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2 2 5 6 A A N A LY T I C A L C H E M I S T R Y / M AY 1,

3 A powerful new approach that goes John R. Engen David L. Smith University of Nebras - Thorough characterization of a protein requires understanding the interplay of three separate but linked features function, structure, and dynamics. Investigating these features demands a wide range of analytical techniques. For example, characterizing protein function, which is the identification of a protein s role in cellular maintenance, is usually more amenable to biochemical, molecular, and cellular types of analyses. On the other hand, studies of protein structure and dynamics most often use physical methods, many of which are common to analytical chemistry. HAMID GHANADAN/GH MULTIMEDIA 2001 M AY 1, / A N A LY T I C A L C H E M I S T R Y A

4 (a) (b) (c) H H H H H H CON CH CON CH CON CH CON CH CON CH CON H CH 2 CONH 2 CH 2 COOH CH 2 OH It was recognized nearly 50 years ago that different protein structures undergo isotope exchange at different rates, and this has led to widespread use of hydrogen exchange as a means to study protein structure and dynamics. Hydrogen exchange has been used to follow protein folding, detect structural changes due to mutation, and study protein protein interactions. Isotope exchange studies have most often used tritium or deuterium with analysis by nuclear magnetic resonance (NMR), IR, or UV spectroscopies. Over the past decade, hydrogen/deuterium (H/D) exchange CH 2 SH (CH 2 ) 4 NH 2 Asn Asp Ser Cys Lys H D 2 O +OD k 1 k 1 N H U H U D k k 1 2 k 1 OD + HOD FIGURE 1. Amide hydrogen exchange in peptides and proteins. (a) Hydrogens used for exchange studies. (b) Protein unfolding dynamics associated with isotope exchange. (c) Model used to link isotope exchange with protein unfolding dynamics. D D N D has also been detected by MS. Combining amide hydrogen exchange (HX) with MS has opened new possibilities for studying protein structure and dynamics. This article discusses models for interpreting HX results, describes studies of the structures and dynamics of small and large proteins, and summarizes the relative merits of NMR and MS for detecting H/D exchange. Structure and dynamics Determining the locations of all atoms in a protein to within 2 3 Å (the average structure of a protein) is a milestone, because this information is intimately linked to a protein s function. For example, with a high-resolution structure in hand, rational drug design can begin. Physical methods used to determine the three-dimensional structures of proteins can be classified according to their resolution. X-ray crystallography and high-resolution multidimensional NMR provide the highest resolution at <2 3 Å. Medium-resolution techniques, which include electron microscopy and neutron diffraction, provide structural data with resolutions of ~5 20 Å. Lower resolution is found with techniques such as circular dichroism (CD) and fluorescence spectroscopy. However, CD can measure the -helix and -sheet content of a protein, whereas fluorescence gives information on the environments of aromatic moieties, such as tryp tophan and tyrosine residues, or covalently attached probes like rhodamine. Low-resolution information can also be deduced from analytical ultracentrifugation, differential binding, chromatography, IR spectroscopy, and reactivities of amino acid side chains. Despite the importance of high-resolution structural information, it is only the beginning of the journey that leads to protein characterization. Proteins may have ,000 atoms, which are joined covalently and arranged in highly compact structures in the native state. However, even under conditions that strongly favor the native state, proteins exhibit highly dynamic behavior, often populating higher-energy states according to the Boltzmann distribution. Examples of dynamics include natural protein breathing or flexing in solution, structural changes resulting from interactions within the protein or with its substrates, and responses to ligand binding and changing solution conditions. Extreme examples of these higher-energy protein forms include partially and completely unfolded structures. Determining which regions of a protein change structure, the rate constants for these changes, and the free energies of different structures are some of the most important and fundamental goals of studying protein dynamics. Although our database of protein structures has grown significantly in recent years, information on protein dynamics is still sparse. Because structure and dynamics are closely intertwined, studies of protein dynamics usually involve detecting populations of different structures. For example, various quench-flow CD and A A N A LY T I C A L C H E M I S T R Y / M AY 1,

