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Protein Science (2003), 12:2768-2781.
Copyright © 2003 The Protein Society

Water and urea interactions with the native and unfolded forms of a ß-barrel protein

Kristofer Modig1, Elizabeth Kurian2,3, Franklyn G. Prendergast2 and Bertil Halle1

1 Department of Biophysical Chemistry, Lund University, SE-22100 Lund, Sweden
2 Department of Biochemistry and Molecular Biology, Mayo Foundation, Rochester, Minnesota 55905, USA

Reprint requests to: Bertil Halle, Department of Biophysical Chemistry, Lund University, Box 124, SE-22100 Lund, Sweden; e-mail: bertil.halle{at}bpc.lu.se; fax: 46-46-222-4543.

(RECEIVED June 15, 2003; FINAL REVISION August 20, 2003; ACCEPTED August 20, 2003)

3 Deceased. Back

Article and publication are at http://www.proteinscience.org/cgi/doi/10.1110/ps.03262603.


   Abstract
TOP
 Abstract
Introduction
 Results and Discussion
 Materials and methods
 References
 
A fundamental understanding of protein stability and the mechanism of denaturant action must ultimately rest on detailed knowledge about the structure, solvation, and energetics of the denatured state. Here, we use 17O and 2H magnetic relaxation dispersion (MRD) to study urea-induced denaturation of intestinal fatty acid-binding protein (I-FABP). MRD is among the few methods that can provide molecular-level information about protein solvation in native as well as denatured states, and it is used here to simultaneously monitor the interactions of urea and water with the unfolding protein. Whereas CD shows an apparently two-state transition, MRD reveals a more complex process involving at least two intermediates. At least one water molecule binds persistently (with residence time >10 nsec) to the protein even in 7.5 M urea, where the large internal binding cavity is disrupted and CD indicates a fully denatured protein. This may be the water molecule buried near the small hydrophobic folding core at the D–E turn in the native protein. The MRD data also provide insights about transient (residence time <1 nsec) interactions of urea and water with the native and denatured protein. In the denatured state, both water and urea rotation is much more retarded than for a fully solvated polypeptide. The MRD results support a picture of the denatured state where solvent penetrates relatively compact clusters of polypeptide segments.

Keywords: Protein denaturation; fatty acid-binding protein; urea; solvent exchange; magnetic relaxation dispersion

Abbreviations: CD, circular dichroism • I-FABP, intestinal fatty acid-binding protein • MRD, magnetic relaxation dispersion • NOE, nuclear Overhauser effect


   Introduction
TOP
 Abstract
 Introduction
Results and Discussion
 Materials and methods
 References
 
Despite the widespread use of urea in studies of protein stability and folding thermodynamics (Kauzmann 1959; Tanford 1970; Myers et al. 1995), the molecular mechanism whereby urea unfolds proteins has not been established. Solvent denaturation is a result of altered protein–solvent interactions, but it is not clear whether denaturants like urea act directly by binding to the protein surface or indirectly by perturbing solvent-mediated hydrophobic interactions or by a combination of these mechanisms. The direct mechanism is made plausible by the structural similarity between urea and the peptide group, suggesting that urea–peptide interactions, like peptide–peptide interactions, can compete favorably with water–peptide interactions. If this is the case, then solvent denaturation can be driven simply by the exposure of more binding sites in the denatured protein (Schellman 1987). The indirect mechanism is supported by the observation that urea enhances the solubility of not-too-small nonpolar solutes or groups (Wetlaufer et al. 1964; Shimizu and Chan 2002) and, by implication, weakens the hydrophobic stabilization of the folded protein.

A fundamental understanding of protein stability, including the mode of denaturant action, must be based on experimental characterization of the structure, solvation, and energetics of the denatured state at the level of detail that has been achieved for the native state (Dill and Shortle 1991; Shortle 1996a). Denatured proteins have traditionally been modeled as fully solvated random coils, but a growing body of experimental evidence is challenging this view (Shortle 1996a; Denisov et al. 1999; Shortle and Ackerman 2001; Choy et al. 2002; Klein-Seetharaman et al. 2002). To quantitatively account for the often marginal stability of native proteins under physiological conditions, we need to examine denatured states from different vantage points using a variety of techniques. Although important progress has been made using NMR (Shortle 1996b) and small-angle scattering (Millet et al. 2002), few methods are available for directly probing the solvation of denatured proteins. One of these, water 17O magnetic relaxation dispersion (MRD), has previously been used to monitor both internal and surface hydration during thermal denaturation (Denisov and Halle 1998) and solvent denaturation by guanidinium chloride (Denisov et al. 1999). Here, we use 17O and 2H MRD to examine hydration as well as denaturant interactions during the urea-induced unfolding of the apo form of intestinal fatty acid-binding protein (I-FABP). 2H MRD has previously been used to study DMSO–protein interactions (Jóhannesson et al. 1997), but this is the first MRD study to monitor solvent and cosolvent/denaturant simultaneously.

Like the other members of the family of lipid-binding proteins (Banaszak et al. 1994; Zimmerman and Veerkamp 2002), the 15-kD cytoplasmic protein I-FABP has a ß-clam structure composed of 10 antiparallel strands that enclose a very large (500–1000 3) internal binding cavity (see Fig. 1Go). Lipids are thought to enter the cavity via a small "portal" lined by two short {alpha}-helices. The folding thermodynamics and kinetics of I-FABP have been studied extensively (Ropson et al. 1990; Ropson and Frieden 1992; Clark et al. 1997, 1998; Kim et al. 1997; Ropson and Dalessio 1997; Burns et al. 1998; Dalessio and Ropson 1998, 2000; Kim and Frieden 1998; Hodsdon and Frieden 2001; Yeh et al. 2001; Chattopadhyay et al. 2002a,b; Nikiforovich and Frieden 2002). The equilibrium denaturation of I-FABP by urea appears to be two-state and cooperative when monitored by optical spectroscopy, but NMR studies have indicated intermediate states (Ropson and Frieden 1992; Hodsdon and Frieden 2001).



