HYDROGEN EXCHANGE MASS SPECTROMETRY FOR THE ANALYSIS OF PROTEIN DYNAMICS

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1 HYDROGEN EXCHANGE MASS SPECTROMETRY FOR THE ANALYSIS OF PROTEIN DYNAMICS Thomas E. Wales and John R. Engen* Department of Chemistry, University of New Mexico, Albuquerque, New Mexico Received 5 April 2005; received (revised) 5 July 2005; accepted 6 July 2005 Published online 5 October 2005 in Wiley InterScience ( DOI /mas Hydrogen exchange coupled to mass spectrometry (MS) has become a valuable analytical tool for the study of protein dynamics. By combining information about protein dynamics with more classical functional data, a more thorough understanding of protein function can be obtained. In many cases, protein dynamics are directly related to specific protein functions such as conformational changes during enzyme activation or protein movements during binding. The method is made possible because labile backbone hydrogens in a protein will exchange with deuterium atoms when the protein is placed in a D 2 O solution. The subsequent increase in protein mass over time is measured with high-resolution MS. The location of the deuterium incorporation is determined by monitoring deuterium incorporation in peptic fragments that are produced after the labeling reaction. In this review, we will summarize the general principles of the method, discuss the latest variations on the experimental protocol that probe different types of protein movements, and review other recent work and improvements in the field. # 2005 Wiley Periodicals, Inc., Mass Spec Rev 25: , 2006 Keywords: deuterium; electrospray; MALDI; protein folding; conformation I. INTRODUCTION Proteins are not static structures in solution. They move and flex naturally or in response to external stimuli. The movements of proteins, collectively termed protein dynamics, can be extremely important for protein function. To ascertain how dynamics play a role in protein function, actual protein motions themselves need to be investigated and understood. Currently, there are not a large number of analytical tools capable of providing the necessary resolution to connect specific protein movements with protein function. The development of new tools and refinement of those that are currently in use is therefore of great interest. To this end, hydrogen exchange (HX) coupled with mass spectrometry (MS) presents an opportunity for the analysis of proteins and protein motions in ways that were not imagined even 5 years ago. Traditionally, hydrogen exchange methodology has been used in conjunction with NMR analysis [see (Dyson & Contract grant sponsor: The National Institutes of Health; Contract grant numbers: R01-GM070590, R01-GM068901, R24-CA088339, P20-RR *Correspondence to: John R. Engen, Clark Hall 242, MSC , Department of Chemistry, University of New Mexico, Albuquerque, NM engen@unm.edu Wright, 2004) for a review]. In comparison, hydrogen exchange MS is a more recent development. The first demonstrated use of HX MS came shortly after the development of electrospray ionization (Chowdhury, Katta, & Chait, 1990; Katta & Chait, 1991). Important developments through the 1990s were led by Prof. David Smith [reviewed in (Smith, Deng, & Zhang, 1997; Engen & Smith, 2001)]. Hydrogen exchange MS has been the subject of several comprehensive reviews (Kaltashov & Eyles, 2002a,b; Hoofnagle, Resing, & Ahn, 2003; Eyles & Kaltashov, 2004; Garcia, Pantazatos, & Villarreal, 2004). Recent developments in this field, which will be summarized in the following sections, offer unparalleled limits of detection, low sample consumption requirements, the promise of single amino acid resolution, potential for automation and the ability to analyze increasingly more complex mixtures. Future refinements that could substantially improve the method will also be discussed. II. OVERVIEW OF THE METHOD An general scheme for hydrogen exchange MS experiments is shown in Figure 1. The integration of deuterium into the protein(s) of interest relies on the natural phenomena of hydrogen exchange. The introduction of deuterium into a peptide or protein can be accomplished in several ways and its incorporation can be analyzed using several methods. The mass spectrometer is used to monitor the increase in mass as hydrogen is exchanged for deuterium. Throughout this review, we discuss several parts in the scheme (Fig. 1, numbered circles) where recent improvements have been made and where future refinements are anticipated. III. HYDROGEN EXCHANGE FUNDAMENTALS The details of hydrogen exchange mechanisms have been widely reviewed (Hvidt & Nielsen, 1966; Woodward, Simon, & Tüchsen, 1982; Englander & Kallenbach, 1984; Kim & Woodward, 1993; Mayo & Baldwin, 1993; Bai et al., 1995; Miller & Dill, 1995; Loh et al., 1996; Clarke, Itzhaki, & Fersht, 1997; Kaltashov & Eyles, 2002b; Hoofnagle, Resing, & Ahn, 2003; Eyles & Kaltashov, 2004; Krishna et al., 2004; Smith, Deng, & Zhang, 1997). Here a short summary, collected from these references, on the basics of hydrogen exchange and how it is applied to the study of protein dynamics is presented. Hydrogens that are located at peptide amide linkages (also referred to as the backbone amide hydrogens) undergo replacement with deuterons within 1 10 s when the peptides are incubated in D 2 O Mass Spectrometry Reviews, 2006, 25, # 2005 by Wiley Periodicals, Inc.

