1 Laboratory of Biochemistry and 2 Laboratory of Biophysical Chemistry, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Nijenborgh 4, 9747AG Groningen, The Netherlands
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Abstract |
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Keywords: disulfide bond engineering/flexible region/haloalkane dehalogenase/molecular dynamics simulation/protein stability
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Introduction |
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Because of its large stabilization potential and relatively well defined conformational requirements, the introduction of disulfide bonds is an attractive method for improving stability, but this strategy has been applied with mixed success. Stability enhancement caused by the introduction of a disulfide is generally attributed to a lowered entropy of the unfolded state (Flory, 1956). This chain entropy model theoretically allows a quantitative prediction of the effect of the introduction of a disulfide bond on the stability (Pace et al., 1988
). However, Doig and Williams (Doig and Williams, 1991
) claim that enthalpic changes of the unfolded state also contribute to the increased stability caused by a cross-link and an increasing amount of evidence indicates that stabilizing effects on the folded state also play a role (Kuroki et al., 1992
; Tidor and Karplus, 1993
; Clarke et al., 1995a
; Vogl et al., 1995
). In addition to the classical thermodynamic approach, Clarke and Fersht (Clarke and Fersht, 1993
) showed that an enzyme can be stabilized kinetically by introducing disulfide bonds between regions involved in the early stages of unfolding. Mansfeld et al. (Mansfeld et al., 1997
) also attributed the extreme stabilization observed in a thermolysin-like protease to a disulfide bond introduced in an area involved in the partial unfolding processes that determine thermal stability.
Although the introduction of disulfide cross-links theoretically should have a stabilizing effect, the opposite is often observed. This destabilization can be attributed to negative effects on packing and flexibility, causing unfavorable enthalpic contributions to offset the entropic profit gained by the cross-link. With the disposition of high-resolution structures, substantial information has become available concerning the spatial requirements of disulfide bonds (Thornton, 1981). This resulted in the development of computer programs that can predict suitable sites for their introduction (Pabo and Suchanek, 1986
; Hazes and Dijkstra,1988; Sowdhamini et al., 1989
). Although these programs have facilitated the search for potentially successful substitutions to a large extent, introduction of a stabilizing disulfide bond remains a matter of trial and error if it is not possible to identify regions in the enzyme that are important for stabilization. The earliest reports on successful introductions of disulfide bonds exploited native unpaired cysteine residues, making the choice for the position of the second cysteine rather straightforward (Villafranca et al., 1983
; Perry and Wetzel, 1984
). When stabilization by de novo introduction of both cysteines was shown to be feasible (Pantoliano et al., 1987
), the number of theoretically possible disulfide bonds again outnumbered the practical possibilities. For haloalkane dehalogenase (Dhla) the program SSBOND (Hazes and Dijkstra, 1988
) found as many as 59 amino acid pairs that fulfilled the positional constraints for introduction of a disulfide bond (see Results). Comparison with related, sometimes more thermostable variants of the protein can lead to a successful choice (Takagi et al., 1990
; Yamaguchi et al., 1996
; Li et al., 1998
), but in most cases the selected location is based on an `educated guess'.
Clearly, at present, a bottleneck in designing more stable proteins is the lack of accurate methods to determine the positions leading to an effective stabilization. Secondary structure elements such as helices and sheets will often complicate efficient disulfide bond formation because of their rather tight steric constraints (Thornton, 1981). Thus, in order to obtain a disulfide bond that positively contributes to the stability of a protein, cross-linking should preferably involve a flexible region (Matsumura et al., 1989
). This will probably have a smaller destabilizing effect on the folded conformation since the adjacent residues are more likely to adjust in order to compensate for negative strain effects. Disulfide introduction in T4 lysozyme demonstrated that the most stabilizing disulfide cross-links were situated in flexible regions as indicated by the temperature factors of the native X-ray structure (Matsumura et al., 1989
). Subsequent analysis of the X-ray structure of one of these mutants (Pjura et al., 1990
) indeed revealed considerable shifts in main chain atoms of neighboring residues and only a slight strain on the disulfide bond itself. The cross-link did not impose rigidity on the residues that showed flexibility in the native enzyme. Weber et al. (Weber et al., 1985
) reported a similar observation after introduction of a chemical cross-link in ribonuclease A.