5 fluorescence methods have been used to detect structural changes associated with changing ph, adding denaturants, or binding ligands. Although several experimental methods have contributed to our understanding of protein dynamics, amide HX has played a particularly important role (1, 2). It will be evident from the following discussion that detecting HX by MS is a powerful new approach for investigating protein structure and dynamics. Fragment the labeled aldolase with pepsin HPLC/MS of digest Native aldolase Destabilize with urea N l 1 l 2 U Pulse label with D 2 O (pd = 6.8, 0 C) Quench H/D exchange with acid (pd = 2.5, 0 C) HPLC/MS of the intact aldolase Some background Of the many different kinds of hydrogens in proteins, only hydrogens located at peptide amide linkages (red in Figure 1) have exchange rates in a range that can be easily measured by MS. Because every residue except proline has one hydrogen at its peptide amide linkage, this class of hydrogens provides a series of discrete sensors along the entire length of the polypeptide chain. Isotope exchange rates for amide hydrogens depend primarily on the amide s intramolecular hydrogen bonding and access to the solvent (3). Thus, amide hydrogens located in -helixes, -sheets, and the hydrophobic core of a protein (where access to solvent is limited) exchange slowly. This slow exchange differs from that of amide hydrogens located in, for example, floppy loops on the protein surface. These amide hydrogens are hydrogen-bonded only to water, and they have good access to solvent; therefore, they exchange much more rapidly, sometimes by a factor of 10 8! The general mechanism of isotope exchange in folded proteins, illustrated in Figure 1b, requires abstraction of the amide proton by base followed by reprotonation/deuteration of the amide nitrogen with a proton/deuteron from the solvent. Exchange of slowly exchanging hydrogens occurs most readily following structural changes that free these hydrogens from intramolecular hydrogen bonding and provide access to the aqueous solvent (4, 5). These structural changes, which may be of low amplitude and involve only a few atoms, facilitate exchange at a single peptide amide linkage. This form of HX occurs while most of the protein remains folded. Alternatively, these structural changes may involve movements of large segments of the polypeptide backbone or even complete unfolding of the protein. This form of HX can be viewed as exchange from unfolded (or partially unfolded) forms of a protein (6). Some of the protein dynamics that can be studied by HX are described by the unfolding and refolding rate constants k 1 and Location of unfolding domains Peptides m/z Intact protein k 1, which are described in Figure 1c. The rate constant for isotope exchange from the unfolded polypeptide, k 2, depends primarily on the ph, temperature, and the nature of the amino acid side chains flanking the peptide amide linkage (7 ). The value of k 2 can be calculated using results obtained for short model peptides. Under physiological conditions, most proteins are stable (k 1 << k 1 ) and k 2 << k 1. In such cases, individual molecules must unfold and refold many times before isotope exchange at all amide linkages is complete. The rate constant for isotope exchange (k ex ) in this case is k ex = k f + k u = ( + K unf )k 2 (1) in which k ex is expressed as the sum of the contributions of exchange from folded (k f ) and unfolded (k u ) forms of the protein (4, 5, 8). The term EX2 has often been used to designate this form of HX kinetics. Equation 1 can be expanded to express k ex as a function of, K unf, and k 2, in which is the probability for exchange from folded forms and K unf is the equilibrium con N l 1 l2 U m/z Populations of N, l 1, l 2, and U Kinetic modeling Free energies and kinetics of N, l 1, l 2, and U Size of unfolding domains FIGURE 2. General strategy used to study the unfolding dynamics of a large protein, rabbit muscle aldolase, by HX MS. (Adapted with permission from Ref. 15.) N represents native protein, I 1 and I 2 are partially unfolded intermediates, and U is the unfolded form of aldolase. M AY 1, / A N A LY T I C A L C H E M I S T R Y A