2 PROTEIN DYNAMICS BY HYDROGEN EXCHANGE MASS SPECTROMETRY & at pd 7.0. In folded proteins, some backbone amide hydrogens exchange quickly while others exchange only after months. The rates of the most slowly exchanging amide hydrogens may be reduced by as much as 10 8 of their rates in unfolded forms of the same protein (Englander & Kallenbach, 1984). Nearly all peptide amide hydrogens in folded proteins are hydrogen bonded, either intramolecularly to another part of the protein or to water. The large reduction of amide hydrogen exchange rates in folded proteins is primarily due to restricted access of solvent to the interior of the protein and to intramolecular hydrogen bonding. It is not possible to differentiate between these two contributing parameters as they occur concomitantly. At physiological ph, base-catalyzed exchange is the dominant mechanism for hydrogen exchange. Base-catalyzed isotope exchange can occur only when a hydrogen bond is severed in the presence of the catalyst (hydroxide) and the source of the new hydrogen (water). The rate constant for isotope exchange at each individual amide linkage in a normally folded protein, k ex, can be described by Equation 1 k ex ¼ k f þ k u ¼ðb þ K unf Þk 2 ð1þ where k ex is expressed as the sum of the contributions of exchange from folded (k f ) and unfolded (k u ) forms of the protein (Kim & Woodward, 1993). The mechanisms described by Equation 1 are illustrated graphically in Figure 2. Exchange from the folded form likely dominates for amide hydrogens that are not participating in intramolecular hydrogen bonding and that are located near the surface. Exchange in unfolded forms requires substantial movement of the backbone. Unfolding to expose backbone amide sites to deuterium can be isolated to small regions (localized unfolding) or may involve the entire protein (global unfolding). The rate constant for exchange from the folded state, k f,is described by Equation 2 k f ¼ bk 2 where b is a probability factor for exchange from folded forms and k 2 is the rate constant for HX at each amide linkage in an unstructured peptide, a value that can be calculated (Bai et al., 1993). The value for b ranges from 0 to 1 and is a function of several parameters including solvent accessibility and intramolecular hydrogen bonding. When b is closer to 1, there is a higher probability that a particular amide hydrogen is exposed to water and catalyst at the same time that it is also exchange competent. K unf in Equation 1 is the equilibrium constant describing the unfolding process. HX NMR studies have used denaturants to distinguish between b and K unf (Bai et al., 1994; Itzhaki, Neira, & Fersht, 1997; Chamberlain & Marqusee, 1998). The rate constant for exchange from unfolded forms of proteins depends on the rate constant for exchange from an unfolded peptide (k 2 ) as well as the unfolding dynamics described by k 1 and k 1 as shown in Equation 3 where F and U are the folded and unfolded forms, respectively. ð2þ F H Ð k1 U H! k2 U k 1 D Ð F D ð3þ k 1 D 2 O k 1 When k 2 >> k 1 (termed EX1 kinetics), the rate constant for exchange from unfolded forms is given by the unfolding rate constant k 1 (Eq. 4). k u ¼ k 1 ð4þ However, under physiological conditions it is more common for k 1 >> k 2. In this case (EX2 kinetics), the rate constant for exchange from unfolded forms is given by Equation 5 where K unf is the equilibrium constant describing the unfolding process. k u ¼ k 1 k 2 ¼ K unf k 2 ð5þ k 1 While only a few proteins undergo EX1 kinetics naturally, all proteins undergo EX2 exchange kinetics under physiological conditions. EX2 kinetics may be envisioned as involving many rapid and random visits to a state capable of exchange. However, the probability of exchange during a single visit is small. EX1 kinetics are described as a cooperative unfolding event involving several residues, all of which exchange before refolding (k 1 ) occurs (Eq. 3). Proteins can be induced to exhibit EX1 kinetics with denaturant (Deng & Smith, 1998) or by increasing ph (Swint-Kruse & Robertson, 1996). Some proteins may contain regions that undergo EX1 and EX2 kinetics simultaneously. Regions in which exchange occurs by either EX1 or EX2 kinetics can be identified by characteristic isotope patterns in mass spectra (Miranker et al., 1993). Structural changes required for HX described by k f and k u differ in the magnitude of atomic displacement(s) required for isotope exchange. Due to the highly compact nature of proteins in their native state, relative to their denatured states, exchange at individual sites is believed to involve small atomic movements, probably less than an angstrom, but sufficient to allow diffusion of OD and D 2 O to the exchange site (Kim & Woodward, 1993). In parallel with this highly localized motion, short segments, as well as the entire backbone of a protein, can exchange through unfolding processes. Molecular motions associated with unfolding of large segments of the backbone require displacing many atoms several angstroms from their equilibrium positions in the native structure and global unfolding requires gross movement of the entire backbone. Thus, exchange from the folded form (k f )of a protein involves primarily low amplitude motions (small displacement) while exchange from unfolded forms (k u ) requires much larger amplitude motions. Results of a theoretical study by Miller and Dill suggest that large structural changes with little changes in free energy are possible, but uncommon (Miller & Dill, 1995). IV. DEUTERIUM INTRODUCTION: DEUTERIUM, MEET PROTEIN; PROTEIN, MEET DEUTERIUM To measure the incorporation of deuterium into a protein, the protein must, of course, be exposed to deuterium. The introduction of deuterium sounds like a relatively simple process. However, different methods of deuterium introduction allow one to probe different aspects of protein dynamics. In addition, the sample quantity requirements of the mass spectrometer, 159

3 & WALES AND ENGEN especially in the case of analysis of protein complexes, may dictate D 2 O introduction of a specific type. The primary method for introducing deuterium into a protein sample is by dilution. Typically, a solution of protein in a protiated buffer is diluted with a deuterated buffer that has a deuterium content of 99% or more. Dilutions of 15-fold or greater will produce final deuterium concentrations of >95%. This serves to force the labeling reaction (k 2 ) in one direction (see Eq. 3). However, with this labeling method the original protein sample is diluted. Such a dilution may not be compatible with the sample quantity requirements of the mass spectrometer and such diluted samples may therefore require an additional experimental step to concentrate the protein sample prior to analysis. Concentration can be accomplished by rapidly trapping the protein with online HPLC at higher flow rates (usually >100 ml/min). The concentration step works best when analyses are performed with online HPLC-ESI as too much deuterium would be lost if this step were done prior to MALDI mass analyses (see below). An alternative to the dilution technique is to carry out a rapid buffer switch with small gel filtration