3.0. Myoglobin Experiments. Stock solutions of 500 µM myoglobin (pI ∼7.0)42 in 20 mM ammonium acetate were centrifuged to remove small amounts of insoluble debris. The supernatant was dialyzed against 20 mM ammonium acetate using the same SlideA-Lyzer system as for lysozyme. Post dialysis, samples were flash frozen in liquid nitrogen as 1 mL aliquots and stored at -80 °C. Consistent with its amino acid sequence the mass of the apomyoglobin was found to be 16952 Da, the heme group in the holoprotein accounts for an additional 616 Da. Immediately prior to use, the 1 mL aliquots were diluted to 112 µM protein by addition of 20 mM ammonium acetate at pH 9.3. Subsequently, 50 µL glacial acetic acid were added, before raising the pH back to 9.3 with ammonium hydroxide. The resulting solutions were mixed with acetonitrile to final concentrations of 27, 30, or 33% (v/v) while maintaining the pH at 9.3. Mass shifts in hydrogen/ deuterium exchange (HDX) experiments were calculated by converting the mass-to-charge ratio values of the original data according to ∆M ) (r × z) - mcharge - M0, where r is the massto-charge ratio, z is the charge state of the ion, mcharge is the combined mass of the z charge carriers (protons and deuterons, their numbers being dependent on the D2O to H2O ratio of the solution), and M0 is the mass of the unlabeled protein. On-Line Pulsed Hydrogen/Deuterium Exchange. Myoglobin was subjected to pulsed HDX using two different on-line rapid mixing approaches. For all HDX experiments the pD of the labeling buffer was adjusted to 9.3, and the acetonitrile concentration used matched that of the protein solution. (i) Direct on-line labeling was carried out using a continuousflow capillary mixing setup that has been described previously (Figure 1a).43 Briefly, the plungers of two syringes (SGE, Austin, TX) were advanced simultaneously by syringe pumps (Harvard Apparatus, South Natick, MA). Syringe 1 (10 µL min-1) contained protein solution, whereas syringe 2 (40 µL min-1) contains labeling buffer. The contents of the syringes were passed through a mixer

and into a reaction chamber, the volume of which had been adjusted to result in a labeling time interval of 60 ms. Isotope exchange was terminated at the reaction chamber exit by solvent evaporation during ESI on the time scale of 1 ms.3 The final protein concentration was ∼15 µM. Effective mixing was verified by using bradykinin as internal standard, which exhibited a labeling level around 79.5%, i.e., very close to the 80% value that would be expected based on the 4:1 (v/v) D2O to H2O ratio of the labeling solvent.44 (ii) On-line labeling followed by acid-quenching employed a modified version of the setup just described (Figure 1b), involving two sequential mixing steps. Following 60 ms of pulsed HDX the solution exiting the reaction chamber was mixed with 10% aqueous formic acid. This quenching solution was delivered by a third syringe pump operating at 50 µL min-1. The resulting mixture was then transferred into a standard Z-spray ESI source by means of a PEEK connector, resulting in a quenching time interval of 30 s at a total flow rate of 100 µL min-1. Mass Spectrometry. All ESI-MS experiments were carried out on a Q-TOF Ultima API instrument (Waters/Micromass, Manchester, U.K.) operated in positive ion mode at a capillary voltage of 3.0 kV. Cone and desolvation gas flow rates were 50 and 500 L h-1 respectively. The settings of the rf-only quadrupole were adjusted to provide constant ion transmission across the range of interest. Lysozyme experiments were carried out by using a cone voltage of 35 V, a desolvation temperature of 120 °C, and a source temperature of 80 °C. For myoglobin experiments the desolvation temperature was lowered to 100 °C, and a cone voltage of zero was used to prevent thermal or collision-induced dissociation of heme-protein complexes during ion sampling.45 All spectra

(41) Glasoe, P. K.; Long, F. A. J. Am. Chem. Soc. 1960, 64, 188-190. (42) Moritz, R. L.; Simpson, R. J. Nat. Methods 2005, 2, 863-873. (43) Wilson, D. J.; Konermann, L. Anal. Chem. 2003, 75, 6408-6414.