6 stant describing the unfolding processes. Whether isotope exchange occurs from the folded or momentarily unfolded forms of a protein, Equation 1 links the experimentally measurable HX rate constant to the dynamics of a protein. Under some conditions (e.g., presence of denaturants or high ph), the rate of exchange from the unfolded polypeptide may be much larger than the refolding rate (i.e., k 2 >> k 1 ). In this case, isotope exchange is complete the first time a segment of a polypeptide backbone unfolds. When this form of HX kinetics prevails, the isotope exchange rate is equal to the unfolding rate (i.e., k ex = k 1 ). The signature for this form of HX kinetics, often referred to as EX1 kinetics, is a bimodal distribution of deuterium among the sample molecules. The ability to easily detect EX1 kinetics has been a particularly important contribution of MS to HX methodology (9 11). Unfolding the large protein aldolase Amide HX has proven to be an excellent tool for studying protein dynamics because H/D exchange in unfolded forms is often several orders of magnitude faster than exchange in the lowestenergy native form. Partially unfolded forms of several small proteins have been identified through elegant HX NMR studies (6, 12). Recent publications from our laboratory demonstrate that using MS instead of NMR extends these measurements to large proteins (13 16). Direct comparisons between NMR and MS for detecting amide HX have been reported (11, 17, 18). In both experiments, denaturants (e.g., guanidine hydrochloride or urea) were used to decrease the free energies and hence increase the populations of unfolded forms. Unfolding domains. The procedure used to study the unfolding dynamics of rabbit muscle aldolase is illustrated in Figure 2. Native aldolase was incubated in H 2 O/urea for times ranging from seconds to hours. The protein was then exposed to D 2 O (same urea concentration and ph) for 10 s, which was sufficient time for complete exchange of all amide linkage hydrogens in unfolded regions of the backbone (7, 17 ). To quench the isotope exchange reaction, the ph and temperature were both decreased. These simple changes decrease the intrinsic rate of H/D exchange (i.e., k 2 ) by ~5 orders of magnitude (7 ). Under quench conditions, the half-life for exchange in unfolded polypeptides is ~1 h. Several features of the unfolding dynamics can be discerned when the labeled and quenched protein is analyzed intact (i.e., as molecular ions) by reversed-phased HPLC electrospray ionization (ESI) MS. Mass spectra of aldolase incubated in 3.5 M urea for 30 min (blue in Figure 2) revealed four peaks, indicating four structural forms during the 10-s labeling period. The centroid of each peak is the average molecular mass of a specific structural form. When used with the appropriate standards (19), these molecular masses give the average number of deuteriums at peptide amide linkages in each structural form. This analysis showed that the peaks labeled N, I 1, I 2, and U have 0, 107 ± 5, 215 ± 5, and 327 ± 5 excess deuterium, respectively. Assuming that each excess deuterium corresponds to one unfolded residue (excluding proline), these results gave the number of unfolded residues in each structural form. It follows that the number of peaks in the ESI mass spectrum and the average molecular masses of the ions represented by these peaks give the number and size of each structural form. The relative intensities of these peaks provide the populations of each structural form. Finding two intermediates in the unfolding of aldolase was surprising because, from a structural point of view, this protein has only one domain (20). X-ray crystallography of aldolase showed that it occurs as a homotetramer (M r = 157 kda), in which the four subunits are joined by noncovalent forces. However, under the acidic conditions used to quench HX, the tetra - mer dissociates into monomers. Therefore, mass spectra of the intact protein (blue in Figure 2) were obtained for the 38-kDa monomers. Rabbit muscle aldolase subunits have 363 residues. To understand the unfolding dynamics of aldolase, we must also know which regions are unfolded in each of the intermediate forms, I 1 and I 2. Unfolded regions in proteins briefly exposed to D 2 O can be identified by locating deuterium along the polypeptide backbone. Although collision-induced dissociation MS/MS shows promise for such measurements (21, 22), we have most often used enzymatic cleavage followed by HPLC/MS (19). Pepsin, an aggressive protease with high activity under conditions of slow H/D exchange (i.e., ph 2.5, 0 C), was used to fragment the labeled protein into peptides. The deuterium levels in the peptides were determined by HPLC/ESI-MS. The ESI mass spectrum of the peptic fragment (including residues ), taken from intact aldolase that was pulse-labeled following equilibration in 3.5 M urea (red in Figure 2), has two envelopes of isotope peaks. The intensities of isotope peaks in the low-mass envelope (left side of the spectrum) corresponded closely to the natural abundance of heavy isotopes ( 13 C, 18 O, etc.) in the protein before labeling. The average molecular mass of ions comprising the high-mass envelope of isotope peaks (right side) corresponded to the peptide with all the amide linkages deuterated. It follows that the backbone region, which included residues , was completely folded (i.e., no deuterium) in some molecules and completely unfolded (i.e., completely deuterated) in others. Choosing the optimal labeling time and ph is an important step in designing effective HX pulse labeling experiments (23). If labeling is inadequate (i.e., short time, low ph), unfolded regions will not be completely deuterated. If labeling is excessive, A A N A LY T I C A L C H E M I S T R Y / M AY 1,

