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1 Department of Chemistry, Loyola University, New Orleans, Louisiana 70118, USA
2 Institute for Cancer Research, Fox Chase Cancer Center, Philadelphia, Pennsylvania 19111, USA
3 Department of Biochemistry, University of Wisconsin, Madison, Wisconsin 53706, USA
4 Department of Biochemistry and Biophysics, University of Pennsylvania, Philadelphia, Pennsylvania 19104-6059, USA
Reprint requests to: W.F. Walkenhorst, Chemistry Department, Loyola University New Orleans, Box 5, 6363 St. Charles Avenue, New Orleans, LA 70118, USA; e-mail: walken{at}loyno.edu; fax: (504) 865-3270.
(RECEIVED July 18, 2001; ACCEPTED October 11, 2001)
Article and publication are at http://www.proteinscience.org/cgi/doi/10.1101/ps.28202.
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Abstract |
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-helical domain of H124L SNase. To further characterize the folding intermediate, protection factors for individual amide sites were measured by varying the pH of the labeling pulse at a fixed refolding time of 16 msec. Protection factors >5.0 were observed only for amide positions in a ß-hairpin formed by strands 2 and 3 of the ß-barrel domain and a single site near the C-terminus. The results indicate that formation of stable hydrogen-bonded structure in a core region of the ß-sheet is among the earliest structural events in the folding of SNase and may serve as a nucleation site for further structure formation. Keywords: Staphylococcal nuclease; pulse labeling; hydrogen exchange; NMR; protein folding
Abbreviations: Cm, midpoint of a GuHCl denaturation transition • DCl, 2HCl; D2O, 2H2O • GuHCl, guanidine hydrochloride • H124L SNase, staphylococcal nuclease with a histidine-to-leucine substitution at position 124 (sequence equivalent to the staphylococcal nuclease isolated from the V8 strain of Streptococcus aureus) • m, slope of the GuHCl denaturation transition in units of kcal mol-1 M-1 • HSMQC, heteronuclear single-quantum, multiple-quantum correlation • pdTp, deoxythymidine-3`,5`-bisphosphate • pH*, pH of a sample dissolved in D2O as determined by an uncorrected glass electrode measurement • Pro-, mutant of SNase with P11A, P31A, P42A, P47G, P56A, and P117G substitutions • SNase, wild-type staphylococcal nuclease with a sequence equivalent to that isolated from the Foggi strain of S. aureus).
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) has been found to strongly increase the stability of SNase (Hinck et al. 1996). Although the structure of SNase can be divided into two subdomains (Fig. 1), an N-terminal ß-barrel domain comprised of five antiparallel ß-strands and a C-terminal cluster of three -helices, the wild-type protein shows the cooperative unfolding transition characteristic of single-domain proteins (Anfinsen et al. 1972; Shortle and Meeker 1986). Structural and mutational data on SNase and several deletion mutants suggested that residual structure existed, either in the denatured state or in a subdomain of the protein (Shortle et al. 1990; James et al. 1992; Shortle 1992, 1993; Shortle and Abeygunawardana 1993; Alexandrescu et al. 1995).
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For further investigation of this system, we chose to work with H124L SNase, the more stable variant originally isolated from the V8 strain of Streptococcus aureus, rather than the wild-type enzyme (SNase), which was isolated from the Foggi strain of S. aureus. H124L SNase differs from SNase by a single residue substitution in helix 3 (Fig. 1
), which stabilizes the protein by 1.3 kcal mol-1 (GuHCl denaturation of H124L SNase: 
G = 6.91 kcal mol-1 at 20°C at pH 7; GuHCl denaturation of SNase: G = 5.62 kcal mol-1 [Shortle 1986]). We describe below the use of pulsed hydrogen exchange methods to measure the time course and magnitude of protection of amide protons as probes for the development of structure during early stages of folding.
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1.5 M (Cm = 1.08 M) at pH 7 (Shortle and Meeker 1986). An unfolded control sample (pH 3.0) exposed to the labeling pulse at high pH was fully labeled when compared to a folded control, confirming that H124L SNase has no residual backbone NH protection at acid pH and low ionic strength. The stability of H124L SNase to the conditions of the labeling pulse were verified by equilibrium CD measurements (see Fig. 2) and kinetically by using stopped-flow fluorescence (see Materials and Methods) to monitor the protein following pH jumps from pH 5 to pH 9 and from pH 5 to pH 10.