spin columns (Engen & Smith, 2000). Although the buffer switch technique takes a bit longer than the dilution method, the protein is not nearly as diluted. There are two kinds of labeling experiments: continuous labeling and pulse labeling [see also (Deng, Pan, & Smith, 1999a)]. In continuous labeling experiments (Fig. 1, top right), protein is exposed to D 2 O while the populations of folded and unfolded species are in flux. The populations may be in flux as a result of natural protein motions that result from some population-altering force (i.e. the addition of denaturant, change in ph or temperature, in response to protein function, ligand binding, or protein protein complex formation, etc.) designed to cause a shift in the population of folded versus unfolded (or the reverse depending on the experiment). Once a protein has made a transition from a folded to unfolded state, it becomes labeled with D 2 O and the mass increases. As the D 2 O concentration is very high, once a molecule is labeled, it is not able to revert to a protiated species (see Eq. 3). In other words, the transition from a protiated species to one that is deuterated is unidirectional. The deuterium level in the protein sample at any point in the course of the labeling experiment integrates the number of molecules in the sample that had unfolded (or folded) up to that point (Miranker et al., 1993). With continuous labeling, it is possible to sample the population of and potentially observe the transition through various intermediate states whose numbers are very small at any given moment and therefore may go undetected using conventional spectroscopic methods. Continuous labeling is most useful for monitoring slow unfolding transitions [i.e. (Engen et al., 1997)], the majority of unfolding events in proteins. Given enough time, all proteins should become totally deuterated during a continuous labeling experiment as a result of protein motions and protein breathing. Because the transition is very slow for many proteins, the addition of denaturants may be used to induce unfolding. Most HX MS experiments involve continuous labeling simply because they are technically simpler to perform. Far fewer experiments are of the pulse labeling variety. Generally in pulse labeling experiments (Fig. 1, top left), a population of protein molecules is either induced to undergo some kind of conformational change by addition of a perturbing agent or it is already in the process of changing its structure through protein folding. The perturbing agent is most often a chemical denaturant, although heat, ph or binding to substrates can also be used. The sample is then exposed to deuterium for a very brief time (the pulse). Only those molecules that are unfolded when the sample is pulse labeled will undergo isotopic exchange; the remainder of the population remains unlabeled. The resulting deuterium levels then indicate the instantaneous population of folded and unfolded molecules. Pulsed labeling has recently been used to identify protein folding mechanisms [see (Wu & Engen, 2004) for a more detailed description] as well as to probe significantly populated kinetic intermediate states in a folding reaction (either on or off pathway intermediates) (Deng & Smith, 1999; Chen et al., 2001; Wintrode et al., 2003; Mazon et al., 2004; Pan & Smith, 2004; Pan et al., 2004; Rojsajjakul et al., 2004). With pulse labeling, it is FIGURE 1. Overall scheme for hydrogen exchange mass spectrometry experiments. A: Pulse labeling. After a protein has been exposed to a perturbant (chemical denaturant, heat, ph, binding, complex formation, pressure, etc.), unfolded regions (gray) become labeled with deuterium (red) during a quick pulse of D 2 O (typically 10 s). Deuterium exchange is quenched by reducing the ph and temperature. B: Continuous labeling. D 2 O buffer is added to a protein (in H 2 O buffer) such that the final D concentration is >95%. After a set period of time, an aliquot of the labeled protein is removed from the original tube and mixed with quench buffer to reduce the ph and temperature. Aliquot removal is repeated for subsequent labeling times. The protein concentration and solution volume are controlled such that all the aliquots are identical upon quench except for the amount of time the protein was exposed to D 2 O. C: Localized exchange information. Quenched samples (from part A, part B, or both) are digested with pepsin or another acid protease. The resulting peptides are analyzed with online HPLC-ESI-MS or with MALDI-MS. The resulting data analysis provides information on deuterium exchange in short fragments of the peptide backbone. D: Global exchange information. Quenched samples (from part A, part B, or both) are directly analyzed with HPLC-ESI-MS or MALDI-MS. The data provide a global picture of how the protein behaves in D 2 O. It is often recommended that Part D be performed prior to Part C. Areas of recent and future improvement have been marked with numbers. 1 Robotic automation of mixing, buffer addition, etc.; 2 Rapid-mixing techniques such as quench-flow analyses (see text for details); 3 Analysis based upon relative deuterium levels instead of absolute levels (see text for details); 4 Use of acid proteases other than porcine pepsin. Alternative proteases include protease type XIII from Aspergillus saitoi and protease type XVIII from Rhizhopus species (Cravello, Lascoux, & Forest, 2003). 5 CID, ECD, and ETD as fragmentation techniques to provide single amino acid resolution of hydrogen exchange information; 6 Solvent-free MALDI; 7 Nano-ESI-MS and other miniaturization; 8 chromatographic and labeling techniques to accomplish HX MS of single proteins in complexes and mixtures; 9 automation of data analysis and increase interpretation speed. 160

4 FIGURE

5 & WALES AND ENGEN FIGURE 2. Models for hydrogen exchange into the folded form (A) and into unfolded forms (B) of proteins. The unfolding and refolding rate constants are described by k 1 and k 1, respectively. k 2 is the rate constant for exchange from an unfolded peptide, a value that can be calculated (Bai et al., 1993). See text and Equation 3 for further details. possible to complement data obtained using conventional spectroscopic and NMR experiments (Nishimura, Wright, & Dyson, 2003). Pulse labeling experiments may also be performed using a quench-flow scheme originally described for HX MS by (Yang & Smith, 1997) and more recently discussed by Konerman & Simmons, 2003; Wintrode et al., The addition of deuterium and other steps in the preparation of pulse or continuous labeled samples has been automated (see section on Automation later in this review). With automation, the reproducibility of sample preparation is improved. It is anticipated that fully automated sample preparation devices will become commonplace (Fig. 1, circles 1,2). during the HPLC step and for MALDI analyses, losses may occur during the sample preparation process. However, an adjustment can be made to compensate for back-exchange. An adjustment calculation was first described by Zhang & Smith, In the appendix to their 1993 publication, they describe the derivation, accuracy, and proper use of the equation used for the