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  		Figure 1. Crystal structure of apo I-FABP from PDB entry 1IFC [PDB] (Scapin et al. 1992) with the 10 ß-strands labeled. Water oxygens in the large binding cavity are colored blue, and the singly buried water oxygen in the D–E turn (labeled W135 in 1IFC [PDB] ) is red. Some of the hydrophobic residues thought to be involved in the hydrophobic folding core are colored green: from bottom to top, Ile40 (barely visible behind strand B), Phe47, Phe62, Trp82, Leu64, Phe68, and Val66. The figure was generated with the program MOLMOL (Koradi et al. 1996).

 
Although the folding of ß-sheet proteins does not appear to differ fundamentally from that of proteins with a classical hydrophobic core (Capaldi and Radford 1998), it is of interest to examine the role of the solvent-filled cavity in solvent denaturation. Another reason for choosing I-FABP for this denaturation study is that its internal and external hydration has been characterized in detail by 17O and 2H MRD (Wiesner et al. 1999; Modig et al. 2003). Briefly, these MRD studies have shown that the 20–25 water molecules that occupy the binding cavity (Scapin et al. 1992) remain trapped for more than 10 nsec before exchanging with external solvent. While residing in the cavity, these water molecules exchange among internal hydration sites on a time scale of 1 nsec. This intracavity exchange has also been characterized by molecular simulations (Likic and Prendergast 2001; Bakowies and van Gunsteren 2002). In addition, a singly buried water molecule (labeled W135 in the I-FABP crystal structure) is buried near a hydrophobic cluster at the turn between ß-strands D and E (see Fig. 1Go). This water molecule is conserved across the family of lipid-binding proteins, and must therefore contribute importantly to the stability of the native protein structure (Likic et al. 2000). W135 has a long residence time (Likic et al. 2000; Likic and Prendergast 2001; Bakowies and van Gunsteren 2002), and is thought to be the main (possibly, the only) contributor to the low-field 17O dispersion (Wiesner et al. 1999; Modig et al. 2003). Because the hydrophobic cluster at the D–E turn forms early on the folding pathway (Ropson and Frieden 1992; Kim et al. 1997; Hodsdon and Frieden 2001; Yeh et al. 2001; Chattopadhyay et al. 2002b; Nikiforovich and Frieden 2002), we can use W135 as an MRD marker for this (un)folding event.

An important aspect of the present work is the separation of water and urea contributions to the observed 2H relaxation. This allows us to directly probe urea interactions with I-FABP across the unfolding transition, while also monitoring the competing water interactions. The available structural data on urea–protein interactions are limited and, with few exceptions (Dötsch et al. 1995; Dötsch 1996), are restricted to native proteins (Lumb and Dobson 1992; Liepinsh and Otting 1994; Pike and Acharya 1994). Computer simulations of proteins in molecular solvent are still a long way from being able to access the time scales on which solvent-induced protein unfolding takes place. Therefore, simulations have so far only provided information about urea–protein interactions in the native state or in partially unfolded states at very high temperatures (Tirado-Rives et al. 1997; Caflisch and Karplus 1999).


   Results and Discussion
TOP
 Abstract
 Introduction
 Results and Discussion
Materials and methods
 References
 
Water 17O relaxation in bulk aqueous urea solutions
For reference purposes, we measured the water 17O relaxation rate in protein-free samples with the same solvent composition as in the I-FABP solutions. The 17O relaxation rate Rbulk in these bulk urea solutions increases with the urea concentration C U (in mol dm-3) as


(1)

In Figure 2Go, this weak concentration dependence is contrasted with the five to six times stronger urea-induced viscosity enhancement (Kawahara and Tanford 1966). The insensitivity of the water 17O (Bagno et al. 1993) and 2H (Yoshida et al. 1998) relaxation rates to the presence of urea has been noted previously. In fact, at the low urea concentrations (C U < 2 M) investigated previously, the effect of urea was barely significant.



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  		Figure 2. Relative variation of the water 17O relaxation rate Rbulk and shear viscosity {eta} with urea concentration in aqueous solutions at 27°C. The Rbulk data (filled circles) were fitted to the cubic polynomial in equation 1Go (solid curve). The viscosity (dashed curve) is described by the empirical relation (Kawahara and Tanford 1966).

 
Assuming that only the nS water molecules in the first hydration shell of urea differ from bulk water, we can express the coefficient of the linear term in equation 1Go as , where {tau}S and {tau}bulk are the rotational correlation times of water molecules in the hydration shell and in bulk water, respectively, and is the molar volume of water. Taking nS = 12.6, as obtained by integrating the first peak in simulated pair correlation functions (Kuharski and Rossky 1984), we find that the rotation of water molecules in contact with urea is retarded by merely 3% ({tau}S/{tau}bulk = 1.025). Concerning rotational dynamics, water in urea solutions can therefore be regarded as essentially unperturbed bulk water. This conclusion is consistent with the nearly ideal thermodynamic behavior of aqueous urea solutions (Ellerton and Dunlop 1966) and the finding, in several molecular simulation studies (Kuharski and Rossky 1984; strand et al. 1994; Vanzi et al. 1998; Kallies 2002), that the intermolecular structure of water is virtually unaffected by urea. Simulations also show that the retardation of water rotation in the urea hydration shell is small, for example, 6% in one study (strand et al. 1994).

The curvature in equation 1Go may be attributed either to overlap of hydration regions, which then would have to extend beyond the first shell (nS= 12.6 corresponds to CU = 3.7 M), or to urea self-association. Molecular simulation studies have provided conflicting results on urea self-association, presumably due to force-field imperfections (Sokolic et al. 2002).