7 folded regions may become partially deuterated. In either case, the difference in deuterium levels found in folded and unfolded forms decreases, making it more difficult to quantify the two envelopes. Furthermore, some folded protein regions (e.g., extended loops) may undergo exchange at rates comparable with the same segment in an unfolded protein. Analysis of the mass spectra of 27 peptic fragments (13) has been used to locate specific regions of the backbone that were unfolded (show no protection to HX) for the two unfolding intermediates, I 1 and I 2. These results are summarized in Figure 3a, in which red identifies regions that unfolded quickly, blue marks regions that unfolded slowly, and yellow designates regions that unfolded at intermediate rates. It is important to note that all of the regions highlighted in a particular color unfolded at the same rate, consistent with the highly cooperative motion expected for protein unfolding reactions. Three short segments in black did not undergo isotope exchange, indicating that they did not unfold. These segments, located in the subunit binding surface (20), failed to exchange because aldolase retains its quaternary structure even when equilibrated in 3.5 M urea (14). Kinetics and thermodynamics. The relative intensities of isotope peak envelopes found in the mass spectra of peptic fragments can be used to determine the fraction of each domain that is unfolded (9, 11). Results from time course studies can be used to determine each domain s unfolding rate. However, this approach does not take into account the folding status of the other domains in the same molecule. We illustrate the problem in Figure 3b using triangles in which each side represents the folding status of a particular domain (red, yellow, or blue). Wavy lines indicate that the domain is folded; straight lines mark an unfolded domain. Analysis of labeled, intact aldolase following incubation in 3 M urea (blue in Figure 2) shows that there are at least four structural forms. These forms are attributed to the four structures illustrated schematically in Figure 3b. The native form has all three domains folded; the intermediates I 1 and I 2 have only the red or the red and yellow domains unfolded, respectively; and the unfolded form has all three domains red, yellow, and blue unfolded. Thus, the folding status of each domain can be related to the specific molecular forms N, I 1, I 2, and U. The populations of these four forms of aldolase destabilized in urea can be determined from the relative intensities of the four envelopes of isotope peaks found in the mass spectra of intact aldolase. Alternatively, they can be determined by a more complicated analysis of the isotope patterns in the mass spectra of peptic fragments (14). Representative results obtained after incubating aldolase in 3.5 M urea for 1 min to 48 h, followed by 10-s pulse labeling, are presented in Figure 3c. These results show that the population of native aldolase decreased rapidly during the first hour, reaching an equilibrium value of ~20%. During this time, the popula- (a) (b) (c) Percent folded N k 1 k 1 k 2 Subunit B k 3 Subunit D k 2 k 3 l 1 l 2 U 3.5 M urea Time (min) FIGURE 3. Unfolding dynamics of aldolase. (a) Ribbon structure of aldolase monomer illustrating the three unfolding domains: fast (red), intermediate (yellow), and slow (blue). Black indicates three short segments, located in the subunit binding surface, which are particularly resistant to unfolding. (b) Model used to link the unfolding of domains to structures proposed for unfolding intermediates. Folded and unfolded domains are indicated by wavy and straight lines, respectively. (c) Populations of variously unfolded forms, which were deduced from bimodal isotope patterns (Figure 2), were used with kinetic modeling to determine rate and equilibrium constants that describe the unfolding dynamics. Colored curves correspond to N (black), I 1 (red), I 2 (green), and U (blue). (Adapted with permission from Ref. 14.)