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shows a plot of the normalized initial amplitudes (A0) for backbone amide protons in H124L SNase after 10 msec of refolding. The most striking observation is that more than half of the 60 observed amide sites show initial amplitudes significantly lower (A0 <0.8) than the value expected for a fully labeled amide site in the unfolded state (A0 = 1). Thus, a species accumulates during the dead time of the quenched flow experiment in which many of the amide sites found to be highly protected in the folded protein are already partially protected against exchange with solvent hydrogens. These rapidly protected sites include positions in all strands of the beta sheet (Fig. 3), in each of the three major -helices (H1–H3, Fig. 1), and in a minor hydrophobic domain, which includes residues 37–40 and 110–112 (Wang et al. 1990b; Wang and Shortle 1996; Loh et al. 1993). The lowest A0 values are observed at sites in strands 2 and 3, whereas many positions in the three -helices show no protection at all (see Figs. 3, 4B). Qualitatively similar results were observed by Jacobs and Fox (1994) for the P117G mutant of SNase for a smaller set of 39 amide protons resolved by homonuclear 1H NMR spectroscopy.
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as a function of time. In panel A, the time course of protection for Val99, located in helix 2, is shown; the data are from all 15 samples measured and cover 10 time points between 10 msec and 160 sec. The time course of protection is also shown for additional amide sites located in the helical (Fig. 4B) and ß-sheet (Fig. 4C) regions of H124L SNase. The protection observed between 10 msec and 160 sec (in addition to the early events that give rise to A0) occurs in two kinetic phases of nearly equal amplitude, which are similar for all probes (Fig. 4A–C). The kinetic data for 53 of the 60 observed amide protons were fitted to two exponentials. The rates from the fits are plotted as a function of residue number in Figure 5. The rates of the faster process fall in the range between 2 and 11 sec-1, whereas those of the slower phase range from 0.08 to 0.25 sec-1. The fits indicate a global protection pattern in which all 53 probes show, within error, a nearly identical rate of 5 ± 1 sec-1 for the faster phase and a rate of 0.13 ± 0.01 sec-1 for the slower process (99% confidence interval from data in Fig. 5).
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), can arise from structure formation during refolding, from intrinsically slow exchange for certain amino acid sequences, or through exchange of these sites with solvent deuterons at a later time before NMR data collection. To distinguish among these possibilities, protection experiments were performed at a fixed refolding time as a function of the pH of the labeling pulse. A tf of 16 msec was chosen as optimal to simultaneously measure a protection factor as early as possible in refolding while allowing for a long-enough labeling pulse (50 msec) to completely exchange most unprotected amide sites at all pH values chosen for the analysis. The proton occupancies compared to fully protonated control spectra were measured after 16 msec of refolding for eight pH values between pH 7.3 and pH 10.7. The data were then fitted as described previously (see Materials and Methods) to extract protection factors for 75 different amide sites in H124L SNase. The fits for sets of amides in ß-strand 2 (V23, K24, L25, and M26) and helix 2 (E101 and R105) and the extended hydrophobic region involving helix 2 (L108 and A109) are compared in Figures 6A and 6B, respectively. The amide protons located in ß-strand 2 showed measurable protection factors with an average value of P = 6.5 (Fig. 6A), indicating that they take on average about six times as long to exchange with solvent as would be predicted for an unstructured region with the same protein sequence at this temperature. The amide protons in and near helix 2, on the other hand, show no protection and exchange at the predicted rate for this sequence (Bai et al. 1993). The protection factors for all amide protons measured in this experiment are plotted in Figure 7 along with an indication of the secondary structure in the native protein sequence. Amides with protection factors Pf that fall in the range between 2 and 5 are observed primarily in ß-strands 2 and 3, but a few residues in strands 4 and 5 and in each of helices H1–H3 also fall in this range as do several amides near the C-terminus. Amide protons with Pf >5.0 are observed primarily in ß-strands 2 and 3. Other residues with slightly elevated protection factors (Pf = 3–7) occur in a minor hydrophobic domain near Trp 140 (Loh et al. 1993), which includes residues 134 in helix 3, and residues 138–141. In Figure 8, amide protons with Pf >5.0 are indicated by black spheres, whereas those with Pf between 2.5 and 5.0 are indicated by gray spheres.