The negligibly small perturbation of water rotational dynamics by urea may be contrasted with that of other small nonelectrolyte solutes (Bagno et al. 1993). Thus, the quantity nS({tau}S/{tau}bulk - 1) is in the range 5–8 for methanol, ethylene glycol, and DMSO, while we obtain 0.32 for urea. This order-of-magnitude difference can be attributed to the dynamic retardation factor ({tau}S/{tau}bulk - 1), because the number nS of water molecules in the hydration shell should vary by less than a factor 2 among these solutes. For proteins, MRD data yield the average of ({tau}S/{tau}bulk - 1) over the heterogeneous surface; typically, this average is in the range 4–5 (Denisov et al. 1996; Halle 1998).

Solvent 17O and 2H relaxation dispersion in I-FABP solutions
The water 17O magnetic relaxation dispersion (MRD) profile R1({omega}0) exclusively monitors the dynamics of water molecules in association with the protein, whereas the 2H MRD profile also contains a pH-dependent contribution from labile hydrogens in the protein that exchange rapidly with the solvent (Denisov and Halle 1995; Halle et al. 1999; Halle and Denisov 2001). In the case of native I-FABP at pH 7, the labile hydrogen contribution appears to be insignificant (Wiesner et al. 1999).

When the solvent contains urea and D2O, hydrogen exchange distributes the 2H nuclei uniformly among water and urea molecules. The 2H magnetization therefore reports on both species. Separate water and urea resonance peaks are observed only at high magnetic fields, where water–urea hydrogen exchange is slow on the chemical shift time scale. Nevertheless, because the exchange remains in the slow to intermediate regime on the relaxation time scale, the individual water and urea 2H relaxation rates can be determined also at low fields from a quantitative analysis of the bi-exponential 2H magnetization recovery (see Materials and Methods).

For most proteins, the water 17O and 2H MRD profiles can be described by a constant term (denoted {alpha}) plus a single Lorentzian dispersion (ß term) with a correlation time {tau}ß that matches the rotational correlation time {tau}R of the protein, which is 7 nsec for I-FABP in water with 50% deuterium at 27°C (Wiesner et al. 1999). For I-FABP and other lipid-binding proteins, the 20–25 water molecules occupying the large internal binding cavity exchange among hydration sites within the cavity on a time scale of 1 nsec (Wiesner et al. 1999; Modig et al. 2003), thus giving rise to a high-frequency dispersion ({gamma} term). Because of the short correlation time ({tau}{gamma} {approx} 1 nsec), only the low-frequency flank of the {gamma} dispersion can be accessed by 17O or 2H MRD. In summary, the MRD profile for native I-FABP is described by a constant plus two dispersive terms (see equation 6Go). The five parameters that define this dispersion profile can be rigorously transformed into well-defined molecular parameters (see Materials and Methods).

Internal hydration of I-FABP during denaturation by urea
17O and 2H MRD profiles were measured in apo I-FABP solutions at pH 7, 27°C, and 10 different urea concentrations from 0 to 8.6 M (see Table 1Go). We shall first discuss the 17O data, which only report on water molecules. The full data set is shown in Figure 3Go. To reduce the number of adjustable parameters, we omit the highest-frequency point in each dispersion profile. This allows us to describe the relaxation data in terms of a single Lorentzian dispersion, ß{tau}ß(1 + [{omega}0{tau}ß]2)-1, and a renormalized constant = {alpha} + {gamma}{tau}{gamma} (see Materials and Methods). The ß parameter can be transformed into (see equation 7bGo). We denote this reduced quantity by and refer to it as the internal hydration parameter. and are the numbers of long-lived (residence time >10 nsec) water molecules in singly occupied cavities and in the large binding cavity, respectively, and the other variables are orientational order parameters with a maximum value of 1 (see Materials and Methods).


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  		Table 1. Composition of MRD samples
 


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  		Figure 3. Water 17O MRD profiles at 27°C in aqueous solutions of apo I-FABP at pH 7.0 with 0–8.6 M urea. The vertical axis measures the excess 17O relaxation rate, R1 - Rbulk, normalized to to remove the effect of slight variations in protein concentration. The curves were obtained from a global fit of all data points to a three-state model (with some of the parameters fixed at plausible values), but is mainly intended as a visual guide.

 
For native I-FABP in the absence of urea, the single-Lorentzian fit yields , in agreement with a previous MRD study (Wiesner et al. 1999). On the basis of an analysis of the 1.2 crystal structure of apo I-FABP (Scapin et al. 1992), this internal hydration parameter can be attributed to singly buried water molecule (W135 in the D–E turn) and water molecules trapped in the large binding cavity (see Fig. 1Go). The single-Lorentzian fit also yields a correlation time {tau}ß = 6.8 ± 0.5 nsec for native I-FABP. This agrees with the rotational correlation time of native I-FABP, {tau}R = 7.2 nsec determined by 15N NMR relaxation (Hodsdon and Cistola 1997) and fluorescence depolarization (Frolov and Schroeder 1997) and scaled to the viscosity (0.968 cP) of our isotope-enriched water.

Figure 4Go shows the variation of the internal hydration parameter with the urea concentration CU along with the far-UV CD denaturation profile, converted to the apparent fraction f of native protein (Santoro and Bolen 1998). The hydration parameter and the combined CD data (at 216 and 222 nm) were analyzed in terms of a two-state denaturation equilibrium N - D with a denaturation free energy linear in CU (see Materials and Methods). The resulting parameters C1/2 and m are given in Table 2Go. Our CD parameters fall within the rather wide range reported from previous CD and fluorescence studies (Ropson et al. 1990; Burns et al. 1998; Dalessio and Ropson 1998, 2000), but do not agree quantitatively with any one of them. Apparently, the salt (and buffer) concentration has a significant effect on the denaturation equilibrium (see Table 2Go).



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  		Figure 4. Variation of the internal hydration parameter (filled circles) during urea-denaturation of 2.3 mM apo I-FABP at pH 7.0 and 27°C. was derived from single-Lorentzian fits to 17O MRD profiles. The ellipticity at 216 nm (+) and 222 nm (x) was measured on 11.5 µM apo I-FABP solutions at 27°C, and is displayed as the apparent native fraction f. The curves resulted from fits according to the standard two-state linear free energy model.

