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NMR Relaxation Studies on the Hydrate Layer of Intrinsically Unstructured Proteins

Mónika Bokor ^*, Veronika Csizmók , Dénes Kovács , Péter Bánki ^*, Peter Friedrich , Peter Tompa  and Kálmán Tompa ^*

^* Research Institute for Solid State Physics and Optics, and Institute of Enzymology, Biological Research Center, Hungarian Academy of Sciences, Budapest, Hungary

Correspondence: Address reprint requests to Peter Tompa, Institute of Enzymology, Biological Research Center, Hungarian Academy of Sciences, H-1518 Budapest, PO Box 7, Hungary. Tel.: 361-279-3143; Fax: 361-466-5465; E-mail: tompa{at}enzim.hu.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL MATERIALS RESULTS AND DISCUSSION CONCLUSIONS ACKNOWLEDGEMENTS REFERENCES

 
Intrinsically unstructured/disordered proteins (IUPs) exist in a disordered and largely solvent-exposed, still functional, structural state under physiological conditions. As their function is often directly linked with structural disorder, understanding their structure-function relationship in detail is a great challenge to structural biology. In particular, their hydration and residual structure, both closely linked with their mechanism of action, require close attention. Here we demonstrate that the hydration of IUPs can be adequately approached by a technique so far unexplored with respect to IUPs, solid-state NMR relaxation measurements. This technique provides quantitative information on various features of hydrate water bound to these proteins. By freezing nonhydrate (bulk) water out, we have been able to measure free induction decays pertaining to protons of bound water from which the amount of hydrate water, its activation energy, and correlation times could be calculated. Thus, for three IUPs, the first inhibitory domain of calpastatin, microtubule-associated protein 2c, and plant dehydrin early responsive to dehydration 10, we demonstrate that they bind a significantly larger amount of water than globular proteins, whereas their suboptimal hydration and relaxation parameters are correlated with their differing modes of function. The theoretical treatment and experimental approach presented in this article may have general utility in characterizing proteins that belong to this novel structural class.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL MATERIALS RESULTS AND DISCUSSION CONCLUSIONS ACKNOWLEDGEMENTS REFERENCES

 
Intrinsically unstructured/disordered or natively unfolded proteins (IUPs), common in living organisms, exist in a highly flexible conformational state mostly devoid of well-defined secondary and tertiary structure (Dunker et al., 2002; Tompa, 2002; Uversky, 2002a; Wright and Dyson, 1999). These proteins fulfill essential functions (Dunker et al., 2002; Tompa, 2002, 2003; Wright and Dyson, 1999), intimately linked with the lack of a well-defined structure. In terms of their modes of action, IUPs can be classified into six broad functional categories (Dunker et al., 2002; Tompa, 2002, 2003; Tompa and Csermely, 2004), in five of which they act via permanent or transient binding of a physiological partner, i.e., another protein, DNA, RNA, or some other ligand. In these cases, termed effectors, scavengers, assemblers, display sites, and chaperones, the disordered protein recognizes a structured partner and undergoes induced folding (Demchenko, 2001; Dunker et al., 2002; Dyson and Wright, 2002; Leulliot and Varani, 2001; Tompa, 2002). Such recognition by an initially disordered protein confers exceptional specificity and versatility to the interaction process, which explains the prevalence of structural disorder in signaling and regulatory proteins (Iakoucheva et al., 2002; Ward et al., 2004).

To understand these binding processes in full mechanistic and thermodynamic detail, the quantitative elucidation of the hydration properties of IUPs is needed. For example, their largely open and solvent-exposed structure has to undergo rapid dehydration-hydration cycles to fulfill functions that rely on transient partner recognition, such as of display sites and chaperones (Dunker et al., 2002; Tompa, 2002; Tompa and Csermely, 2004). In concordance, their flexible and adaptable structure abounds in an extended and fully hydrated motif, the polyproline II helix (Syme et al., 2002; Tompa, 2003; Uversky et al., 1998), often implicated in intermolecular recognition processes (Bochicchio and Tamburro, 2002; Williamson, 1994). In addition, hydration and water retention is the very function of certain IUPs involved in dehydration or other types of osmotic stresses (Goyal et al., 2003; Kiyosue et al., 1994; Lisse et al., 1996). Last, but not least, IUPs often are not fully disordered but have permanent or transient global or local structural organization (Tompa, 2002, 2003; Uversky, 2002b). Such a residual structure closely associated with binding functions (Fuxreiter et al., 2004) must manifest itself in suboptimal hydration of the polypeptide chain (Konno et al., 1997).