calculation. While other adjustment methods have been described (Resing, Hoofnagle, & Ahn, 1999; Hoofnagle, Resing, & Ahn, 2004), they do not significantly improve upon this original comprehensive description. When properly controlled, the back-exchange in most modern and well tuned ESI mass spectrometers is on the order of 1 3%. An additional 10 20% of the deuterium label may be lost during in-solution digestion and HPLC separation depending on the length of time that it takes to perform each step. However, peptide and protein recovery differ widely [discussed in (Pan & Smith, 2004)]. More losses can be expected when MALDI is used for the analysis [see below and (Kipping & Schierhorn, 2003)]. However, even when 15% of the label reverts back to hydrogen, less than 1 in 6 deuterium is lost. The end result is that if two curves indicating changes in deuterium levels in a given protein or peptide are obtained and show a difference of 2 3 deuterium, a correction for back-exchange will only change the difference between the two curves by Da. Therefore the comparison of exchange curves with and without the correction yields almost no new information except for a closer approximation of the number of deuterium that were incorporated into the protein or peptide of interest. An example of this is shown in Figure 3. Here, the back-exchange correction was applied to raw data where D 2 O losses were abnormally high (35%). After correction for the V. IMPROVING MASS ANALYSIS A. Retaining the Label Once the incorporation of the isotope label is complete, the task becomes the identification of which amides have been deuterated. To maximize the amount of retained label, the deuterium back-exchange to hydrogen must be controlled. Back-exchange is the undesirable exchange of the deuterium for hydrogen and results in loss of some of the label. To minimize back-exchange, sample analysis must be as rapid as possible and be done at 08C. The majority of back-exchange occurs because proteolytic digestion and the subsequent analysis of the deuterium levels are done with protiated solvents. For ESI analyses, losses occur FIGURE 3. Back-exchange correction. A: Back-exchange adjustment equation as described by (Zhang & Smith, 1993). D is the adjusted deuterium level, m is the experimentally observed mass, m 0% is the 0% or undeuterated control, m 100% is the totally deuterated control, and N is the total number of exchangeable amide hydrogens in the sequence of interest. B: Example of the use of the back-exchange correction in part A. The average deuterium loss for this example peptide was 35%. Closed circles: raw data; Open squares: raw data adjusted using the equation in (A); Dashed line: the calculated amount (Bai et al., 1993) of deuterium in an unstructured peptide with the same sequence as the example peptide examined under identical conditions (ph 7.0 and 228C). 162

6 PROTEIN DYNAMICS BY HYDROGEN EXCHANGE MASS SPECTROMETRY & samples and all other variables related to back-exchange cancel out. FIGURE 4. Model peptide showing the fragmentation sites for b/y and c/z ions [see Roepstorff & Fohlman, 1984]. 35% D 2 O loss, the curve is shifted upward, with the most significant deviation from the raw, uncorrected data occurring at larger deuterium levels. This is an extreme example, as many peptides do not loose 35% of their label during a well controlled experiment. The back-exchange correction is only necessary when one wishes to know the exact number of deuterium atoms in a given protein or peptide fragment. In many cases where biological functional information is being probed, the actual number of deuterons that have exchanged is not as important as where the exchange has occurred. For example, if deuterium levels are being compared in peptic fragments of a folded and denatured version of the same protein, the location of the unfolding is of primary interest. The same protein is being analyzed, but under different experimental conditions. In such cases, a relative deuterium level can be used rather than making a back-exchange correction to obtain the absolute number of deuterons for each exchange time-point (Fig. 1, circle 3). There are several advantages of using relative levels rather than absolute deuterium levels. First, as proper back-exchange correction relies on analysis of a totally deuterated form of the molecule of interest, a totally deuterated form must be prepared. It can be difficult to prepare such a control, especially for larger proteins. As one is never really certain that the totally deuterated control sample is really 100% deuterated at all backbone amide positions, one can never really be sure of the validity of a correction that assumes 100% deuteration. Using relative deuterium levels does not require the preparation or analysis of a totally deuterated sample. Second, because back-exchange is a complex process that depends to some extent on the sequence (Zhang & Smith, 1993), recoveries differ widely between different proteins and peptides [discussed in (Pan & Smith, 2004)]. Sequence variation is no longer a factor when using the relative method because the same sequence is being compared to itself. The only variable that has changed is the conformation of the protein(s). An added benefit of relative results is that other variables such as slight changes in buffer ph, temperature, concentration, etc. all cancel out. Ordinarily these types of variables, in addition to back-exchange variability, are significant enough to prevent the use of relative deuterium levels to compare samples obtained at different times. To take advantage of the relative method, all experiments that one wishes to compare must be completed together under identical experimental conditions. For experiments performed at the same time and under identical conditions, back-exchange is statistically the same for all B. Localizing the Label Determination of deuterium levels in whole proteins does not provide the type of local information desired. In other words, the spatial resolution is very low. To increase the resolution, proteolytic digestion is primarily used [first described by Zhang & Smith, 1993]. Because digestion of the deuterium labeled protein(s) must occur under quench conditions, acid proteases must be used. To date, the best acid-protease for these purposes is pepsin. A few complications exist however. As pepsin is a nonspecific protease (it generally cleaves at hydrophobic residues), the sites of backbone cleavage cannot be predicted from the amino acid sequence. However, pepsin will cut in the same place given the same conditions, so reproducibility is not a problem. To be certain of the identity of each peptic peptide, it becomes necessary to sequence those that are generated. Most commonly, this is accomplished using tandem MS techniques. A further complication is that peptic digestion of large proteins may produce undesirably long peptides (>15 residues long) rather than the more desirable short ones (5 10 residues). Eric Forest and colleagues recently demonstrated the use of acid proteases other than pepsin for HX MS analyses (Cravello, Lascoux, & Forest, 2003) (Fig. 1, circle 4). However, pepsin was still the most efficient