(44) Hossain, B. M.; Simmons, D. A.; Konermann, L. Can. J. Chem. 2005, 83, 1953-1960. (45) Collings, B. A.; Douglas, D. J. J. Am. Chem. Soc. 1996, 118, 4488-4489.

Figure 1. Mixing schemes employed for pulsed hydrogen/deuterium exchange (HDX) experiments. (a) On-line mixing setup where protein in aqueous solution is exposed to a D2O-based labeling buffer at mixer M1. Following 60 ms of pulsed HDX, the mixture reaches the ESI source of the mass spectrometer. (b) Double-mixing setup incorporating an acid quenching step. Labeling of the protein proceeds as in (a), but following 60 ms of HDX the mixture is exposed to 10% formic acid at mixer M2, resulting in pH 2.5. ESI occurs after 30 s of acid quenching.

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Figure 2. ESI mass spectra of 1 µM lysozyme in aqueous solution at pH 3.0. The spectrum of folded, disulfide-intact lysozyme (lyso-F) is depicted in (a); (b) shows data for the unfolded, disulfide-reduced protein (lyso-U). The spectra in (c) and (d) were obtained for mixtures of lyso-U and lyso-F in a 1:1 molar ratio. The solution flow rate was 5 µL min-1 for (a)-(c) and 100 µL min-1 for (d).

shown represent the average of at least 15 scans, acquired with an integration time of 10 s. RESULTS AND DISCUSSION Comparative Studies on Native and Disulfide-Reduced Lysozyme. The presence of four disulfide bridges in native lysozyme makes this protein an excellent model system for studying the ESI-MS signal response of coexisting conformers. Reductive cleavage of these -S-S- bonds greatly destabilizes the native conformation.46 Disulfide-intact lysozyme (referred to as “lyso-F”) retains a tightly folded structure under mildly acidic conditions, whereas the reduced protein (“lyso-U”) is extensively unfolded.47,48 This behavior is illustrated by the ESI mass spectra in Figure 2, which were recorded in water at pH 3.0. At a solution flow rate of 5 µL min-1 and a protein concentration of 1 µM the spectrum of lyso-F is dominated by the 10+ charge state (Figure 2a). In contrast, the spectrum of lyso-U shows a much wider charge state distribution with a maximum around 17+ (Figure 2b). As expected, the mass measured for lyso-U (14313 Da) is larger than that of lyso-F (14305 Da) by eight units as a result of disulfide reduction. (46) Acharya, A. S.; Taniuchi, H. Mol. Cell. Biochem. 1982, 44, 129-148. (47) Loo, J. A.; Edmonds, C. G.; Udseh, H. R.; Smith, R. D. Anal. Chem. 1990, 62, 693-698. (48) Konermann, L.; Douglas, D. J. J. Am. Soc. Mass Spectrom. 1998, 9, 12481254.

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Figure 3. ESI mass spectra of lysozyme under conditions identical with those of Figure 2, except that the total protein concentration was increased to 100 µM. Insets represent expansions of the 11+ charge state, illustrating the mass increase associated with reduction of four disulfide bonds. The insets in panels c and d demonstrate the Gaussian decomposition procedure used to extract relative peak intensities for lyso-F (dotted lines) and lyso-U (dashed lines) in the 11+ to 8+ charge state range. The resulting fits (bold lines) are superimposed on the experimental data.