Here we demonstrate that a technique so far unexplored with respect to IUPs, solid-state NMR relaxation measurements, yields information on the amplitude and dynamics of their hydration processes. The information provided by this NMR technique is complementary to that obtained by other, more often used methods (homo- and heteronuclear nuclear Overhauser effect, relaxation dispersion (nuclear magnetic relaxation dispersion), and spin-spin-lattice relaxation (for reviews, see Antzutkin, 2002; Dyson and Wright, 2004; Otting, 1997; Wider, 1998). We have selected three IUPs for these studies. Calpastatin, the specific inhibitor of the Ca²⁺-activated intracellular cysteine protease, calpain (Emori et al., 1988; Maki et al., 1989), is capable of very rapid and specific interaction with the enzyme. Microtubule-associated protein 2c (MAP2c) binds microtubules (Matus, 1994; Sanchez et al., 2000), and its major function is to ensure proper spacing in the cytoskeleton by long-range entropic repulsion (Chen et al., 1992; Mukhopadhyay and Hoh, 2001). Early responsive to dehydration 10 is a dehydration-stress protein, which shifts the osmotic balance of plant cells due to its putative large hydration capacity (Alsheikh et al., 2003; Kiyosue et al., 1994). To study the hydration of these proteins, their solutions are gradually frozen down, to separate free induction decays (FIDs) of the hydrate layer from those of the overwhelming background of bulk water. The full methodological and theoretical background of how to carry out and interpret these measurements in four different temperature ranges above and below the freezing point has been outlined (Noack, 1971; Racz et al., 1983; Slichter, 1990; Tompa et al., 2001, 2003). As a result, we report the amplitude of hydrate water and dynamic parameters, such as activation energy and various correlation times of the first inhibitory domain of calpastatin (CSD1), MAP2c, ERD10, and bovine serum albumin (BSA) as reference, which are all interpreted in terms of the structure and function of these IUPs.

Theoretical background
Protein solutions are composed of protein molecules, a hydration shell, and unbound water as main components. The protein molecules have rapidly and slowly movable protons, which act as relaxation centers of all the magnetic dipoles. Whereas the rapidly moving centers originate from side chains with rotatable groups or any other flexible parts of the protein, the slowly moving centers correspond to the more rigid backbone. The spin polarization of the water protons is transferred first by rapid material exchange from the bulk solvent to the hydration shell and from there to the relaxation centers by dipolar interaction and spin diffusion. Also, there exists a slight material exchange between the water and protein protons (Noack, 1971). To separate the various water phases present in the samples, the protein solutions are frozen down: the narrow proton signal (time-domain FID) of hydrogen atoms in the ice phase falls within the dead time of the spectrometer. The phases of ice protons, protein protons, and (unfrozen) water protons are clearly separated in the FID signal by virtue of large differences in the spin-spin relaxation rate R₂. Ice protons yield a solid signal fraction with a typical value of R₂ > 200,000 s^–1, which is completely buried in the dead time of the spectrometer. Protein protons yield a solidlike signal fraction also with a large value of R₂ > 20,000 s^–1. The water signal has a much smaller spin-spin relaxation rate, typically R₂ < 2000 s^–1. This enables specific recording of FIDs that belong to the hydrate layer of proteins (Fig. 1). In terms of the behavior of water molecules, four distinct temperature regions (a–d) could be identified, within which the contribution of water phases to the NMR signal show characteristic differences.

View larger version (23K): [in this window] [in a new window]   FIGURE 1  Illustration of the method applied to measure the fraction of the unfrozen water component xunfrozen (MAP2c solution, T = –10.4°C). The slowly decaying part of the time-domain FID signal was extrapolated to t = 0 by applying Lorentzian approximation (dashed line). The extrapolated signal intensity was then compared to the signal intensity measured above 0°C when the whole sample is in liquid state, to get the xunfrozen value. The inset shows the typical spread in time of FID signals produced by ice protons, protein protons, and unfrozen water protons.

Region b
The process of freezing initially acts only upon bulk water outside the hydration shell. Thus, the protein solution can be treated as a two-phase (hydration shell and free water) spin system at about the freezing point, with molecular exchange between the phases. The case of rapid exchange applies if the lifetime of spins in each phase is much shorter than their relaxation times in the given phase. In this case, the relaxation of nuclear magnetization is exponential and

(1)

where R_1av is the measured spin-lattice relaxation rate measured just before freezing, whereas ‘h’ and ‘f’ stand for the hydration shell and free water, respectively. x denotes the probability density of the spins in a phase and (Zimmerman and Brittin, 1957). In this picture, the relative amount of unfrozen water just below the freezing temperature is equal to x_h and the spin-lattice relaxation rate at this temperature is taken as R_1h. The fraction x_h results directly from the FID intensity measurements (see Experimental). The spin-lattice relaxation rate in the free water phase, R_1f, at about the freezing point can then be calculated from the equation above.