 

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  		Table 2. Results of two-state analysis of I-FABP denaturation curves
 
In line with previous reports, our CD data are well described by a two-state model. When probed by the internal hydration parameter , however, denaturation is seen to require significantly higher urea concentration: C1/2 = 6.5 M versus 5.1 M from CD. Moreover, the data are not well described by a two-state model if is taken to be independent of CU (as was done in the fit of Fig. 4Go). Specifically, increases by nearly one unit in the range 0–3 M urea, where the CD data indicate that the protein is fully native. For native apo I-FABP, is thought to have roughly equal contributions from one singly buried water molecule and 20–25 water molecules in the binding cavity (Modig et al. 2003). The vanishing of at 8.6 M urea (see Fig. 4Go) therefore indicates that the binding cavity has vanished (or, at least, has opened up sufficiently to allow subnanosecond water exchange with the bulk solvent) and that the hydrophobic cluster at the D–E turn has disintegrated (or, at least, has only a transient existence). Our data show that this unfolding takes place at a higher urea concentration than the secondary structure changes probed by CD. Because {alpha}-helices contribute twice as much as ß-strands to the specific ellipticity in the far-UV range (Kelly and Price 1997), the CD data are expected to overemphasize the two short {alpha}-helices (see Fig. 1Go), which may unfold at lower urea concentration than the cooperatively stabilized ß-barrel.

The increase of below 3 M urea may reflect an equilibrium folding intermediate, but could also result from trapping of one or two previously short-lived water molecules by urea molecules in long-lived association with I-FABP. The former explanation is supported by a 1H–15N HSQC NMR study that revealed an intermediate protein structure with maximum population in the range 2.0–3.5 M urea (Hodsdon and Frieden 2001). That study also demonstrated that native-like structural elements persist up to 6.5 M urea, where CD and fluorescence data suggest that the protein is fully denatured. Also, this observation is consistent with our data, which exhibit a denaturation midpoint at 6.5 M. The observation of a substantial 17O dispersion at such high urea concentrations implies that the residual structure is sufficiently permanent to trap water molecules for periods longer than 10 nsec. This residual structure may be related to the equilibrium folding intermediate detected at high urea concentrations (4–7 M) by 19F NMR on fluorinated Trp82 (Ropson and Frieden 1992), the backbone NH of which donates a hydrogen bond to the long-lived internal water molecule (W135) in the D–E turn.

The correlation time {tau}ß obtained from the 17O dispersion can be identified with the tumbling time {tau}R of the protein. Unlike Rbulk (see Fig. 2Go), {tau}ß should therefore be proportional to the solvent viscosity; that is, a hydrodynamic continuum description should apply. To remove the trivial dependence of {tau}ß on the urea concentration CU via the viscosity {eta} (Kawahara and Tanford 1966), we multiply {tau}ß by {eta}(0)/{eta}(CU). For a rigid globular protein, {tau}R is proportional to the hydrodynamic volume. The viscosity-corrected {tau}ß should therefore reflect any global changes in protein structure during denaturation. As seen from Figure 5AGo, the viscosity-corrected {tau}ß hardly varies with urea concentration. This finding is not unexpected. The disappearance of the 17O dispersion at 8.6 M urea (see Fig. 4Go) shows that there are no long-lived water molecules in the fully denatured protein; hence, ß(D) = 0. The frequency-dependent part of R1 is thus entirely due to the native protein fraction f (see equation 10Go). However, this argument is only valid for two-state denaturation. The invariance of {tau}ß in Figure 5AGo therefore tells us that the hydrodynamic volume of the intermediate species indicated by the CU variation of the internal hydration parameter does not differ markedly from that of the native state. Moreover, it shows that the overall structure of the native state is essentially independent of urea concentration. This finding is consistent with previous studies showing that the native structures of BPTI (Liepinsh and Otting 1994) and hen lysozyme (Lumb and Dobson 1992; Pike and Acharya 1994) are essentially unaltered at high urea concentrations.



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  		Figure 5. Variation of (A) the viscosity-corrected correlation time {tau}ß (filled circles) and (B) the apparent surface hydration parameter (open circles) during urea-denaturation of 2.3 mM apo I-FABP at pH 7.0 and 27°C. {tau}ß and were derived from single-Lorentzian fits to 17O MRD profiles. The horizontal line in A corresponds to the independently determined rotational correlation of native I-FABP.

 
Surface hydration of I-FABP during denaturation by urea
The third piece of information obtained from the 17O dispersion is the renormalized parameter = {alpha} + {gamma} {tau}{gamma}, which can be transformed into (see equations 7aGo,cGo). We denote this reduced quantity by and refer to it as the (apparent) surface hydration parameter. In the first term, denotes the number of water molecules in contact with the external protein surface, estimated to 460 for native I-FABP (see Materials and Methods). The dynamic retardation factor measures the relative slowing down of rotational diffusion for these water molecules. For native proteins, (Denisov and Halle 1995; Halle 1998). For native I-FABP, the surface hydration contribution to should therefore be close to .

For native I-FABP in the absence of urea, the 17O dispersion yields . The contribution to from the water molecules trapped in the ligand-binding cavity (the second term in ) is therefore comparable to the surface water contribution. The quadrupole frequency {omega}Q and bulk relaxation rate Rbulk are known (see Materials and Methods and Fig. 2Go) and the correlation time for water exchange among hydration sites inside the cavity is {tau}{gamma} = 1.1 ± 0.1 nsec for native I-FABP (Wiesner et al. 1999; Modig et al. 2003). Combining all this, we obtain for native I-FABP, in agreement with previous MRD studies (Wiesner et al. 1999; Modig et al. 2003).