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Discussion |
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To detect protection of amide protons during refolding, NH groups must be shielded from solvent, either by being buried away from solvent or as is more often the case, by becoming involved in an intramolecular hydrogen bond during refolding. A second prerequisite is that the amides protected in refolding must also be protected in the native structure. Despite this requirement, the patterns of protected amide protons can provide information even on nonnative structure in folding intermediates. This was shown recently for ß-lactoglobulin, in which the formation of nonnative helical structure was detected preceding the formation of native ß-sheet (Kuwata et al. 2001). More generally, observation of nonnative structure requires that amide positions be protected in both the intermediate and the native structures and that the patterns or rates of protection contain clues that allow the existence of nonnative structure to be inferred. For example, if a ß-hairpin is formed in an intermediate, alternating amide sites in that region would be protected. If this region converted to an -helix (or a pair of -helices) in the fully folded protein, the resulting pattern of protection would differ from that of a sequence that contained an -helix in both the intermediate and the native protein.
To establish that protection of amide positions is attributable to the formation of structure during folding, it is necessary to verify that there is no residual structure under denaturing conditions. Fluorescence measurements following denaturation by acid (Walkenhorst et al. 1997) or by GuHCl (Shortle 1986) suggested that nuclease was completely unfolded by acid or by GuHCl alone. Previous hydrogen exchange studies by Loh et al. (1993) also suggested that there was no residual structure in denatured SNase. On the other hand, a number of observations by NMR and other methods indicates that SNase and some of its deletion mutants contain residual structure under some denaturing conditions (Wang and Shortle 1995; Gillespie and Shortle 1997; Filfil and Chalikian 2000). To resolve this question for these experiments, we looked for residual NH protection by performing real-time exchange measurements under several sets of denaturing conditions. The observed rates were within a factor two of the predicted rates (Bai et al. 1993) and were identical within error for solutions at pH 3 with or without added GuHCl (2.5 M). This is consistent with previous hydrogen exchange studies on SNase deletion mutants, which found that protection factors for a denatured state were no larger than 2.4 (Mori et al. 1997), whereas those in a structured deletion mutant showed protection factors between 1 and 190 (Alexandrescu et al. 1996). This justified our use of pH 3 alone under low ionic strength conditions to denature SNase.
Our results for protection in the dead time of the pulse-labeling experiment (10 msec) are similar to those of Jacobs and Fox (1994) on the P117G variant of SNase. In particular, both studies show evidence for population of a folding intermediate consisting largely of the ß-sheet domain of SNase with some missing amplitude for amide sites outside the ß-barrel. In our studies, we were able to observe between 60 and 75 amide protons (depending on field strength), which included many more residues in the helices and turn regions than the 39 amides reported by Jacobs and Fox (1994), despite the addition of stabilizing salts in their buffer solutions. We attribute this to two main factors: (1) The use of heteronuclear 1H-15N HSMQC experiments, rather than the proton NMR used by Jacobs and Fox, allowed us to identify a larger number of signals; (2) the addition of the stabilizing ligands Ca2+ and pdTp directly in the quench buffer to prevent the loss of information because of exchange during sample workup.
To measure protection factors in the intermediate, we chose to vary the pH of the labeling pulse in a series of experiments with a fixed refolding time of 16 msec. Using a refolding delay of 100 msec followed by a 50-msec labeling pulse, Jacobs and Fox (1994) observed measurable protection factors throughout the protein with the largest protection observed for residues in the ß-barrel. The main result from our studies is that at an earlier stage of folding (16 msec), we observe protection factors >5.0 only in a ß-hairpin consisting of strands 2 and 3 of the ß-barrel, suggesting that this region of the protein is the first to acquire stable hydrogen bonds and may act as a nucleation site for structure formation. This is consistent with previous evidence that this ß-hairpin is partially structured in a deletion mutant of SNase (Wang and Shortle 1995; Gillespie and Shortle 1997). Recent studies of the pentapeptide N-acetyl-YKGQP-NH2, which contains the central residues of this ß-hairpin, indicated that turn conformations were partially populated in solution (Ramakrishna and Sasidhar 1997). Studies on a longer 15-residue peptide containing the entire ß-hairpin were inconclusive because of its strong tendency to aggregate (Ramakrishna and Sasidhar 1998).