An equimolar mixture of lyso-F and lyso-U analyzed at a total protein concentration 1 µM and a flow rate of 5 µL min-1 results in the bimodal charge state distribution of Figure 2c. We define the ion abundance ratio, RESI, as

∑I

U

RESI )

z

∑

(1) IF

z

where the two sums represent the total signal intensity of lyso-U and lyso-F, respectively, for all charge states z. The data in Figure 2c correspond to an RESI value of 1.9. Virtually the same result was obtained when the flow rate was lowered to 1 µL min-1 (data not shown). At an elevated flow rate of 100 µL min-1, however, the relative contributions of lyso-U is much higher, resulting in RESI ) 4.4 (Figure 2d). Figure 3 shows the results of ESI-MS measurements under the same conditions as in Figure 2, except that the total protein concentration was increased to 100 µM. The spectra obtained for an equimolar mixture of lyso-U and lyso-F at this elevated concentration are strongly dominated by signals corresponding

to the unfolded protein, with RESI values close to 7.0 for both 5 µL min-1 (Figure 3c) and 100 µL min-1 (Figure 3d). The determination of RESI in these cases necessitated the use of a decomposition procedure due to peak overlap in the 11+ to 8+ charge state region (Figure 3, insets). The data in Figures 2 and 3 show that lyso-U tends to result in considerably higher signal intensities than lyso-F, when electrosprayed from equimolar solutions. We propose that this behavior is caused by the larger number of solvent-exposed nonpolar residues for lyso-U, which results in a more hydrophobic character. Therefore, this protein species has a higher affinity for the droplet surface from where it is preferentially transferred to offspring droplets that will eventually generate gas-phase protein ions. Lyso-F, on the other hand, tends to reside within the electrically neutral droplet interior where its hydrophilic solventexposed residues can interact favorably with water. Consequently, the transfer of lyso-F to smaller progeny droplets is not as effective. This scenario is analogous to partitioning effects seen for other types of analytes.40 The extent to which lyso-U dominates the ESI mass spectrum depends on the experimental conditions. At low flow rate and low concentration RESI deviates from the [lyso-U]: [lyso-F] concentration ratio only by a factor of 2 (Figure 2c). A much stronger bias is observed when the flow rate (Figure 2d) and/or the total protein concentration (Figure 3c,d) are increased. These trends can be rationalized on the basis of the following considerations: The average ESI charge state, qavg, of a protein is given by

∑zI

z

qavg )

z

∑I

(2)

z

z

where Iz is the abundance for ions of charge state z. For the following calculations we will use qavg ) 15, which represents the mean value derived from the data in Figures 2c,d and 3c,d. As noted previously by others,12,49 it is the molar concentration of excess charge, Cq, on the initially formed electrospray droplets that determines the upper limit of the ionization efficiency. For a protein concentration Cprot one can consider two different regimes: (1) “Charge surplus” is characterized by Cq > (Cprot × qavg). Under these conditions the amount of excess charge is sufficient to allow, in principle, the ionization of all analyte molecules in the droplet.31,39,50 (2) “Charge deficiency” prevails if Cq < (Cprot × qavg), corresponding to a regime where only a fraction of the proteins can undergo ionization. In the latter scenario, coexisting conformers within the ESI droplet will compete for the available charge, and the species that ionizes most effectively (lysoU, in the present case) will suppress the signals of other conformers. It is noted that that scenario 1 does not automatically imply an analyte ionization efficiency of 100%, because a significant fraction of Cq may be consumed in side reactions, such as the formation of protonated water clusters. (49) Wang, G.; Cole, R. Anal. Chem. 1995, 67, 2892-2900. (50) Tang, K.; Page, J. S.; Smith, R. D. J. Am. Soc. Mass Spectrom. 2004, 15, 1416-1423.