Region c
The temperature range from freezing point down to 250 K is characterized by rapidly changing spin-lattice relaxation rates and amounts of unfrozen water in the hydrate layer. The measured spin-lattice relaxation rates can be interpreted as weighted averages of the relaxation rates in different unfrozen water fractions. The measurable relaxation rate R_1M of an n-fraction sample is (Noack, 1971)

(2)

where m_i is the number of protons in the ith fraction and R_1i is the relaxation rate of the ith fraction. The original theoretical description (Noack, 1971) was developed for vacuum dehydration of biological samples at room temperature. It can be applied without change to our case with the only difference that instead of the dehydration process, the freezing process was used to separate the contributions of the different water fractions to R_1M. When the first hydration-shell phase freezes, the first term disappears from the summation. Upon further cooling, the next term becomes zero and so on until only one, quasi-rigid phase is left, which is analyzed in the next section.

Region d
The unfrozen water fraction can be treated as a system with identical ¹H nuclei below 250 K. The amount of unfrozen water changes only slightly in this temperature range. The relaxation rate R₁ is described by the Redfield-Slichter model (Racz et al., 1983). The spin-lattice relaxation in an applied magnetic field B₀ is interpreted as the statistical ensemble average of local field B_loc induced transitions between two Zeeman levels. The dipoles interact with the fluctuating magnetic field B_loc. The spin-lattice relaxation rate formula of this model with isotropic fluctuations is

(3)

where is the gyromagnetic ratio of the ¹H nuclei, ₀ is the Larmor frequency, and is the mean jump time characteristic to the fluctuation of B_loc. The mean jump time obeys the Arrhenius law: The mean squared amplitude (variance) of the isotropic fluctuations is The spin-spin relaxation rate R₂ is given by the equation

The relative magnitudes of the spin-spin (R₂) and the rotating-frame spin-lattice (R₁) relaxation rates compared to the spin-lattice relaxation rate at the temperature of the R₁ maximum is informative with respect to the nature of the correlation time(s). When R₂ and R₁ are greater by one or two orders of magnitude than that predicted by the Redfield-Slichter or the Bloembergen-Purcell-Pound (Bloembergen et al., 1948) model fitted to the R₁ values and the relaxation cannot be described by a single correlation time. Two distinct molecular motions characterized by two different correlation times _s and _r can be assumed to account for the measured relaxation rates. The relaxation rates at the temperature of the R₁ maximum then can be expressed as given in Slichter (1990):

where A and B are the strengths of interactions responsible for the relaxation and _s and _r are the correlation times for slow molecular motion and reorientational fast motions, respectively. The parameters A, B, _s, and _r can be calculated from the above equations at the temperature of the R₁ maximum.

Region b–d
The frozen water (ice) fraction acts as a rigid spin system. Obviously, it is present in increasing amounts when cooling the sample below T_FP. Its NMR signal is characterized by a rapid decay (Barnaal and Lowe, 1967) lost in the dead time of the spectrometer and by a very small spin-lattice relaxation rate. The contribution of ice therefore cannot be detected directly, only its absence is experienced.

	EXPERIMENTAL

TOP ABSTRACT INTRODUCTION EXPERIMENTAL MATERIALS RESULTS AND DISCUSSION CONCLUSIONS ACKNOWLEDGEMENTS REFERENCES

 
DNA constructs
The clones of human calpastatin domain 1 (CSD1, Ala¹³⁷-Lys²⁷⁷ of CST) and rat MAP2c were kindly provided by Prof. M. Maki from Nagoya University (Nagoya, Japan) and Prof. A. Matus from the Friedrich Miescher Institut (Basel, Switzerland). The coding regions were amplified by PCR and subcloned into the NdeI-XhoI sites of the expression vector pET22b (Novagene, Darmstadt, Germany). The gene of ERD10, including an intron, was amplified from Arabidopsis thaliana genomial DNA, with Erd10F (GGAATTCCATATGGCAGAAGAGTACAAGAACAC) and Erd10R (ATAGTTTAGCGGCCGCATCAGACACTTTTTCTTTCTTCTCTTCC) primers. The exons from this primary product were amplified with Erd10F and Erd10R, and two inner primers (Erd10endoF, CCAACAGCTCTTCTTCCTCT TCGAGTGATGAAGAAGGTGAAG and Erd10endoR, CTTCACCTTCTTCATCACTCGAAGAGGAAGAAGAGCTGTTGG, which encode for an overlap in the middle). The exons were then joined by simple annealing PCR, to produce ERD10 cDNA, which was then ligated into pET22b. All the constructs were checked by sequencing at MWG-AG Biotech (Ebersberg, Germany).