Interestingly, residues in strands 2 and 3 contain some of the most slowly exchanging amide protons under native conditions (Loh et al. 1993). In particular, the exchange rates for residues 23–26 are consistent with exchange via global unfolding. Thus, the most stable core region of the native SNase structure is also among the earliest to acquire persistent hydrogen bonds during folding, as is observed for a number of other proteins (e.g., Roder et al. 1988; Woodward 1993; Bai et al. 1994; Raschke and Marqusee 1997; Kuwata et al. 2001).
The global rates of 5 sec-1 and 0.13 sec-1 observed in our time-resolved labeling experiments are consistent with similar rates detected by other methods (Sugawara et al. 1991; Chen and Tsong 1994; Kalnin and Kuwajima 1995; Maki et al. 1999; Walkenhorst et al. 1997). There appears to be no detectable protection of amides associated with a 1-sec-1 phase observed by stopped flow, which is consistent with our assignment of this phase to a nonproline cis-trans isomerism in our previous stopped-flow fluorescence studies (Walkenhorst et al. 1997). We also see no additional changes in proton occupancy at folding times >20 sec after Pro117 has isomerized to cis, suggesting that this process causes no additional protection detectable in our experiment.
Although we have not directly followed the time course of formation of the early intermediate containing the ß-hairpin by pulsed hydrogen exchange, this process may be related to the observation of a lag phase over the 2–10 msec time range in our earlier stopped-flow fluorescence experiments on H124L nuclease and a proline-free variant (Walkenhorst et al. 1997). The amide protection patterns observed at a folding time of 16 msec (Fig. 7) differ substantially from those measured previously at a folding time of 100 msec followed by a 50-msec labeling pulse (Jacobs and Fox 1994). At these longer times, well-protected amide sites (protection factors in the range 10–60) were found not only in ß-strands 2 and 3, but also strands 4 and 5, as well as isolated residues in helices 2 and 3. Because the time-resolved pulse-labeling experiments show only minor changes over the 10–100 msec time range (Fig. 4), the increase in the level and number of protected sites is probably attributable to the partial accumulation of a more stable native-like species in the subsequent kinetic phase, which has a time constant of 200 msec.
Initiation sites for protein folding
An important goal of protein-folding studies is to identify regions of the chain that become structured during early stages of refolding and, therefore, are potential nucleation sites for folding. For many small, single-domain proteins, the first appearance of a nucleus of native-like contacts is also the rate-limiting step in folding, resulting in a concerted (two-state) folding process. However, for many other proteins, including SNase, this is not the case. The initial stabilization of a core region of the structure (in the present case a central pair of ß-strands that forms on the 10-msec time scale) is succeeded by a series of slower conformational events, giving rise to transient accumulation of partially structured states. For proteins with two-state folding mechanisms, Plaxco et al. (1998) found a striking (inverse) correlation between the rate of folding and the contact order, a measure of the average sequence separation of contacting residues. Thus, the kinetic barrier encountered in small proteins comprising a single folding unit appears to be dominated by the overall topology of the chain rather than specific aspects of their amino acid sequence. However, structurally more complex proteins that do not follow a simple two-state folding mechanism often fold more slowly than predicted based on their contact order (Plaxco et al. 2000). This is also the case for SNase. The folding rate predicted based on its relatively low contact order (10%) is several orders of magnitude faster than the experimentally observed rate for the major folding step (5 sec-1). Thus, the rate-limiting barrier in the formation of the native SNase structure is clearly dominated by factors other than the overall topology of the chain, such as specific tertiary interactions or docking of subdomains. On the other hand, topological features such as the ß-hairpin in SNase described here are more likely to be an important factor in determining the rate at which the early intermediate is formed.