The total charge Q on the initially formed ESI droplets can be estimated on the basis of the relationship12,51

Q ) 0.7 × 8π(0γR3)1/2

(3)

where the radius R can be calculated according to12,39,51

R ) (Vf κe 0/K)1/3

(4)

with a dielectric constant of κe ) 80 and a conductivity of K ) 0.25 S m-1.12,39 Equation 4 predicts that the radius of the initial ESI droplets is 0.62 µm for a flow rate of 5 µL min-1 and 1.7 µm for 100 µL min-1. For the experiments described here the expected number of protein molecules contained within each of the droplets is between 6 × 102 (Figure 2c) and 1.2 × 106 (Figure 3d). By combination of eqs 3 and 4, Cq can be expressed as

Cq )

( )

4.2 Kγ F V f κe

1/2

(5)

where F ) 96 485 C mol-1 and the surface tension of water is γ ) 0.07 N m-1. For a flow rate of Vf ) 5 µL min-1, eq 5 predicts that Cq ) 70 µM, whereas, for 100 µL min-1, Cq ) 16 µM. The conditions of Figure 2c with (Cq ) 70 µM) > (Cprot × qavg ) 15 µM), therefore, represent a charge surplus scenario. Under these conditions charge is so abundant that a substantial fraction of lyso-F can undergo ionization, despite its relatively low surface activity. As a result, the overall ion intensity of lyso-U is not much larger than that of lyso-F. Figure 2d with (Cq ) 16 µM) ≈ (Cprot × qavg ) 15 µM) represents an intermediate regime where suppression effects become more noticeable, as evidenced by the considerably lower intensities of lyso-F ions. The data in Figure 3c with (Cq ) 70 µM) , (Cprot × qavg ) 1.5 mM) represent conditions of charge deficiency, the same is true for Figure 3d where the amount of charge/protein is even lower, (Cq ) 16 µM) , (Cprot × qavg ) 1.5 mM). Ion suppression in the latter two cases is very pronounced because a major fraction of the available charge is being utilized for the ionization of lyso-U, thereby causing exceedingly low signal intensities for lyso-F. All the ESI mass spectra in Figure 3 obtained for a protein concentration of 100 µM are shifted to lower charge states, when compared to the corresponding data for 1 µM in Figure 2. Similar effects have been noted previously,12,52 and they can be attributed to a higher degree of charge deficiency caused by the elevated analyte concentration. Similarly, the amount of charge available/ protein molecule can be reduced by raising the flow rate (eq 5). This provides an explanation for the observation that data recorded at 100 µL min-1 (Figure 3d) are skewed toward lower charge states when compared to those obtained for 5 µL min-1 (Figure 3c). Close inspection of Figures 2c,d reveals that this effect also occurs at a protein concentration of 1 µM, albeit to a much lesser extent because charge deficiency is not as severe. In summary, the experiments discussed in this section show that coexisting protein conformers can have drastically different (51) Loscertales, I. G.; de la Mora, J. F. J. Chem. Phys. 1995, 103, 5041-5060. (52) Benkestock, K.; Sundqvist, G.; Edlund, P.-O.; Roeraade, J. J. Mass Spectrom. 2004, 39, 1059-1067.

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Figure 4. (a) Mass spectrum of semi-denatured myoglobin (Mb) in 27% acetonitrile at pD 9.3. The data were obtained by electrospraying the protein immediately after exposure to deuterated labeling buffer for 60 ms, using the setup of Figure 1a. Notation: a, apoMb; h, holoMb; h*, Mb bound to a heme dimer. The signal of monomeric heme has been rescaled by a factor of 0.5 for better visualization. The peak denoted with a double asterisk corresponds to dimeric heme. Arrows highlight peak splitting for the holoMb 9+ and holoMb 8+ signals. (b) Spectrum of Mb obtained in the same way as for (a), except that the 60 ms HDX step was followed by 30 s of acidic quenching (pH 2.5) prior to ESI (using the setup of Figure 1b).