One might expect to increase with urea concentration as denaturation leads to enhanced solvent exposure (and, hence, larger ). In contrast, Figure 5BGo shows that decreases monotonically across the denaturation transition. This behavior can be understood by recognizing that three different processes contribute to the CU dependence of . Two of these processes are directly linked to the N - D equilibrium. Denaturation greatly increases the solvent-accessible surface area AS (see below), leading to a corresponding increase of the surface contribution to . But denaturation also disrupts the binding cavity, thereby eliminating the second contribution to . These two effects are large, but tend to cancel out.

The third process is the competition of water and urea molecules for surface sites, causing the number of water molecules per unit surface area to decrease with CU. This competition can be taken into account by writing , where NW is the number of external hydration sites on the protein (proportional to AS) and {theta} is the fraction of the surface occupied by bound urea molecules. According to the solvent exchange model (Schellman 1990, 1994), {theta} can be expressed in terms of the mean urea-binding constant KU and the known urea and water activities (see Materials and Methods).

For proteins without large internal cavities, solvent denaturation only involves increase of surface area and solvent competition while thermal denaturation only involves the surface area effect. In such cases, the N -> D transition is clearly reflected in (Denisov and Halle 1998; Denisov et al. 1999). For I-FABP, the near cancellation of the effects of increased surface area and disrupted binding cavity precludes a quantitative analysis of the variation of with urea concentration (see Fig. 5BGo).

Nevertheless, we can extract useful information about the denatured state from the value obtained at 8.6 M urea, where the cavity is disrupted and . We correct for urea competition with the aid of the relation . The urea-binding constant KU is expected to lie in the range 0.05–0.2 M-1 (Pace 1986; Liepinsh and Otting 1994; Schellman and Gassner 1996; Wu and Wang 1999). For this range, equation 12Go yields {theta} = 0.33–0.67, whereby . For KU = 0.1 M-1, . A previous MRD study gave a similar value, , for bovine {alpha}-lactalbumin denatured by guanidinium chloride (Denisov et al. 1999). That protein is nearly the same size as I-FABP (123 versus 131 residues), and was found to be unaffected by cleavage of the four disulfide bonds (I-FABP has no cysteine).

The experimentally derived value can be used as a constraint on models of the denatured state. In particular, a fully solvent-exposed polypeptide chain can be ruled out categorically. For this extreme model, the dynamic retardation should be essentially the same as for an aqueous mixture of amino acids, (Ishimura and Uedaira 1990; Denisov et al. 1999). Note that, because urea has little effect on water dynamics in the bulk solvent (see Fig. 2Go), it should have negligible effect on the relative dynamic retardation factor . With this value and the relation AS/nm2 = 0.15 NW (see Materials and Methods), the experimental constraint yields for the denatured-state solvent-accessible surface area, AS(D) = 0.15 x (6.2 ± 0.6) x 103/(1.3 ± 0.1) = 715 ± 90 nm2. This value greatly exceeds all computational estimates of AS(D) for unfolded models of I-FABP (Miller et al. 1987; Creamer et al. 1997), ranging from 150 nm2 (based on the exposure of the central residue in 17-mer polypeptide segments excised from 43 native protein structures) to 190 nm2 (based on the same polypeptide segments in an extended conformation) to 225 nm2 (based on extended Gly-Xaa-Gly tripeptides), in all cases with a probe of radius 1.4 .

The denatured state of I-FABP must therefore be much more compact than a fully exposed polypeptide chain. It is difficult to be more quantitative, because (solvent-mediated) contacts between polypeptide segments not only reduce NW (or AS), but are also expected to increase . The typical value for native proteins is thought to be strongly dominated by a small number of water molecules in clefts and pockets on the surface, with {tau}S values of several hundred psec (Denisov and Halle 1996; Halle 1998). For a denatured state without rigid and persistent structural constraints, such special hydration sites are improbable. More likely, denatured I-FABP contains a large number of water molecules that are all substantially more perturbed than are water molecules at the surface of the native protein because they act as hydrogen-bond cross-links between polypeptide segments in transient clusters.

Persistent urea binding to the native and denatured states of I-FABP
Up to this point, we have only discussed water 17O MRD data. We now turn to the 2H MRD data, which report on water as well as urea. By explicitly taking into account the slow to intermediate hydrogen exchange between water and urea (Vold et al. 1970; Hunston and Klotz 1971) in the analysis of the magnetization recovery, we could determine the individual water and urea 2H relaxation rates and at most of the investigated urea concentrations (see Materials and Methods). These 2H MRD profiles were then subjected to the same single-Lorentzian analysis as the 17O MRD data.

The reduced parameters , , and {tau}ß derived from the water 2H MRD profiles, and their dependence on CU, conform closely to the corresponding 17O parameters. For example, decreases from 2.2 ± 0.2 in the absence of urea to 0.9 ± 0.3 at CU = 7.5 M (the highest urea concentration investigated by 2H MRD). This agreement indicates that the contribution to from labile hydrogens in the protein is negligible at pH 7, as previously found for the native state (Wiesner et al. 1999). The agreement between the water 2H and 17O parameters also supports the protocol used to separate the water and urea contributions to the 2H magnetization recovery (see Materials and Methods).

Figure 6Go shows urea 2H dispersion profiles at three urea concentrations. The reduced parameters resulting from single-Lorentzian fits are collected in Table 3Go. Within the experimental uncertainty, the viscosity-corrected correlation time {tau}ß does not deviate significantly from the water 17O correlation time (see Fig. 5AGo). As in the case of the water 17O and 2H dispersions, we can therefore identify {tau}ß with the tumbling time {tau}R of the protein. This means that the species giving rise to the urea 2H dispersion has a residence time longer than 10 nsec. In principle, this species could be either bound urea or labile hydrogens in the protein.



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  		Figure 6. Urea 2H MRD profiles at 27°C in aqueous solutions of 2.3 mM apo I-FABP at pH 7.0 with 3.1–7.5 M urea. The figure shows the excess 2H relaxation rate , normalized to to remove the trivial dependence on urea concentration. The curves were obtained from single-Lorentzian fits and the resulting parameter values are given in Table 3Go.