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Pulsed hydrogen-exchange experiments
A Biologic QFM-5 quenched flow instrument was used to label unprotected backbone amide positions at various times during refolding. A 3-mM solution of H124L SNase in 99.9 % D2O (Isotec) was unfolded by lowering the pH* (uncorrected glass electrode reading) to below 3.0 by the addition of DCl. All exchangeable NH groups were deuterated by first placing the unfolded protein solution at 65°C for 1 h, followed by incubation overnight at 25°C. Refolding was initiated at 15°C by a 1:2 dilution of the unfolded protein with a refolding buffer containing 150-mM KCl and 75-mM sodium acetate (pH 6.8) such that the final refolding mixture had a pH* of 5.3 and contained 100-mM KCl and 50-mM sodium acetate. All subsequent buffers used in the pulse-labeling experiment contained 100-mM KCl. All pH values were verified by manual mixing at 15°C before running the experiments. Refolding was allowed to continue for a variable length of time for 10 samples covering folding times (tf) of 10, 33, 60, 115, 250, and 750 msec, and 2.5, 20, 60, and 160 seconds. A number of these time points (115 msec, 750 msec, 2.5 sec, 20 sec, and 160 sec) were repeated in independent experiments as controls for procedural variations such as the choice of continuous or interrupted mixing mode (necessary for time points >200 msec) or variation of solvent composition. For example, the refolding buffer contained H2O for shorter refolding times, but D2O was used for samples with a tf longer than a few seconds to prevent exchange with solvent protons from occurring during folding. As a control, the 2.5-sec time point was run independently under each condition: one trial with refolding buffer containing H2O and one trial with refolding buffer containing D2O.
After refolding for the designated amount of time, an H2O labeling pulse (tp) was introduced at high pH to exchange deuterons for protons at unprotected amide sites. For samples refolded in H2O, this was accomplished by a 1:1 dilution of the refolding mixture with a solution containing 100-mM glycylglycine adjusted to provide a pH of 8.8 after mixing. For samples refolded in D2O, a 1:4 dilution was used to achieve a sufficiently high molar fraction of H2O during the labeling pulse. Because the initial unfolded protein solution always contained 99.9% D2O, the final conditions during the labeling pulse were 83% H2O for the samples refolded in H2O and 80% H2O for those refolded in D2O. The labeling was allowed to continue for 57 msec before quenching and completion of folding at lowered pH was initiated in a third and final mixing event. The quench buffer contained 300-mM sodium acetate in D2O, adjusted to provide a final pH* of 5.3 after mixing, as well as 20-mM CaCl2 and 1 mM of the competitive inhibitor pdTp (Cal Biochem). The latter components were included because the binding of Ca++ and pdTp are known to stabilize SNase and to drastically decrease the rates of hydrogen exchange for many backbone protons (Loh et al. 1993). The samples for each time point were kept on ice before workup. Final concentration of the samples and transfer into a suitable NMR buffer (50-mM d4-acetic acid, 0.1-M KCl, 20-mM CaCl2, 1-mM pdTp in 99.9% D2O) was accomplished by three sequential cycles of dilution and concentration at 4°C using Centri-Prep concentrators (Amicon) with a MW cutoff of 10 kD.
Measurement of protection factors
Protection factors after 16 msec of refolding were measured on a Biologic stopped flow/quenched flow instrument (SFM-4/QFM-4). The extent of protection for backbone amide sites was probed by systematic variation of the labeling pulse pH in separate experiments at a constant refolding time. The pH of the 50-msec labeling pulse was varied for eight samples in the range between 7.0 and 11.0 following a fixed refolding time of 16 msec. The pH values after mixing were 7.3, 8.0, 8.3, 8.5, 8.9, 9.5, 9.9, and 10.7 at 15°C. The labeling buffers all contained 100-mM KCl in H2O and were 100 mM in either HEPES (pH 7.3, 8.0), glycylglycine (pH 8.3, 8.5), CHES (pH 8.9, 9.5, 9.9), or CAPS (pH 10.7). All other details are as described above for the time-course data.