ionization efficiencies. The relative signal response of the different solution-phase species depends strongly on the experimental conditions. The interpretation of these lysozyme data has been particularly straightforward because interconversion of the two conformers was blocked by covalent modifications of the protein. As a result, the experiments could be carried out under highly controlled conditions with a well defined [lyso-U]:[lyso-F] concentration ratio. We will now proceed to explore a more complex scenario, involving coexisting protein conformers that are in equilibrium with each other. Signal Response of Coexisting Protein Conformers in Different Ligand-Binding States. The native state of myoglobin (Mb) contains a heme group that is bound in a hydrophobic pocket where it makes numerous noncovalent contacts with the protein. Additionally, the heme iron is ligated by the side chain of His93.53 Exposure of Mb to organic cosolvents under mildly basic conditions is known to induce the formation of several semiunfolded conformations, along with partial loss of heme.54 Figure 4a shows an ESI mass spectrum of Mb recorded in the presence of 27% acetonitrile at pD 9.3 (the reasons for using a D2O-based solvent system are discussed below). The spectrum is dominated by heme-free ions (apoMb) in a bimodal charge state distribution. The apoMb charge states 8+ to 10+ represent (53) Evans, S. V.; Brayer, G. D. J. Mol. Biol. 1990, 213, 885-897. (54) Simmons, D. A.; Dunn, S. D.; Konermann, L. Biochemistry 2003, 42, 58965905.

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compact solution-phase conformers, whereas more highly charged ions (around 14+) are attributed to increasingly unfolded species.16 A similar bimodal distribution is observed for holoMb, corresponding to heme-containing conformers in various degrees of compactness. The spectrum also reveals the presence of free heme in monomeric and dimeric form. Binding of the protein to heme dimers leads to the formation of double-heme Mb, denoted as h* in Figure 4a.54 Pulsed hydrogen/deuterium exchange (HDX) techniques can provide structural information complementary to that obtained from the ESI charge state distribution.55-57 The HDX characteristics of a protein are highly conformation-dependent because exchange takes place preferentially at solvent accessible sites that are not involved in hydrogen bonds.58 HoloMb has a total of 263 exchangeable hydrogens, 148 of which are located along the amide backbone, 110 on the amino acid side chains, 3 on the termini, and 2 on the heme propionates. The data in Figure 4a were obtained using a pulsed HDX strategy where the protein was exposed to deuterated labeling buffer for a short time interval (60 ms) immediately prior to ESI, using the approach outlined in Figure 1a. Slightly basic conditions had to be employed to ensure the complete exchange of freely accessible sites during this very brief labeling pulse.59 With this methodology the different chargeand ligand-binding states in the mass spectrum can be directly correlated with the deuteration behavior of the corresponding solution-phase species.23 Pulsed HDX causes peak splitting for the 9+ and 8+ holoMb ions. The high mass portion of both signals displays a shift of 185 Da, whereas the low mass portion is shifted by 105 Da relative to the unlabeled protein (Figure 5a). This behavior reveals that the holoMb 9+ and 8+ signals contain contributions from two different solution-phase conformers. Previous work has shown that the low mass component can be attributed to native holoMb, whereas the high mass contribution represents a structurally perturbed heme-bound state.44,54 All other protein signals in the spectrum exhibit HDX characteristics matching those of the holoMb 9+ and 8+ high mass component, with an average mass shift around 185 Da (Figure 5b). The observation of virtually identical mass shifts for all of the apoMb, double-heme Mb and high mass holoMb signals implies that the corresponding solutionphase protein molecules rapidly interconvert during labeling.54,60 On the basis of these HDX data, the various coexisting Mb species can be categorized into two classes. The natively folded, heme bound protein exhibits a small HDX mass shift and will be referred to as “Mb-F”. All other species, collectively termed “MbU”, represent non-native conformations exhibiting large mass shifts. Partial heme loss and unfolding are both expected to contribute to a partially hydrophobic character for the Mb-U conformers. As with the lysozyme data discussed above, it is possible to calculate the ion abundance ratio RESI which reflects the relative response of Mb-U and Mb-F in the mass spectrum. (55) Pan, J. X.; Wilson, D. J.; Konermann, L. Biochemistry 2005, 44, 8627-8633. (56) Katta, V.; Chait, B. T. J. Am. Chem. Soc. 1993, 115, 6317-6321. (57) Miranker, A.; Robinson, C. V.; Radford, S. E.; Aplin, R.; Dobson, C. M. Science 1993, 262, 896-900. (58) Krishna, M. M. G.; Hoang, L.; Lin, Y.; Englander, S. W. Methods 2004, 34, 51-64. (59) Bai, Y.; Milne, J. S.; Mayne, L.; Englander, S. W. Proteins: Struct. Funct. Genet. 1993, 17, 75-86. (60) Wagner, D. S.; Anderegg, R. J. Anal. Chem. 1994, 66, 706-711.