protease. The use of multiple enzymes produced many overlapping fragments. Overlapping peptides are highly desirable (Garcia, Pantazatos, & Villarreal, 2004) as they increase the spatial resolution. Multiple shorter peptides were also obtained from regions where pepsin cleavage produced single, long peptide fragments. A form of pepsin that cleaves with a greater specificity would significantly improve the digestion step for HX MS. Enzyme engineering should be able to generate recombinant analogues of pepsin that have increased specificity, as seen with trypsin or other specific proteases. In addition to the complications already mentioned, another significant problem with using pepsin that is often overlooked is the interpretation of small deuterium changes in the long peptic fragments. For example, a change of 2 deuterium in a peptide of 20 amino acids cannot be attributed to changes in dynamics within the whole peptide. While curve fitting (Zhang & Smith, 1993) serves to classify the hydrogens into categories, it cannot identify them. Careful data interpretation is necessary when large fragments are involved. A method to further improve the spatial resolution is needed. C. New Fragmentation Methods It would be most desirable to be able to monitor deuterium exchange at individual amino acids with MS. Attempts have been made to use CID to fragment peptic peptides into shorter pieces (b/y ions, see Figure 4) (Deng, Pan, & Smith, 1999b; Demmers et al., 2000, 2002; Kim et al., 2001; Hoerner et al., 2004). It was originally observed that b ions from high-energy CID with argon as the collision gas yielded deuterium levels that were consistent with NMR measured deuterium levels (Deng, Pan, & Smith, 1999b; Kim et al., 2001). However, deuterium was apparently scrambled in most y ions via migration during the fragmenta- 163

7 & WALES AND ENGEN tion process. The scrambling process seemed to depend on the sequence (Demmers et al., 2002) and other multiple factors (Hoerner et al., 2004). Other work has shown that scrambling appears to be 100% in both b and y ions (Jorgensen et al., 2005). Other methods of fragmentation may offer a solution to the scrambling issue. Kaltashov and Eyles have extensively discussed the use of FTMS and electron capture dissociation (ECD) methodology towards this end (Kaltashov & Eyles, 2002a,b; Eyles & Kaltashov, 2004). It has been shown that the fragmentation of ions in ECD occurs before there is an opportunity for the energy to be randomized to a more probable bond-cleavage site (Turecek & McLafferty, 1984; Zubarev, Kelleher, & McLafferty, 1998; Horn, Ge, & McLafferty, 2000). This rapid and effective activation method may therefore result in a decrease in the hydrogen scrambling as opposed to the slower CID activation of ions. A report on the use of ECD to fragment intact proteins after HX labeling has recently appeared (Charlebois, Patrie, & Kelleher, 2003). Fragments produced by ECD of whole proteins (c/z ions, see Figure 4) may still result in incomplete sequence coverage, especially for larger proteins. It may still be advisable to digest larger proteins and protein complexes with pepsin and perform ECD on the peptides, thereby providing single amino acid resolution (Fig. 1, circle 5). Other instrumental methods have also reported on fragmentation of proteins after HX (Akashi, Naito, & Takio, 1999; Lanman et al., 2003). Unfortunately, the ECD methods described to date are limited to Fourier transform instruments for technical reasons and are therefore cost prohibitive for many research laboratories. Recent descriptions of ECD and ETD (electron transfer dissociation) in ion traps may bring this technology into the hands of researchers using standard mass analyzers (Baba et al., 2004; Syka et al., 2004). ECD/ETD technology should facilitate the eventual analysis of deuterium exchange at individual amino acids without concern for potential label scrambling. Such experiments would be a major improvement to the HX MS method (Fig. 1, circle 5). VI. WHICH KIND OF MASS SPECTROMETRY? ESI VERSUS MALDI The use of ESI-MS for the analysis of hydrogen/deuterium exchange experiments has become very popular since the mid 1990s. Approximately 80% of the articles published with respect to HX MS in the past 5 years have employed electrospray as the ionization source. ESI-MS has become the most commonly used ionization method for HX MS partially because the quenched sample is introduced directly via HPLC into the electrospray source. The advantages of using electrospray MS are described below. MALDI-MS can also be used for HX MS protein dynamics studies. E. A. Komives reported the first use of MALDI for the analysis of hydrogen exchange content in the mapping of a protein protein interface (Mandell, Falick, & Komives, 1998a). She has used MALDI HX MS to identify several binding sites in the cyclic-amp-dependent protein kinase (PKA) complex with the kinase inhibitor and ATP (Mandell, Falick, & Komives, 1998a) and to determine the antibody antigen recognition site for a monoclonal antibody raised against human thrombin (Baerga-Ortiz et al., 2002). Others have reported on the use of MALDI to probe the folding and assembly of viral capsids (Tuma et al., 2001) and to investigate protein conformational changes as a result of polyvaline and polyleucine a-helix aggregation (Hosia, Johansson, & Griffiths, 2002). Topological information about yeast F1-ATPase, a supramolecular hetero-oligomeric protein complex of 370 kda, was obtained using MALDI as the ionization method (Nazabal et al., 2003). The advantages of MALDI are that it is, in principle and generally in practice, easier than ESI for non-experienced users. All ions are singly charged and this simplifies data interpretation. There is no HPLC separation and desalting. However, the disadvantages seem to outweigh the advantages: only 20 % of all HX MS publications have used MALDI as the ionization technique. Deuterium losses are significantly higher in MALDI than in ESI, although attempts have been made to reduce these loses (Kipping & Schierhorn, 2003). While all of the peptides are present in one spectrum and no HPLC steps are required (seemingly a simplification over ESI methods), for large and complex systems, the spectra become too crowded to interpret the data. Simple HPLC separation is necessary to temporally resolve a large number of peptides. Coupling HPLC to ESI has other advantages: the HPLC step washes away deuterium label present in the amino acid side-chains (Zhang & Smith, 1993), it is compatible with all buffers and denaturants, it can afford rapid concentration of very dilute samples (although ZipTips and the like can do some concentration in MALDI analyses) and ESI can handle some buffers and matrices that would be deleterious to MALDI sample ionization. The development of new solvent-free sample preparation methods (Trimpin et al., 2001, 2002) may eliminate some of the aforementioned complications for MALDI analysis of HX samples. Eluent from a chromatographic source can be continuously deposited onto a MALDI sample stage (that has been pre-coated with matrix) by spraying the eluent at elevated temperatures (Falkenhagen et al., 2003; Falkenhagen & Weidner, 2004). The advantages of this over the conventional dried droplet method and the solvent-free grinding method are improved compatibility of analyte and matrix as well as greater sensitivity. It remains to be seen if coupling of chromatography to solventfree MALDI spotting will make the use of MALDI more attractive for HX MS analyses (Fig. 1, circle 6). VII. THINKING SMALLER Although an obvious extension of the method, the use of nano- HX MS has not yet become widespread. Off-line nano-esi-ms analyses of continuous labeled samples have been used to probe lipid interactions with transmembrane peptides (Demmers et al., 2000). The a-helical structures in the molten globule state of wild type human a-lactalbumin and several proline analogues were also investigated with off-line nano-esi-ms (Last et al., 2001). Complete online digestion, separation and ESI-MS analysis on a nano scale have also been described (Wang & Smith, 2003) and a 100-fold increase in sensitivity was reported. Nano-HX MS can be particularly valuable for proteins that are hard to obtain in large quantities or those that aggregate at higher concentrations. 164

8 PROTEIN DYNAMICS BY HYDROGEN EXCHANGE MASS SPECTROMETRY & Many rare, disease-relevant signaling proteins fit into this category. It is anticipated that more and more proteins and protein systems will require the use of nano-hx MS in the coming years (Fig. 1, circle 7). VIII. MORE COMPLEX WITH COMPLEXES IN THE MIX While analysis of proteins in vitro is providing much information about them, the Holy Grail will be to analyze proteins in vivo. Further, the analysis of protein complexes and protein machines (Gavin et al., 2002) is becoming more and more commonplace. Techniques that can work with large protein assemblies and complex matrices such as cell lysates or organelle preparations should make this dream a reality. HX MS seems positioned to make this advance and with proper method development, it may soon be possible to investigate the dynamics of a single protein from a large, multi-protein complex or a single protein in a very complex mixture like cytoplasm. In vivo hydrogen exchange has already been reported (Ghaemmaghami & Oas, 2001). Other studies investigating proteins in cell lysates have also appeared (Engen, Bradbury, & Chen, 2002). The problem in all these experiments is the separation or isolation of the protein of interest from other cellular components. When such complex systems are involved, mass alone is not sufficient to distinguish the protein of interest from the components of the matrix. Systems in which protein fragmentation occurs after initial mass separation (ECD as described above, or several tandem stages of MS and HPLC) may help alleviate this problem. Other methods to get around the complexity problem include tagging the protein of interest so that it stands out against the background of the matrix (Engen, Bradbury, & Chen, 2002). These methods, although demonstrated in principle, have yet to be used practically. It would also work to isolate the protein by affinity purification just prior to mass analysis. However, the ph requirements for HX MS hinder this possibility, although other chromatographic methods (IEX) could potentially be better suited for this purpose. HX MS methods development is moving in this direction and future investigators will no doubt solve these technical problems (Fig. 1, circle 8). IX. AUTOMATION In an ideal automated system, an operator would select a time point for analysis and automated sample preparation and analysis instrumentation would do the rest. The end result would be a graph of hydrogen exchange for a given region of a protein, also automatically represented in a 3-dimensional modeling program. While the plumbing and sample introduction (Fig. 1, circles 1, 2) parts of this ideal scenario have been automated fairly easily (Woods, 1997, 2001a,b,c), the rest has not come so easily. Data processing at the end of data acquisition is substantial and mostly has been done manually. Automation of the data analysis [mentioned in (Garcia, Pantazatos, & Villarreal, 2004)] aids in processing but is not yet ideal. Commercial versions of the software are also not available. The coming years should see more development in this area (Fig. 1, circle 9). X. RECENT EXAMPLES OF WORK USING HXMS There are many examples of analyses performed with HX MS over the past 5 years. We would like to highlight several different types of analyses with specific examples from the literature to demonstrate the range of possibilities. A. Protein Assemblies MS has opened the door to investigating the dynamics of large protein assemblies. The HIV-1 capsid assembly is one such system that has been studied recently with HX MS (Lanman et al., 2003, 2004). HX MS experiments were performed on unassembled and assembled capsid tubes to determine the putative N domain to C domain interactions in the assembled capsid tubes. Interactions between helices I and II of the N domain were identified. In addition, a previously unrecognized inter-subunit N domain C domain interaction was observed (Lanman et al., 2003). Further experiments on both the immature and mature virion revealed that helix I and helix II were involved in intersubunit interactions in both the mature and immature virion (Lanman et al., 2004). Together, these results illustrate the use of HX MS methodologies to identify possible therapeutic targets that cause disruption of viral capsid assembly. Other groups have also investigated viral capsids with HX MS (Wang, Lane, & Smith, 2001). B. Protein Dynamics Conformational changes may occur as a result of phosphorylation or by means of single amino acid mutation. Two recent studies used HX MS to describe the effects of both phosphorylation and single amino acid mutation on the COOH-terminal Src kinase (Csk), an enzyme that regulates signaling by the Srcfamily of tyrosine kinases. To provide a structural framework for understanding phosphorylation-driven protein conformational changes, HX MS was used to monitor the effects of nucleotide binding on the solution conformation of Csk in the presence of ADP and AMPPNP (a non-hydrolysable ATP analogue) (Lanman et al., 2003). The results implied that phosphorylation of Csk results in conformational changes that may influence regulatory motions in the catalytic pathway. The HX MS data also showed that the conformational states of the protein are different depending on whether substrate or product is bound (Lanman et al., 2003). To probe the conformational consequences of a single amino acid substitution at amino acid position F183 in Csk, believed to be of significant importance in the communication between the SH2 and kinase domains of Csk, HX MS experiments for three substitution analogues at this specific position were performed. Three substitutions were explored: F! G, F! Y, F!A. HX MS experiments revealed that compared to the wild-type protein, glycine substitution at F183 reduced flexibility in several peptide regions in Csk, tyrosine mutations increased flexibility, and alanine mutations showed mixed results. In addition, the data suggested that each mutant was well folded, since major regions in all 3 domains had exchange patterns that were indistinguishable from those for the wild-type protein (Lanman et al., 2004). 165