Folded and unfolded controls
A separate, protonated control sample was run along with each set of folding data to control for differences in NMR instrumentation and sensitivity. The protonated controls were prepared by dissolving 30 mg of H124L SNase in 2.0 mL of 100-mM glycylglycine buffer (pH 9.0) containing H2O and D2O in a 80:20 ratio. The sample was equilibrated by raising the temperature to 65°C for 10 min. The sample was then cooled and the pH lowered to 5.0, followed by dilution with four volumes of cold quench buffer. The control samples were concentrated and worked up in parallel with the experimental samples to ensure that all samples had the same history.
A number of other controls were run to test the unfolding and labeling conditions used in the pulse-labeling studies. To test whether the protein was unfolded by acid pH alone, real-time NH exchange was monitored at 15°C by proton-detected 1H-15N HSMQC experiments (described below) for protein samples prepared at pH 3.0 in the presence and absence of 2.5-M GuHCl. Exchange was initiated at 15°C using a spinning Sephadex column to transfer the protein into a 99.9% D2O solution buffered to pH* 3.0 (±GuHCl) with 10-mM citrate, and spectra were collected every 30 min for a period of 4 h to monitor exchange.
To ensure that the labeling pulse was neither too strong nor too weak, both hydrogen exchange and stopped-flow fluorescence controls were run. To assess the stability of the protein to the high pH conditions of the labeling pulse, we measured the unfolding kinetics for H124L SNase by monitoring changes in Trp fluorescence following pH jumps from pH 5 to 9 and pH 5 to 10 using a six-fold dilution into 100-mM glycine buffer. An alkaline titration of H124L SNase by CD was performed over the pH range from 7 to 12 to verify that nuclease remained stable to the pH values used in the labeling pulse. The stability of the folded protein to the labeling pulse was also assessed by the use of a very long tf of 160 sec preceding the high pH pulse. To verify that the calculated labeling strength (calculated D-H exchange rate at a given pH times the length of the labeling pulse) was sufficient to completely label all unprotected amide positions, an unfolded protein sample was subjected to the labeling pulse and compared to a fully protonated (folded) control sample prepared as described above.
NMR experiments
NMR experiments were run on an AM-500 for the time course and real-time exchange data and on a DMX-600 for the protection factor measurements. The time-course and protection data were collected at 295K, whereas the real-time exchange data were collected at 288K. 1H-15N HSMQC experiments (Zuiderweg 1990) with proton detection were run with 256 experiments (t1) consisting of 32 scans and 2048 data points (t2), except for the real-time exchange experiments, which were run with eight scans and 128 experiments. To aid in assignment, HSMQC spectra were also collected between 295K and 315K in increments of 5K. All spectra were processed identically, with apodization by squared sine-bell functions in each dimension before zero-filling to a digital resolution of 2 points/Hz. One-dimensional 1H reference spectra were collected for each folding time for use in normalization between samples.
Data analysis
NMR peak assignments previously made at 310 K and 318K (Wang et al. 1990a; Loh et al. 1993) were extended to 295K via analysis of the temperature titration data described above. Peak heights and volumes were measured using macros written in the Felix software package (Molecular Simulations). The uncertainty in the peak height for a given NH proton (Pi) was calculated from Pi/Pi = Ii /Ii (tf) + Ii /Ii (control), in which P is the peak height and I is the intensity of that peak in the control spectra and in a particular folding time point. Differences in sample concentration were taken into account by normalizing each 2D spectra according to the averaged heights of five nonexchangeable proton resonances in the 1D reference spectra (described above) collected separately for each sample. Time-course data were fit to two exponentials using a nonlinear least-squares algorithm. Protection factors (Pf) were determined by fitting the pH-dependent proton occupancies, f(pH), to the expression f = 1 - (exp(-kc*tp/Pf)) (tp = 50 msec), in which kOH = 8.063*107 and kc = kOH*10 (pH-14.35), based on (Elöve and Roder 1991). Intrinsic hydrogen exchange rates as a function of pH, temperature, and amino acid sequence were estimated on the basis of the model-peptide data of (Bai et al. 1993) using the program HX-Pred (www.fccc.edu/research/labs/roder).
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1 (open circles) fall between 2 and 11 sec-1 with an average value of 5 ± 1 sec-1, whereas those for 