 

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  		Table 3. Results derived from urea 2H MRD parameters
 
A labile-hydrogen contribution to can be ruled for the following reasons. First, the labile-hydrogen contribution to was found to be negligible, and in such a contribution is further reduced by a factor 2 because urea contains twice as many hydrogens as water. Second, sufficiently fast (meaning submillisecond) exchange of labile hydrogens between I-FABP and urea, whether direct or via water, requires catalysis by an ionized species. In the indirect pathway, H3O+ and OH- catalyze both the protein -> water and the water -> urea proton transfer steps. At pH 7, the second step occurs on a time scale of 1 sec (see Materials and Methods), thus effectively eliminating this pathway for magnetization transfer. The direct pathway would presumably be catalyzed by H2NCONH3+, which is present at similarly low concentration as H3O+ (protonated urea has a pKa of 0.1), and is likely to be a less potent catalyst. Consistent with these arguments, exchange between protein and urea was shown to be slow on the NOESY mixing time scale (40 msec) in an intermolecular NOE study of BPTI at pH 7 (Liepinsh and Otting 1994).

On the basis of these considerations, we conclude that the urea 2H dispersion, observed at all investigated urea concentrations from 3.1 to 7.5 M, demonstrates that urea binds to I-FABP with a residence time longer than 10 nsec but shorter than ca. 0.2 msec (the intrinsic relaxation time of urea bound to I-FABP; see Materials and Methods). To our knowledge, this is the first demonstration of such long-lived urea binding to proteins. The dispersion could result from urea molecules trapped in the binding cavity, but then should decrease with increasing urea concentration and vanish at CU = 7.5 M, where the CD data indicate that the cavity is disrupted (see Fig. 4Go). In contrast, we find that increases with CU (see Table 3Go), indicating that urea binds to specific sites in (or on) the native as well as the denatured protein. Because trapping in the large cavity is apparently not involved, we may write , where NI,U is the number of long-lived (specific) urea-binding sites, {theta}I their mean occupancy, and SI,U the orientational order parameter of the bound urea molecule(s). The latter two factors cannot exceed unity, so the values in Table 3Go imply that both the native and denatured forms of I-FABP contain at least one specific urea-binding site.

The increase of with CU does not necessarily indicate a higher affinity for urea in the denatured state, but can be explained by mass action even if the native and denatured states have the same number of specific binding sites with the same urea binding constant. With KU = 0.1 M-1, equation 12Go yields a twofold higher occupancy {theta}I at 7.5 M than at 3.1 M urea. (The maximum in at CU = 5.5 M may be a systematic error; the product increases monotonically with CU.) On the other hand, a residence time longer than 10 nsec implies that KU > 1 x 10-8 kon, where kon is the second-order association rate constant. If urea binding is close to diffusion controlled and/or if the residence time is much longer than 10 nsec, so that KU >> 1 M-1, then the long-lived urea binding site(s) will be essentially saturated at the investigated urea concentrations. The increase of with CU would then suggest a larger number of long-lived urea-binding sites in the denatured state. In any event, the observation of long-lived urea binding to the native and denatured states of I-FABP raises the possibility that strong urea binding contributes significantly to the unfolding thermodynamics and thereby calls into question the validity of the linear extrapolation method widely used to determine the stability of the native protein in the absence of urea (Myers et al. 1995).

Urea–protein interactions have also been studied by other NMR methods than MRD, in particular, intermolecular NOEs and chemical-shift titration. In a study of the small stable protein BPTI, which retains its native structure up to 8 M urea, four urea binding sites were detected in surface pockets and grooves (Liepinsh and Otting 1994) with KU = 0.2 M-1 and residence times of a few nsec at 4°C. In a similar study of the urea-unfolded (7 M) state of the DNA-binding domain of the 434-repressor at -8°C (Dötsch et al. 1995; Dötsch 1996), positive NOESY cross-peaks were observed between urea and most aliphatic protons, indicating urea residence times longer than 0.3 nsec. In both studies, the urea cross-peaks vanished at higher temperatures without exhibiting the expected sign reversal. It should be noted that the model used to transform the sign of the cross-peak into a bound on the residence time may not be appropriate for denatured proteins.

Transient urea interactions with the protein surface during denaturation of I-FABP
The high urea concentrations needed to denature proteins implies that weak binding to many sites is involved. Information about such interactions is contained in the parameter (see Table 3Go). Having rejected the possibility of urea trapping in the large cavity, we can attribute this parameter entirely to urea molecules in short-lived (<1 nsec) association with the external protein surface, so that . To rationalize the observed variation of with CU , we write


(2)

with the mean urea occupancy {theta}(CU) given by equation 12Go (with the same binding constant KU for the native and denatured states) and the native protein fraction f(CU) obtained from equation 11Go. The parameters m and C1/2 may be taken from either the CD or the denaturation curve (see Fig. 4Go; Table 2Go). Because the available data do not allow us to determine all of the three remaining parameters, we fix the value of KU. Acceptable fits are obtained for binding constants in the plausible range 0.05–0.2 M-1 (see Fig. 7Go). For this KU range, the ratio of the two adjustable parameters is . For water, the corresponding ratio is 3.0 ± 0.3 if KU = 0.1 M-1.



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  		Figure 7. Variation of urea–protein interactions during urea-denaturation of 2.3 mM apo I-FABP at pH 7.0 and 27°C. The parameter was obtained from the urea 2H MRD parameter after division by the fractional urea occupancy {theta}, calculated from the solvent exchange model with KU = 0.1 M-1 (see Table 3Go). The curve resulted from a fit where the value of in the native and denatured states were adjusted, while the parameters m and C1/2 characterizing the N - D equilibrium were fixed at the values deduced from the denaturation curve (see Fig. 4Go).