9 & WALES AND ENGEN Another example of the use of HX MS to probe protein conformational dynamics involved the heat shock transcription factor s 32 (Rist et al., 2003). HX MS proved particularly valuable for this task as previous CD and fluorescence measurements were only able to detect global and local changes, respectively. As this protein tended to aggregate at high concentrations, NMR was not an available tool to study the protein solution conformation. HX MS experiments were designed to probe whether or not s 32 acted as a thermosensor. The folded states of s 32 were investigated at two separate temperatures: 37 and 428C, optimal growth conditions, and heat-stress conditions, respectively. The results indicated that there was a high degree of protein flexibility at 378C, and that there was reversible unfolding of a small structural motif at 428C. The location of this unfolded region was identified by analyzing the HX into pepsin fragments of the deuterium labeled samples. From these data, a map of the region that unfolded at elevated temperatures was produced. C. Protein Unfolding/Refolding Studies of protein folding/unfolding have traditionally been performed using the more conventional spectroscopic based techniques: NMR, IR, CD, and Fluorescence. Hydrogen exchange coupled to MS is also a useful tool for the study of protein folding/unfolding reactions. To illustrate this, some recent examples using pulse labeling HX MS are presented below. In studies of rabbit muscle triosephosphate isomerase (TIM) (Pan et al., 2004), pulse labeling denaturation experiments have revealed a bimodal isotope pattern of labeled protein in the raw MS data, which is evidence for two-state unfolding behavior. However, during the renaturation experiments there were three envelopes of isotope peaks present, suggesting that there was an intermediate in the refolding pathway. To obtain information about this intermediate, peptic digestion was performed after the pulse labeling experiment. The data showed that the intermediate was a form in which the C-terminal half was folded while the N- terminal half was not. Further, TIM folding fit a 4 þ 4 model of folding for (ba) 8 -proteins. In this set of experiments, HX MS provided location information regarding the folding intermediates as well as following both the refolding and unfolding properties. In another example, pulse labeling HX MS was employed to better characterize the molten globule intermediate state during the unfolding of the multidomain dimeric protein MM-CK (Mazon et al., 2004). In 0.8 M GdmCl, where the molten globule state of this protein was maximally populated, MM-CK exhibited a highly fluctuating structure that allowed for total deuteration. The study also used intrinsic fluorescence, ANS binding, and far- UV CD to probe the molten globule state of the protein. With HX MS, only two species were detected during GdmCl denaturation as opposed to more intermediate states detected by the conventional methods. The a-subunit of tryptophan synthase is a 29 kda, single domain protein that unfolds according to a four-state equilibrium unfolding model. There are two intermediate states, I1 and I2. The intermediate state I1 contains a significant amount of secondary structure while I2 contains no detectable secondary structure and mostly resembles the denatured state. Both of these intermediates are on-pathway kinetic species. HX MS was used to map the stable secondary structure in the I1 equilibrium intermediate (Rojsajjakul et al., 2004). Such information provided insights into the relationship between sequence, structure, and folding in the a-subunit. The identification of protected regions in the I1 intermediate was accomplished. The identified regions (most of the N-terminal region with 5 peptides regions of exception three of which are exposed loops) represented a contiguous domain (a-helices 1 3 and b-strands 1 4). It was also shown that the C-terminus, residues , was either unfolded or weakly folded in this intermediate. A refolding study of the urea denatured a-subunit of tryptophan synthase using pulse-quench HX MS has shown that there is an on pathway kinetic folding intermediate that shares a similar folded protein core with the I1 equilibrium intermediate (Wintrode et al., 2005). Together the data obtained for the equilibrium and kinetic intermediates show that the latter stages of the folding reaction for the a-subunit of tryptophan synthase are under thermodynamic control (Wintrode et al., 2005). When studying protein folding/unfolding reactions, it is not uncommon to see HX MS technologies combined with other methods. One such recent example is the integration of HX MS with a cyanylation-based methodology (Li et al., 2004) to better understand the conformation of disulfide-bonded proteins and intermediates during refolding reactions. In the cyanylationbased methodology, a protein containing disulfide bonds is trapped and the disulfide structure of a given cystinyl protein folding intermediate is identified and preserved. HX MS is then used to assess the other conformational features of the intermediate. These technologies were used to trap a 1-disulfide bond (early folding) intermediate and a 2-disulfide bond (later forming) intermediate of long Arg 3 insulin-like growth factor-i (LR 3 IGF-I) (Li et al., 2004). HX MS data showed an increasing degree of protection from exchange as a function of disulfide bond formation. There was significantly more secondary structure after formation of the second disulfide bond (specifically in helix 3). It is clear that HX MS technology is even more valuable when combined with conventional methods for the analysis of protein folding and unfolding reactions. As spectroscopic methods such as CD, IR, and Raman spectroscopy monitor globally averaged changes to protein secondary structures, and fluorescence monitors the exposure of aromatic residues to solvent, there is the possibility that they might not be sensitive to specific, local changes in protein conformation and dynamics. One can see from the above examples that HX MS can be an important complement to the traditional biophysical techniques to probe protein folding/unfolding reactions. HX MS may reveal a wealth of additional information that would otherwise go undetected. D. Binding Experiments As a result of their significance to protein folding, stability, association and function, binding interactions are under intense investigation. These interactions include protein:small molecule, protein:polypeptide, protein:lipid, protein:nucleic acid, and 166