Protein purification
CSD1 was expressed in the Escherichia coli strain BL21 and purified on diethylaminoethyl-cellulose anion exchange column as described in (Yang et al., 1994). MAP2c was prepared according to (Ferralli et al., 1994). ERD10 was also expressed in E. coli BL21 and purified by Ni-nitrilotriacetic acid affinity chromatography. Purified proteins were dialyzed into distilled water and dried down by lyophilization. For NMR experiments, the proteins were dissolved on the day of use in buffer 20 mM Tris + 150 mM NaCl + 0.1 mM EDTA at 50 mg/ml (CSD1, MAP2c, and BSA) or 25 mg/ml (ERD10).

NMR spectroscopy
The measurements and data acquisition were accomplished by a Bruker (Rheinstetten, Germany) SXP 4-100 pulse NMR spectrometer of 10^–6 resolution at ₀/2 = 44.14 and 82.57 MHz. Spin-lattice relaxation rates were measured by the inversion-recovery method (Fig. 2). As an addition to the known NMR methods, the standard in situ measurement of ¹H concentration was also solved (Tompa et al., 2003), which is operative in the time range of minutes and enables the simultaneous measurement of NMR relaxation rates and hydrogen content. Determination of the unfrozen water fraction (x_unfrozen) is based on the comparison of the FID signal intensity extrapolated to t = 0 (Fig. 1) with that measured at a temperature where the whole sample is in liquid state. The zero-time FID signal intensity is proportional to the number of resonant nuclear spins in the sample.

View larger version (9K): [in this window] [in a new window]   FIGURE 2  Typical magnetization versus pulse spacing curve of -t-/2 experiments (inversion-recovery method). Spin-lattice relaxation rate R1 was obtained by fitting the equation M0 – M(t) = 2 M0 x exp(–t x R1) (line) to the experimental data (circles).

	MATERIALS

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For DNA purification, the Nucleo-Spin extract kit (Macherey-Nagel, Püren, Germany) was used. BSA and all other chemicals were purchased from Sigma Chemical, St. Louis, MO. Buffers were made in Millipore (Billerica, MA) MilliQ water.

	RESULTS AND DISCUSSION

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Region a spin-lattice relaxation above freezing point
Spin-lattice relaxation was exponential in the studied samples above the freezing point. R₁ data showed simple Arrhenius type behavior, and approximate activation energies (E_a) were calculated (Table 1). The results can be interpreted as mean activation energy values characteristic to the dynamics of water molecules. As seen, the values are within experimental error for the solutions of IUPs and the globular control, i.e., the proteins have a negligible effect on bulk water. The 18–20 kJ/mol value accords with the typical 20 kJ/mol energy of a hydrogen bond in water.

View larger version (15K): [in this window] [in a new window]   FIGURE 3  1H spin-lattice relaxation rate (circles) and unfrozen water fraction (squares) in MAP2c solution (50 mg/ml) at 0/2 = 44.14 MHz. (Solid line) Redfield-Slichter model fitted to R1 data; dotted lines are guides to the eye. SEs are represented by the size of the symbols.

View larger version (15K): [in this window] [in a new window]   FIGURE 4  1H spin-lattice relaxation rate (circles) and unfrozen water fraction (squares) in CSD1 solution (50 mg/ml) at 0/2 = 44.14 MHz. (Solid line) Redfield-Slichter model fitted to R1 data; dotted lines are guides to the eye. SEs are represented by the size of the symbols.

View larger version (15K): [in this window] [in a new window]   FIGURE 5  1H spin-lattice relaxation rate (circles) and unfrozen water fraction (squares) in ERD10 solution (25 mg/ml) at 0/2 = 82.57 MHz. (Solid line) Redfield-Slichter model fitted to R1 data; dotted lines are guides to the eye. SEs are represented by the size of the symbols.

View larger version (14K): [in this window] [in a new window]   FIGURE 6  1H spin-lattice relaxation rate (circles) and unfrozen water fraction (squares) in BSA solution (50 mg/ml) at 0/2 = 44.14 MHz. (Solid line) Redfield-Slichter model fitted to R1 data; dotted lines are guides to the eye. SEs are represented by the size of the symbols.