Figure 5. Close-up view of selected myoglobin (Mb) peaks obtained after 60 ms of on-line-pulsed HDX, using the setup depicted in Figure 1a. Acetonitrile concentrations used were 27% (a, b), 30% (c, d), and 33% (e, f). The x-axes display HDX mass shifts relative to the unlabeled protein. Panels on the left represent holoMb 9+; panels on the right show data for apoMb 17+, holoMb 16+, and doubleheme Mb 10+ (top to bottom; these peaks have been rescaled for better visualization). Peaks marked Mb-F refer to natively folded holoprotein, exhibiting a small HDX mass shift. All other peaks represent partially unfolded species (Mb-U) that show a much larger mass shift.

The data in Figure 4a for an acetonitrile concentration of 27% are strongly dominated by Mb-U, resulting in an RESI value of 11. Close-up views of selected peaks are depicted in Figure 5a,b. Slight increases in the organic content of the solution have minimal effects on the overall appearance of the mass spectrum (data not shown); however, they cause a dramatic reduction of the Mb-F intensities. RESI increases to 56 when the acetonitrile concentration is raised to 30% (Figure 5c,d). In the presence of 33% acetonitrile the Mb-F contribution becomes almost indiscernible, with RESI around 200 (Figure 5e,f). These trends reflect a progressive destabilization of the native Mb structure with increasing acetonitrile concentration, which is consistent with earlier reports.61,62 The key question in the context of this study is how the measured RESI values compare to the actual Mb-U and Mb-F mole fractions. The concentration ratio in solution can be measured by employing an acid-quench approach. In this strategy, the protein is initially labeled for 60 ms under conditions matching those discussed in the previous paragraph. However, instead of terminating HDX by desolvation during ESI (as for the data in Figures 4a and 5) the labeling pulse is ended by a second mixing step that exposes the solution to pH 2.5 for 30 s. The acidic environment lowers amide HDX rates by more than 6 orders of magnitude, such that the labeling characteristics of all the species (61) Clark, S. M.; Konermann, L. J. Am. Soc. Mass Spectrom. 2003, 14, 430441. (62) Babu, K. R.; Moradian, A.; Douglas, D. J. J. Am. Soc. Mass Spectrom. 2001, 12, 317-328.

Figure 6. Mass distributions obtained after exposing myoglobin (Mb) to 60 ms of on-line pulsed HDX, followed by 30 s of acid quenching at pH 2.5 (using the setup of Figure 1b). Acetonitrile concentrations were 27% (a), 30% (b), and 33% (c). The peak intensities IMb-U and IMb-F were determined as illustrated in (b), following a peak shape extrapolation on the basis of the data in panel a; Gaussian decomposition could not be employed in this case because of peak tailing effects. Details of the procedure used have been described elsewhere.54 For reasons of consistency the same extrapolated peak shape was used for the analysis of all three data sets. The IMb-U: IMb-F intensity ratios obtained in this way directly reflect the corresponding [Mb-U]:[ Mb-F] concentration ratios in solution.