10 PROTEIN DYNAMICS BY HYDROGEN EXCHANGE MASS SPECTROMETRY & protein:protein binding. Much information about the conformational changes that a protein undergoes during ligand binding have been determined with high-resolution X-ray crystallography or NMR structural analyses. HX MS has been used to probe the location(s) of binding sites of specific ligands on target proteins for which X-ray crystallographic and NMR methods are not applicable. Interpretation of these data, however, must be done with caution as binding is known to cause changes in protein dynamics and conformation at sites distant from the actual binding site or interface. In the discussion below there are a few examples of this observable fact [see also ref. (Mayne et al., 1992)]. An label and chase method that tries to compensate for this problem has been described (Garcia, Pantazatos, & Villarreal, 2004). There are many examples in recent years of HX MS being used in conjunction with high resolution structures to explore the organization and dynamics of complex molecular assemblies. The 42 kda eukaryotic protein actin, for example, has been investigated with HX MS (Chik & Schriemer, 2003). G-actin, F- actin (formed by polymerization of monomeric G-actin), F-actin bound to phalloidin, and a DNaseI:G-actin complex were studied to provide further information about the structure of F-actin and the structural effects of phalloidin and DNaseI ligand interactions. Note that the size and nature of polymerized actin (F-actin) excludes use of X-ray diffraction and NMR. HX MS results suggested a conformational transition from a closed to an open state of actin. The changes were localized to the phosphate binding loops upon polymerization of G-actin. Additionally, phalloidin binding to F-actin induced the monomer conformation to adopt a more G-actin-like state with the phosphate groups excluded from solution, a conformational change that inhibited phosphate release thereby reducing the rate of monomer dissociation. HX MS also provided evidence for conformational changes that occurred away from the DNaseI binding site of G-actin. These distal changes indicated a possible alteration of conformational flexibility consistent with previously published data in which the C-terminal residues were found more accessible to trypsin digestion. A study aimed at identifying regions in MAP kinases specific for binding two peptide docking motifs (DEJL and DEF) included the presentation of the distal effects of these ligand interactions (Lee et al., 2004). The data revealed that DEJL motif interactions with p38a MAPK induced enhanced backbone flexibility in the activation lip, an effect that was shown to be conserved between different MAP kinases. There were backbone conformational changes away from the region of the DEF motif interaction with ERK2 kinase. However, it remained unknown whether the interactions were attributable solely to allosteric effects or occurred as a result of alternate binding sites or nonspecific binding of the synthetic peptide to the kinase. Protein binding to small molecules is of great interest especially for development of small-molecule therapeutics. A recent HX MS study illustrating this type of experiment involved the binding of ATP to a-crystallin (Hasan, Smith, & Smith, 2002). ATP decreased the accessibility of amide hydrogens in multiple regions of both aa and ab subunits of a-crystallin. Four regions of a-crystallin, two in aa and two in ab, showed a significant decrease in the uptake of deuterium. Location information from HX MS data allowed for the comparison of the regions affected by ATP binding with proposed substrate binding sites. The authors concluded that ATP binding releases substrate by means of both direct displacement and a global conformational change. Recent HX MS binding studies between papain (target enzyme) and cystatin (thiol protease inhibitor) demonstrated that enzyme inhibitor interactions can be characterized by HX MS coupled to CID in a hexapole ion-guide using ESI-FTICR MS (Akashi & Takio, 2000). Binding of cystatin to papain reduced the flexibility throughout the papain molecule, data that are consistent with previous structural studies. HX MS protein:protein binding studies with the complex of UBC9 and SUMO-1 illustrated several key issues about protein binding experiments by HX MS (Engen, 2003). First, HXMS titrations can be used to estimate the K d for complexes. Further, backbone amide hydrogen exchange may not be altered in all proteins during complex formation, especially if the protein complex is formed primarily via electrostatic side-chain interactions. Finally, by combining site-directed mutagenesis with HX MS, much more information about the nature of complexes can be obtained. XI. LOOKING AHEAD While HX MS has come a long way in the past years, many more improvements can be made. It has only recently been widely realized that this technique can provide valuable information about proteins and protein dynamics that cannot be easily obtained with other techniques. Improvements in the method, as suggested by the circles in Figure 1, will only increase its power. We hope that many new researchers will realize this and join in the analysis of all proteins with HX MS technologies. ACKNOWLEDGMENTS We gratefully acknowledge support for this work from the National Institutes of Health (R01-GM070590, R01-GM068901, R24-CA088339, and P20-RR016480). 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13 & WALES AND ENGEN Woods VL, Jr. 2001b. Methods for identifying hot-spot residues of binding proteins and small compounds that bind to the same. In. USA: Carta Proteomics, Inc. US patent 6,599,707. Woods VL, Jr. 2001c. Method for characterization of the fine structure of protein binding sites. In. USA: Carta Proteomics, Inc. US patent 6,331,400. Woodward C, Simon I, Tüchsen E Hydrogen exchange and the dynamic structure of proteins. Mol Cell Biochem 48: Wu Y, Engen JR What mass spectrometry can reveal about protein function. Analyst 129: Yang H, Smith DL Kinetics of cytochrome c folding examined by hydrogen exchange and mass spectrometry. Biochemistry 36: Zhang Z, Smith DL Determination of amide hydrogen exchange by mass spectrometry: A new tool for protein structure elucidation. Protein Sci 2: Zubarev RA, Kelleher NL, McLafferty FW Electron capture dissociation of multiply charged protein cations. A nonergodic process. J Am Chem Soc 120: Dr. Thomas E. Wales received his B.A. degree in Chemistry and Spanish (1998) from Assumption College and his Ph.D. in Analytical Chemistry (2003) from Duke University. He is currently a postdoctoral fellow with John R. Engen at the University of New Mexico. His research has focused on the analysis of the protein backbone from both a chemical and structural perspective. Currently he is investigating the minute time-scale unfolding of protein backbone motions using hydrogen-deuterium exchange mass spectrometry. Dr. John R. Engen received B.S. degrees in Molecular Biology (1994) and Biochemistry (1995) from Union College and his Ph.D. in Analytical Chemistry (1999) from the University of Nebraska-Lincoln. As an EMBO Fellow, he did postdoctoral work (2000) in molecular biology and cellular signaling at the European Molecular Biology Laboratory in Heidelberg, Germany followed by a second postdoctoral appointment (2001) at Los Alamos National Laboratory. He was appointed an Assistant Professor of Chemistry at the University of New Mexico in January His research centers around the analysis of protein function with mass spectrometry, specifically hydrogen exchange analysis of structural activation in oncogenic kinases and protein folding during cellular processes. 170