8 (a) Percent folded (c) 3 min tions of unfolding intermediates I 1 and I 2 passed through maxima, then decreased to equilibrium values. After ~15 h, the population of the unfolded form of aldolase rose to an equilibrium value of ~55%. Thus, the populations of these structural forms (b) SH3 SH2 15 min 8 hr m/z SH2-kinase linker 3 min 12 min 45 min Kinase domain FIGURE 4. ESI mass spectra of Hck SH3 domain and a SH3 domain peptic fragment. (a) Mass spectra of intact human Hck SH3 domain incubated in D 2 O. The incompletely resolved experimental data were fitted with two Gaussian functions. The fraction of molecules that remained folded during the labeling time is given by the area of the low-mass envelope (red), whereas the fraction of molecules that unfolded during the labeling period is indicated by the high-mass envelope (blue). (b) Mass spectra of the peptic fragment with residues (c) The unfolding regions located in intact Hck SH3 (33). Two segments of Hck SH3 participate in local unfolding and are indicated as red and yellow. (Adapted from Ref. 24.) can be determined as the system approaches equilibrium. The populations of the variously unfolded forms of aldolase can be used to investigate the kinetics and thermodynamics of its unfolding, pending selection of a specific unfolding mechanism (14, 15). For example, each of the three unfolded forms may be generated in a single, cooperative step. In this case, I 1 would be formed when only the red domain unfolded, I 2 when the red and yellow domains unfolded simultaneously, and U when all three domains unfolded simultaneously. Alternatively, the unfolding domains in aldolase may unfold sequentially. In this model, the yellow domain could unfold only after the red domain had unfolded, and the blue domain only after the red and yellow domains had unfolded. Modeling of these two possibilities suggests that aldolase unfolds sequentially (14). Fitting the experimental populations of N, I 1, I 2, and U to populations predicted for the models not only points to a particular unfolding mechanism but also gives rate constants for each step in the unfolding and refolding of the protein (15). The free energies of each unfolded form, relative to the native form, were determined using data from two different time zones. When equilibrium populations of unfolded forms had been established (t > 15 h), the equilibrium constants for each step of the unfolding process were determined directly from the populations of N, I 1, I 2, and U. Alternatively, the equilibrium constants were determined from the ratios of unfolding and refolding rate constants as determined by modeling. Estimated free energies for the same unfolding processes using these two approaches differed by <0.15 kcal/mol. Structure and dynamics of small proteins Slow unfolding of the Hck SH3 domain. HX MS has been used to investigate the dynamics of the regulatory domains in the tyrosine kinase Hck domain (24 26). Within signal transduction pathways, many interactions are facilitated by two small protein modules, the Src homology 3 (SH3) and Src homology 2 (SH2) domains, first discovered in the Rous sarcoma virus oncogene src (27 29). The properties of these domains have been of particular interest because of their roles in controlling kinase activity. SH2 domains typically consist of ~100 residues and bind A A N A LY T I C A L C H E M I S T R Y / M AY 1,

9 sequences containing phosphotyrosine. SH3 domains have ~60 residues, which form five -strands that create a hydrophobic binding surface for proline-rich sequences. We have probed various constructs of Hck SH2 and SH3 domains to look for changes in dynamics that may occur during activation or facilitate regulation of the kinase domain (24 26). Although the SH3 and SH2 domains are small enough to be analyzed by high-resolution HX NMR, HX MS studies can provide information outside of NMR s detection range. For example, HX MS studies of Hck SH3 led to the discovery of a particularly unusual unfolding process (24). The SH3 domain was labeled at ph 7 and 25 C using a procedure somewhat different from that illustrated in Figure 2. Pulse labeling was replaced with continuous labeling, and no denaturants were used to destabilize the protein. Mass spectra of the intact SH3 domains following incubation in D 2 O for 15 min exhibited two envelopes of isotope peaks (Figure 4a), which indicate a bimodal distribution of deuterium in the SH3 molecules. Although bimodal isotope