 
The fit in Figure 7Go yields . This may be compared with the corresponding water 17O result, , obtained with the same KU. If we assume that , we can regard the ratio of these numbers as the ratio of water to urea binding sites on the surface of the denatured protein, that is, NU(D)/NU(N) = 1.9 ± 0.3. If water and urea compete for the same space at the polypeptide surface, this should equal the ratio aU/aW of areas occupied by single urea and water molecules at the surface. The latter ratio can be approximated as (U/W2/3). The ratio of partial molar volumes varies from 2.45 to 2.55 in our CU range (Gucker et al. 1938). With U/W = 2.5, we get aU/aW = 1.8. The urea 2H and water 17O MRD data are thus consistent with a similarly large dynamic retardation for urea and water.

To summarize, the urea 2H and water 17O MRD data support a picture of the denatured state where much of the polypeptide chain participates in clusters that are more compact and more ordered than a random coil, but nevertheless, are penetrated by large numbers of water and urea molecules. These solvent-penetrated clusters must be sufficiently compact to allow side chains from different polypeptide segments to come into hydrophobic contact, while, at the same time, permitting solvent molecules to interact favorably with peptide groups and with charged and polar side chains. The exceptional hydrogen-bonding capacity and small size of water and urea molecules are likely to be essential attributes in this regard. In such clusters, many water and urea molecules will simultaneously interact with more than one polypeptide segment, and their rotational motions will therefore be more strongly retarded than at the surface of the native protein. Although the hydrogen-bonding capacity per unit volume is similar for water and urea, the 2.5-fold larger volume of urea reduces the entropic penalty for confining a certain volume of solvent to a cluster. The energetics and dynamics of solvent included in clusters is expected to differ considerably from solvent at the surface of the native protein. This view is supported by the slow water and urea rotation in the denatured state, as deduced from the present MRD data. Further studies are needed to test and refine this tentative picture of the denatured state and to establish whether it applies to a wider range of proteins and denaturing conditions.


   Materials and methods
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 Abstract
 Introduction
 Results and Discussion
 Materials and methods
References
 
Preparation and characterization of protein solutions
Recombinant apo I-FABP was expressed, purified, and delipidated as previously described (Kurian 1998; Wiesner et al. 1999). Lyophilized protein was dissolved in 2H and 17O enriched water (52 atom % 2H, 17 atom % 17O) with 10 mM phosphate buffer. A small fraction insoluble protein was removed by centrifugation. The pH (uncorrected for isotope effects) was 7.00 ± 0.05 in all samples.

The protein concentration was determined, with an estimated accuracy of 5%, from absorbance measurements at 280 nm, using an extinction coefficient of 18.6 mM-1cm-1, calibrated against the complete amino acid analyses performed by Wiesner et al. (1999). Three samples were used for MRD measurements, with I-FABP concentration CP = 2.3–2.4 mM (see Table 1Go). The total number of water molecules per protein molecule was obtained as \mathit|<|N|>|^|<|W|>|_|<|T|>| = (1/[6.022 |<|\times|>| 10^|<||<|-|>|7|>| |<|\times|>| \mathit|<|C|>|_|<|P|>|/mM] |<|-|>| \mathit|<|V|>|_|<|P|>|/^|<|3|>|)/(\mathit|<|V|>|_|<|W|>|/^|<|3|>|), where VP = 18,600 3 is the solvent-excluded protein volume, determined with the program GRASP (Nicholls et al. 1993), and VW = MW/({rho}WNA) = 30 3 is the volume occupied by a single bulk water molecule.

Urea (BDH, ultrapure) was added directly to the NMR samples, and its molar concentration CU was determined from the mass of added urea and the volume of the solution. The pH increase caused by addition of urea was corrected by addition of small volumes of HCl. Because is the ratio between the numbers of water and protein molecules in the sample, it does not change on addition of urea. The total number of urea molecules per protein molecule was obtained by multiplying with the factor xU/(1 - xU), where xU is the mole fraction urea in the solvent. To obtain the mole fraction xU from the molarity CU, we used the following empirical relation for the density d of aqueous urea solutions (Gucker et al. 1938): , with d0 = 0.997 g cm-3 the density of pure H2O at 25°C. Note that the relation between xU and CU is essentially independent of H/D isotope substitution. Table 1Go lists CU, xU, and for the investigated samples.

Far-UV (216 and 222 nm) circular dichroism (CD) denaturation profiles were recorded at 27°C on a Jasco J-720 spectropolarimeter equipped with a Peltier thermostat, using a cell length of 1 mm. The CD samples (pH 7.0) were prepared by mixing a protein solution (approximately 0.2 mM) with appropriate volumes of 10 mM phosphate buffer with or without 10 M urea. The final protein concentration was 9.7 µM.

Magnetic relaxation dispersion measurements
Magnetic relaxation dispersion profiles of the 2H and 17O longitudinal relaxation rate R1 = 1/T1 were acquired for each of the 10 samples. Each dispersion profile comprised nine magnetic field strengths, accessed with the aid of four different NMR spectrometers, including Varian 600 Unity Plus, Bruker Avance DMX 100, and DMX 200 spectrometers and a field-variable iron-core magnet (Drusch EAR-35N) equipped with a field-variable lock and flux stabilizer and interfaced to a Bruker MSL 100 console. The 17O resonance frequencies ranged from 2.2 to 81.4 MHz and the 2H frequencies from 2.5 to 92.1 MHz. The sample temperature was adjusted to 27.0 ± 0.1°C by a thermostated airflow and was checked with a copper-constantan thermocouple referenced to an ice bath.

The relaxation time T1 was measured by the inversion recovery method, using a 16-step phase cycle, 20 delay times in random order, and a sufficient number of transients to obtain a signal-to-noise ratio of at least 100 (Halle et al. 1999). The 17O magnetization recovered as a single exponential and T1 was determined from the standard three-parameter fit. The accuracy of R1(17O) is estimated to ±0.5% (one standard deviation).