In qualitative terms, the behavior of this "loose" hydration layer of IUPs correlates with their unstructured nature. First, IUPs seem to have a spatially more extended outer hydration layer, as they lose significantly more of their bound water in this phase (between 260 and 250 K), than BSA (5%–7% vs. 3.5%). This may suggest that their effect on water structure and dynamics extends deeper into space than that of globular proteins. The rapid interaction of IUPs with their partners may stem from their enhanced capture radius (Pontius, 1993; Shoemaker et al., 2000; Tompa, 2002) but also from an increased electrostatic screening effect (Schreiber and Fersht, 1996) that results from their high net charge (Uversky et al., 2000). This screening effect may be related to the observed hydration pattern. The second point of note is that the behavior of CSD1 differs significantly from the other proteins studied. This deviation becomes most apparent below 240 K but is also unmistakable in this range. In effect, the hydrate layer of CSD1 is more heterogeneous than that of the other proteins, as the R₁ value is worst fitted by the Redfield-Slichter model and becomes clearly two componential below 250 K.

In principle, this inhomogeneity might result simply from the heterogeneity of the protein surface, spotted with hydrophobic patches and charged clusters. This, however, would apply equally to the other IUPs studied and would not explain the difference observed. Rather, our contention is that the inhomogeneity reflects residual structure, as it competes with water binding. This is most characteristic of CSD1, shown to have a significant residual structure apparent by local structural preferences (Mucsi et al., 2003; Todd et al., 2003), suboptimal hydration (Konno et al., 1997), and also transient long-range interactions shown by circular dichroism and limited proteolysis (P. Tompa, unpublished observations). This feature appears to ensure that calpastatin is structurally primed for very rapid and effective interaction with calpain (Emori et al., 1988; Maki et al., 1989). Such a structural organization and priming for fast interaction is not required in the case of MAP2c, the major function of which is to ensure spacing in the cytoskeleton (Mukhopadhyay and Hoh, 2001) or ERD10, which is mostly implicated in water binding (Alsheikh et al., 2003; Kiyosue et al., 1994). A final note on this section is that lowering the temperature to 250 K leaves about one equivalent of hydrate water (w/w) mobile, which supports our assertion that from here on only water molecules directly in contact with the protein remain.

Region d "frozen solution"
Below 240–250 K, the amount of unfrozen water changes only slightly and behaves single phased. This region is the most informative when studying the hydration properties since this is the innermost water fraction bound to the protein. Further, the spin-lattice relaxation rates show a maximum in this low-temperature region (Figs. 3–6), and the Redfield-Slichter model can be used for the quantitative evaluation of water dynamics. By fitting Eq. 3 to the measured R₁ data (Table 3), the dynamics of water molecules in the hydration shell of BSA can be characterized by significantly lower activation energy (20 kJ/mol) than that of the IUPs (32 kJ/mol). Thus, IUPs not only bind more water, they also bind the inner hydration layer much more strongly. The related correlation time constants also differ according to the intrinsically unstructured (10^–16 s) or globular (10^–13 s) nature of the protein, perhaps due to more rotational freedom of water molecules on the surface of IUPs than on globular proteins. This marked difference may be of physiological relevance in terms of the rapid rearrangements of hydrate layer that IUPs have to undergo, as outlined in the introduction. This is most apparent with CSD1 (Emori et al., 1988; Maki et al., 1989) but may also apply to MAP2c that binds microtubules (Matus, 1994; Sanchez et al., 2000) and ERD10, in light of its possible membrane association (Bussell and Eliezer, 2003; Koag et al., 2003). The inhomogeneity of CSD1 hydration, pointed out in the previous section, is relevant here, due to the significant residual structure and rapid binding function of this protein.

View this table: [in this window] [in a new window]   TABLE 3  Activation energy (Ea), correlation time constant (0), and average local field values obtained from the Redfield-Slichter model fitted to R1 data below 240–250 K

1

View this table: [in this window] [in a new window]   TABLE 4  Spin-spin (R2) and rotating-frame spin-lattice (R1) relaxation rates and correlation time values (r, s) obtained from the Redfield-Slichter model fitted to R1 data below 240–250 K

	CONCLUSIONS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL MATERIALS RESULTS AND DISCUSSION CONCLUSIONS ACKNOWLEDGEMENTS REFERENCES

	ACKNOWLEDGEMENTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL MATERIALS RESULTS AND DISCUSSION CONCLUSIONS ACKNOWLEDGEMENTS REFERENCES

 
This work was supported by the Hungarian Scientific Research Fund (OTKA; grant T34255) and by the International Senior Research Fellowship GR067595 from the Wellcome Trust. Monika Bokor and Peter Tompa acknowledge the support of the Bolyai János Scholarship.

Submitted on August 27, 2004; accepted for publication December 9, 2004.

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