patterns are common in proteins destabilized in denaturants (Figure 2), they are uncommon for proteins under native conditions. The bimodal distribution of deuterium was attributed to partial unfolding of SH3 during which the unfolded region remained open long enough for all of the amide hydrogens to be replaced with deuterium; that is, k 2 >> k 1 (Figure 1). Deconvoluting the two envelopes provided the date for determining the populations of molecules that unfolded during the labeling time (blue in Figure 4a). The rate constant for unfolding, calculated from the change in this population with time, showed that part of the SH3 domain unfolds and refolds every min, which is far too slow to be detected by conventional HX NMR methods. This unfolding, however, is only partial. The difference in mass between the two envelopes (red and blue in Figure 4a) indicates that at least 17 residues participated in this cooperative unfolding process. The location of these unfolding residues was determined by a similar analysis of peptic fragments of labeled SH3. Two fragments displayed bimodal isotope patterns similar to those in the mass spectra of the intact protein. This bimodal isotope pattern is illustrated in Figure 4b for a peptic fragment that includes residues Spectra for the same fragment taken from SH3 but labeled for only 3 min before a significant population had unfolded as well as from SH3 labeled for 45 min after most of the SH3 had unfolded at least once are included in Figure 4b. Mass spectra of a peptic fragment with residues suggest that some of the residues in this segment also participate in the cooperative unfolding process. The locations of these peptic fragments are indicated by red in Figure 4c. Although not directly in the binding site, the region undergoing partial cooperative unfolding was positioned to potentially influence the binding site conformation. Results from an additional experiment with an SH3 domain bound to a 12-aminoacid peptide showed that binding slowed the unfolding rate but did not substantially change the structure of the unfolding region (24). Further studies in which the concentration of the peptide was varied indicated that unfolding persisted even when the domain was bound to the peptide ligand. The unfolding was observed in the isolated SH3 domain and in larger constructs containing the SH2 domain, suggesting that it has a role in the function of Hck (25). Unfolding of other SH3 domains has been reported, but these domains unfold on timescales faster than 1 s (30). Interactions between SH3 and SH2 domains. The properties of large, multidomain proteins are often investigated by studying the isolated structural domains of the parent protein. This approach opens the possibility that some functions may not be the same in the isolated domain because they are in larger, multidomain constructs. Extrapolating the NMR structures of isolated domains to the structures of the intact protein is of particular concern. Because HX MS can compare structures by their exchange properties, this method may be useful for extrapolating the structures of isolated domains to the intact protein structure. This approach has been used, for example, to compare the structures of isolated SH3 and SH2 domains with their structures in a larger construct (25). Structures of joint constructs SH(3+2), in which SH2 directly follows SH3 in sequence, as well as structures of intact human Hck protein in the inactive state, have been reported (31). Results of these studies showed that the structures of the SH2/SH3 domains in larger constructs are very similar to those of the isolated SH2/SH3 constructs, which is consistent with the SH2 and SH3 domains folding as individual units that contact each other only through the SH3/SH2 linker. Continuous labeling experiments were used to label isolated SH3 and SH2 domains, as well as the joint SH3/SH2 construct. Differences in deuterium levels indicated minor structural changes that had escaped detection by NMR or X-ray crystallography. Results for two peptic fragments derived from an isolated SH3 domain (closed data points) and from SH3 joined to SH2 (open data points) are presented in Figures 5a and 5b. At the shortest labeling time, the peptic fragment with residues (the N terminus of the SH3 construct), which was derived from the joint construct, had two more deuteriums than the same fragment from an isolated SH3 domain. This observation indicates that the structures of the two constructs differ in the region of residues The deuterium level in another segment was the same for both constructs (e.g., residues 87 99, Figure 5b), suggesting similar structures. M AY 1, / A N A LY T I C A L C H E M I S T R Y A