Hydrogen exchange between water and urea makes the 2H magnetization recovery bi-exponential. In the absence of exchange, the water and urea 2H magnetizations are assumed to relax exponentially with intrinsic relaxation rates and . In the presence of exchange, the nonequilibrium longitudinal magnetization {Delta}M(t) = MZ(t) - M0 in the two states then evolves according to (Slichter 1989)


(3a)


(3b)

This can be written succinctly as


(4)

where {Delta}M is a column vector formed from the two magnetizations, R is a diagonal relaxation matrix with elements and , and K is an exchange rate matrix with rows [-k1 k-1] and [k1 -k-1]. Because the forward and backward rates must balance at equilibrium, the rate constants are not independent: k1 (1 - PU) = k-1 PU. The urea-deuteron fraction PU is related to the urea mole fraction xU as PU = 2 xU/(1 + xU). The single independent rate parameter is conveniently chosen as the overall exchange rate kex = k1 + k-1.

The formal solution to equation 4Go is {Delta}M(t) = S exp(-Dt) S-1 {Delta}M(0), where S is the matrix that diagonalizes (R - K), that is, D = S-1 (R - K) S. The nonequilibrium magnetization present immediately after the 180° pulse is described by the vector {Delta}M(0) = -M0(1 + {phi}), where {phi} = 1 for an ideal 180° pulse and the elements of M0 can be identified with the relative equilibrium populations PU and 1 - PU. Finally, the observed water and urea 2H magnetizations are computed from and the analogous relation for MU(t), with instrumental scaling factors {sigma}W and {sigma}U. At magnetic fields below 2 T, the water and urea resonances could not be resolved. At these fields, we analyzed the total magnetization MW + MU, taking {phi} and {sigma} to be the same for water and urea. At the three highest fields, separate relaxation experiments were performed on the two resonances, with independent {phi} and {sigma} parameters and different sets of relaxation delays.

The combined water and urea inversion recovery data recorded at all fields were fitted simultaneously with PU and kex as common parameters and , , {phi}W/U, and {sigma}W/U, and as field-dependent parameters (but with common {phi} and {sigma} at low fields). In a typical case, we thus fitted 44 parameters to 240 data points. Figure 8Go shows that this bi-exponential fit substantially improves upon a single-exponential fit. The PU values deduced from the fits agree quantitatively with the urea-deuteron fractions calculated from sample compositions (see Table 1Go). The rate constants k-1 obtained from the fits range from 0.87 sec-1 at 3.1 M to 1.22 sec-1 at 7.5 M urea, in agreement (considering differences in temperature and H/D isotope composition) with previous results (Vold et al. 1970; Hunston and Klotz 1971). The observed increase in k-1 with urea concentration is consistent with the report that acid catalysis is no longer first order in urea concentration above 3 M (Vold et al. 1970). The accuracy of and obtained from these fits is estimated to ±(0.5–1.0)% (one standard deviation) at the three highest fields, with somewhat inferior accuracy at lower fields.



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  		Figure 8. Fits to 2H inversion recovery data from an apo I-FABP solution containing 5.8 M urea. (Left) Sum of the water and urea magnetizations at 0.45 T. (Right) Urea magnetization at 4.7 T. In both panels, the curves resulted from a simultaneous fit to data from all nine magnetic fields. The residuals shown in the upper panels are the differences (in percent) between measured and calculated M(t) for an individual three-parameter single-exponential fit (filled circles) and for the simultaneous bi-exponential fit (open circles).

 
Using the protocol described above, we extracted the 2H relaxation rates and at urea concentrations from 3.1 to 7.5 M (no 2H measurements were done at 8.6 M). In addition, was determined at CU = 0 and 0.5 M, where the urea contribution to the observed R1 is negligible. At CU = 1.1 and 1.8 M, we could not separate and because of the small urea-deuteron fraction (PU {approx} 0.05).

2H relaxation experiments on protein-free reference samples were carried out at 3.1 M, 5.5 M, and 7.5 M urea at the three highest fields, to obtain and. By fitting the measured and rates to cubic polynomials, the bulk relaxation rates were then recalculated at all investigated urea concentrations. For , we also included the relaxation rates at 0 M and 0.5 M urea in the fit. Considering the spread in the data at the investigated concentrations, the error introduced this way is less than 1% for and around 2% for .

Analysis of magnetic relaxation dispersion data
All magnetic relaxation dispersion (MRD) profiles were analyzed with an in-house Matlab implementation of the Levenberg-Marquardt nonlinear {chi}2 minimization algorithm (Press et al. 1992). To estimate the uncertainty in the fitted parameters, we performed fits on a Monte Carlo generated ensemble of 1000 data sets, subject to random Gaussian noise with 0.5% standard deviation for the 17O data. For the 2H data, the standard deviations were set equal to the estimates made above. Quoted uncertainties correspond to a confidence level of 68.3% (one standard deviation).

The water 17O and 2H MRD profiles, R1({omega}0), were modeled by a bi-Lorentzian spectral density J({omega}0) according to (Halle et al. 1999; Wiesner et al. 1999; Halle and Denisov 2001)


(5)


(6)

where {omega}0 = 2{pi} |gu0 is the resonance frequency in angular frequency units and Rbulk is the relaxation rate of the bulk solvent. The five adjustable parameters in equation 6Go were interpreted according to the dynamic cluster model (Modig et al. 2003), a generalization of the standard model (Halle et al. 1999; Halle and Denisov 2001) adapted to proteins with large water-filled internal cavities. This model distinguishes three types of protein-associated water: (1) water molecules in contact with the protein surface (subscript S) are responsible for the frequency-independent {alpha} term, (2) singly buried internal water molecules (subscript I) give rise to the dispersive ß term, and (3) water molecules trapped in the large binding cavity (subscript C) account for the dispersive {gamma} term.

The parameters in equation 6Go are related to the molecular parameters of the model in the following way (Modig et al. 2003):


(7a)


(7b)


(7c)


(7d)