are partially preserved.58,63 More importantly, the pH change converts all of the differentially deuterated species to highly denatured apoMb, thereby removing any ionization bias (Figure 4b).64 Therefore, after acid quenching every individual charge states displays exactly the same mass distribution, resulting in an IMb-U:IMb-F ion intensity ratio that directly reflects the [MbU]:[Mb-F] concentration ratio in solution.63 The quench-flow data exhibit mass shifts of 100 and 58 Da, for Mb-U and Mb-F, respectively (Figure 6). The fact that these shifts are smaller than for the data of Figure 5 is attributed to side chain back exchange during the 30 s quenching period, an effect that is unavoidable for this type of experiment.58,63 The mass distribution obtained in the presence of 27% acetonitrile is strongly dominated by Mb-F (Figure 6a), whereas Mb-U dominates at 33% (Figure 6c). Data measured for 30% acetonitrile represent a situation where the solution-phase concentrations of Mb-U and Mb-F are nearly equal (Figure 6b). Even without a detailed analysis, it is obvious that the [MbU]:[Mb-F] concentration ratios resulting from these quench-flow experiments are strikingly different from the corresponding RESI values determined under nonquench conditions. In the presence of 27% acetonitrile, for example, the overall ion abundance of Mb-F (63) Smith, D. L.; Deng, Y.; Zhang, Z. J. Mass Spectrom. 1997, 32, 135-146. (64) Konermann, L.; Rosell, F. I.; Mauk, A. G.; Douglas, D. J. Biochemistry 1997, 36, 6448-6454.

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in Figures 4a and 5a,b is miniscule when compared to the corresponding quench-flow mass distribution (Figure 6a). The accurate determination of peak intensity ratios in Figure 6 is slightly complicated by the overlap of the two contributions, thus necessitating the application of a baseline extrapolation (see caption of Figure 6). When using this procedure, it is found that the [Mb-U]:[Mb-F] concentration ratios for 27%, 30%, and 33% acetonitrile are 0.18, 0.61, and 3.6, respectively. These ratios are almost 2 orders of magnitude smaller than the corresponding RESI values. Thus, the contributions originating from Mb-U are drastically over-represented in the ESI-MS data of Figures 4a and 5. In other words, partial unfolding leads to a greatly enhanced ionization efficiency of the non-native Mb species. This finding is consistent with the behavior seen for lysozyme in the experiments discussed above; it is also supported by the results of ultracentrifugation54 and optical studies61 under conditions closely resembling those of Figure 4a. Once again, it is instructive to consider the amount of charge available for the conditions employed here. For the on-line HDX data of Figure 5 with Vf ) 50 µL min-1, qavg ) 14, and Cprot ) 15 µM it follows from eq 5 that (Cq ) 23 µM) < (Cprot × qavg ) 210 µM), corresponding to charge deficincy. As discussed above, this implies that the conformers that can most effectively compete for the available charge (Mb-U) will suppress ionization of other species (Mb-F). It would be interesting to investigate the effects of lower flow rates and protein concentrations on the appearance of the Mb mass spectrum under pulsed HDX conditions. Unfortunately, lower protein concentrations result in HDX data with poor S/N ratios. Lower flow rates result in longer labeling times, where interconversion of Mb-U and Mb-F is no longer negligible. The interpretation of data under those conditions is considerably more difficult.54 CONCLUSIONS The results of this work suggest that ESI-MS analysis of coexisting protein conformers in solution generally tends to result in higher signal intensities for more unfolded species. Ion intensity and actual concentration ratios can differ by as much as 2 orders of magnitude. It is proposed that this effect is due to the partially nonpolar character of unfolded proteins which enhances their affinity for the air/liquid interface of the early ESI droplets. From

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the parent droplet surface layer the proteins are transferred to small progeny droplets that eventually produce gas-phase ions. Natively folded proteins have a higher propensity to remain in the interior of the parent droplet, where all solvent-exposed hydrophilic side chains can favorably interact with water. As a result, the transfer of these folded species to ion-producing offspring droplets is not as efficient. In addition, the increased hydrophobicity of unfolded proteins may facilitate desolvation and, therefore, the transition from solution into the gas phase. The extent to which signals arising from natively folded proteins are suppressed depends on the experimental conditions. Low flow rates (