10 (a) Deuterium level (b) Time (min) Results for other peptic fragments are summarized in Figure 5c. The intensity of red indicates the increasing levels of deuterium in the joint construct, whereas the intensity of blue indicates the decreasing levels of deuterium. These results suggest that the structure of SH3 in the joint construct is significantly more mobile less hydrogen bonding and with better access to the solvent than in the isolated domain. In contrast, H/D exchange was slightly lower in the SH2 domain when part of the joint construct. These observations illustrate how HX MS can be used to determine whether the structures of isolated domains can be extrapolated to larger constructs. (c) SH3 bf Increased HX ba bb ab bc be SH(3+2) compared with isolated SH3 & SH2 bb be bd ba Decreased HX RT-loop aa SH2 FIGURE 5. Deuterium levels found in peptic fragments and in SH3 and SH2 domains. The differences between SH(3+2) and isolated SH2/SH3 is color coded as increasing (red) or decreasing (blue) deuterium levels in SH(3+2) relative to the levels found in the isolated SH2 and SH3 domains. The intensity of the color indicates the amount of change. (a) The peptic fragment for residues from the SH3 domain alone (filled points) and the SH(3+2) construct (filled points). (b) Residues from SH3 alone (filled points) and the SH(3+2) construct (filled points). (c) Isolated SH3/SH2 versus SH3/SH2 as part of the SH(3+2) construct. (Adapted with permission from Ref. 25.) bc bd Isotope exchange in proteins Initially, tritium was used for detecting protein structural changes by amide HX. Although H/D exchange has been detected by UV and IR methods, NMR has become the reference technique on which other methods are judged because it can detect exchange at specific protein sites. The growing use of MS to detect H/D exchange suggests that the technique offers some advantages, including the ability to detect peptides and proteins with very high sensitivity; study partially exchanged proteins when exchange has been quenched by acid; analyze large proteins, either intact or as proteolytic fragments; and determine the intermolecular distributions of deuterium. The high sensitivity of MS allows time course studies to be completed with subnanomolar quantities of proteins. More importantly, MS can analyze proteins at submicromolar concentrations. This feature aids studies of minimally soluble proteins, as well as partially unfolded proteins. Whether studies use NMR or MS, isotope exchange is usually quenched before the analysis. Quenching in NMR experiments is typically achieved by rapidly refolding the protein to its native state, where resonance signals have been previously assigned. Although refolding slows exchange at many amide linkages, exchange at many others remains too fast to be readily detected. In MS measurements, H/D exchange is quenched by decreasing the ph and temperature, which effectively slows exchange at all peptide linkages. As a result, structural changes along the entire polypeptide chain can be detected. Adding acid and lowering the temperature to quench exchange will likely be particularly useful for protein studies of large complexes in which the protein must be isolated from a complex matrix. For example, this approach has been used to simultaneously quench isotope exchange and disassemble deuterium-labeled viral capsids (32). Both ESI and laser desorption/ionization MS have been used to analyze intact proteins with molecular masses >100,000. On the other hand, although methods for analyzing large proteins by NMR are advancing, most proteins studied this way have molecular masses <30,000. Proteolytic digestion followed by analysis of the peptide fragments is the second route through which H/D exchange in large pro-

11 teins can be detected by MS. Although some proteins may resist digestion, the high activity of immobilized proteases suggests that it will be possible to digest most proteins under conditions where exchange is minimal. The simple view of protein folding envisions all the protein molecules folding to a common structure, but it is becoming increasingly evident that there may be considerable structural heterogeneity along these pathways. The intermolecular distribution of naturally occurring heavy isotopes is normally random, giving the binomial distribution of isotope peaks found in most mass spectra. However, the structural heterogeneity that alters H/D exchange rates may lead to bi- or multimodal isotope patterns from which the populations of the various structural forms can be determined. Challenges ahead HX MS will play a greater role in studies of protein structures and dynamics and will be particularly useful for bridging the gap between high-resolution three-dimensional protein structures determined by NMR and X-ray diffraction under nonideal conditions versus structures when proteins are doing interesting things. For example, being able to label proteins on the millisecond timescale will facilitate studies of relatively fast processes. In addition, most proteins can be studied by HX MS under a wide range of conditions, including different phs, high concentrations of denaturants, and as a component of large heterogeneous complexes. Nevertheless, HX MS needs to develop further in several areas, including automating sample preparation and data processing, scaling down chromatography to take advantage of ultrasensitive MS, and validating gas-phase fragmentation methods that determine deuterium levels at specific peptide linkages. For those who do not wish to develop such mundane aspects of the methodology, there is ample room to imagine challenging new applications of HX MS. This work was supported by a grant from the National Institutes of Health (GM RO1 40,384) and the Nebraska Center for Mass Spectrometry. John Engen works in the biological sciences section at Los Alamos National Laboratory. His research interests center on using MS to study higher order structures of proteins. David Smith is a professor at the University of Nebraska. His research interests focus on using MS to study the structure and dynamics of proteins. Address correspondence about this article to Smith at the Department of Chemistry, University of Nebraska, Lincoln, NE (dsmith7@unl.edu). References (1) Englander, S. W.; Mayne, L.; Bai, Y.; Sosnick, T. R. Protein Sci. 1997, 6, (2) Raschke, T. M.; Marqusee, S. Curr. Opin. Biotechnol. 1998, 9, (3) Englander, S. W.; Kallenbach, N. R. Q. Rev. Biophys. 1984, 16, (4) Miller, D. W.; Dill, K. A. Protein Sci. 1995, 4, (5) Li, R.; Woodward, C. Protein Sci. 1999, 8, (6) Bai, Y.; Sosnick, T. R.; Mayne, L.; Englander, S. W. Science 1995, 269, (7) Bai, Y.; Milne, J. S.; Mayne, L.; Englander, S. W. Proteins: Struct., Funct., Genet. 1993, 17, (8) Kim, K.-S.; Woodward, C. Biochemistry 1993, 32, (9) Miranker, A.; Robinson, C. V.; Radford, S. E.; Aplin, R. T.; Dobson, C. M. Science 1993, 262, (10) Zhang, Z.; Post, C. B.; Smith, D. L. Biochemistry 1996, 35, (11) Yang, H.; Smith, D. L. Biochemistry 1997, 36, (12) Chamberlain, A. K.; Handel, T. M.; Marqusee, S. Nature Struct. Biol. 1996, 3, (13) Deng, Y.; Smith, D. L. Biochemistry 1998, 37, (14) Deng, Y.; Smith, D. L. J. Mol. Biol. 1999, 294, (15) Deng, Y.; Smith, D. L. Anal. Biochem. 1999, 276, (16) Chen, J.-W.; Smith, D. L. Biochemistry 2000, 39, (17) Thévenon-Emeric, G.; Kozlowski, J.; Zhang, Z.; Smith, D. L. Anal. Chem. 1992, 64, (18) Zhang, Z.; Li, W.; Logan, T. M.; Li, M.; Marshall, A. G. Protein Sci. 1997, 6, (19) Zhang, Z.; Smith, D. L. Protein Sci. 1993, 2, (20) Gamblin, S. J.; Cooper, B.; Millar, J. R.; Davies, G. J.; Littlechild, J. A.; Watson, H. C. FEBS Lett. 1990, 262, (21) Deng, Y.; Pan, H.; Smith, D. L. J. Am. Chem. Soc. 1999, 121, (22) Eyles, S. J.; Speir, J. P.; Kruppa, G. H.; Gierasch, L. M.; Kaltashov, I. A. J. Am. Chem. Soc. 2000, 122, (23) Deng, Y.; Zhang, Z.; Smith, D. L. J. Am. Soc. Mass Spectrom. 1999, 10, (24) Engen, J. R.; Smithgall, T. E.; Gmeiner, W. H.; Smith, D. L. Biochemistry 1997, 36, 14,384 14,391. (25) Engen, J. R.; Smithgall, T. E.; Gmeiner, W. H.; Smith, D. L. J. Mol. Biol. 1999, 287, (26) Engen, J. R.; Gmeiner, W. H.; Smithgall, T. E.; Smith, D. L. Biochemistry 1999, 38, (27) Cohen, G. B.; Ren, R. B.; Baltimore, D. Cell 1995, 80, (28) Kuriyan, J.; Cowburn, D. Curr. Opin. Struct. Biol. 1993, 3, (29) Pawson, T.; Schlessinger, J. Curr. Biol. 1993, 3, (30) Zhang, O.; Forman-Kay, J. D. Biochemistry 1995, 34, (31) Sicheri, F.; Moarefi, I.; Kuriyan, J. Nature 1997, 385, (32) Wang, L.; Smith, D. L. Protein Sci. 2001, in press (33) Schindler, T.; Sicheri, F.; Pico, A.; Gazit, A.; Levitzki, A.; Kuriyan, J. Mol. Cell. 1999, 3, M AY 1, / A N A LY T I C A L C H E M I S T R Y A