A powerful method to investigate dynamic properties of a protein at the atomic level is provided by molecular dynamics (MD) simulations. Modern computers allow simulations of several nanoseconds, which is long enough to determine the dominant contributions to atomic fluctuations. Therefore, MD simulations should provide an attractive means to identify flexible regions in proteins that can serve as a target for stability enhancement. Using Dhla as a model, we exploited the possibility of identifying such regions and studied the effect of cross-linking on stability.
Dhla from Xanthobacter autotrophicus GJ10 (DhlA) catalyzes the hydrolytic dehalogenation of a range of small halogenated alkanes (Keuning et al., 1985). From the X-ray structure (Franken et al., 1991
) the enzyme appears to belong to the
/ß-hydrolase fold family. The members of this family share a common main domain, consisting of an eight-stranded ß-sheet surrounded by
-helices, on which the position of the catalytic residues is preserved (Ollis et al., 1992
). Most of the
/ß-hydrolase fold enzymes show an excursion between ß-strands 6 and 7, which appears to be highly diverse in size and structure. In Dhla this second domain consists of five
-helices and is called the cap domain, since it more or less shelters the active site that is located in a hydrophobic cavity on the interface of the two domains. Previous work on DhlA showed that the cap domain is very important in determining substrate specificity (Pries et al., 1994
; Schanstra et al., 1996
, 1997
) and also indicated that flexibility of this domain is probably controlling the release of the halide ion, which is the main rate-limiting reaction step in the conversion of 1,2-dichloroethane (Schanstra and Janssen, 1996
; Krooshof et al., 1999
).
In this report we describe the results of the MD simulation of Dhla with a chloride ion bound in the active site. The results indicate the presence of a mobile helixloophelix region in the cap domain, involving residues 184211. Introduction of a disulfide bond between residues 201 and 16, anchoring this flexible region to the main domain, led to an enhanced stability towards both thermal and chemical denaturation.
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Materials and methods |
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The MD simulation was performed using the GROMOS87 forcefield (van Gunsteren and Berendsen, 1987) with adjusted LennardJones interactions between aliphatic carbon atoms and water oxygen atoms (van Buuren et al., 1993
) and explicit hydrogens added to aromatic residues (van der Spoel et al., 1996b
). Polar hydrogens were included explicitly, whereas non-polar hydrogens were implicitly included by the use of united atoms. For all but the initial simulations, a simple form of polarization was included in the force field. This was necessary because in an initial simulation the chloride ion, present in the starting structure, left the active site almost instantaneously (within 20 ps). This is due to the fact that the interaction of the chloride ion with its hydrophobic environment is too weak in the force field used.
A direct charge-induced dipole polarization term:
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The polarization term adds >30% to the stabilization energy of chloride (in a typical structure from the simulation the polarization energy added 26.8 kcal mol1 to the Coulomb energy of 72.9 kcal mol1 between chloride and protein atoms) and keeps the chloride ion in place.
As a starting structure, the X-ray structure at pH 6 with a resolution of 2.1 Å (Brookhaven Data Bank entry 2DHE) was used (Verschueren et al., 1993b). To compensate for the charge of the chloride ion, His298 was assumed to be protonated at both ring nitrogens. All other histidines were assumed to be electrically neutral and protonated at only one ring nitrogen. To select the protonated nitrogen, the structure was checked for the presence of possible hydrogen bonds by looking at the closest hydrogen bond acceptor. Histidine residues 37 and 54 were protonated at the N
1 position, and residues 102 and 305 at the N
2 position. Periodic boundary conditions were applied by the use of a truncated octahedron filled with equilibrated water molecules. Each water molecule of which the oxygen atom had a distance to any non-hydrogen protein atom or water molecule of <2.3 Å was removed. The minimum distance between protein atoms and the walls of the box was taken as 7.5 Å and the volume of the box was 174 000 Å3. The protein (including the chloride ion) possessed a negative charge of 17 e. To obtain an electrically neutral system, 17 sodium ions were added as counterions. This was done by calculating the electric potential at the positions of all oxygen atoms of water molecules and replacing 17 water molecules with the lowest potential by a sodium ion. In total the system contained 16 047 atoms of which 3171 were protein atoms (including the chloride ion).
An energy minimization was performed using the steepest descent method for 100 steps without position restraining. During the first 10 ps of the simulation, harmonic position restraints were applied to all protein atoms, using a force constant of 9000 kJ nm1 mol1. After that, the position restraints were removed and the system was initially simulated for 1 ns and later continued for another 500 ps.
Initial velocities were taken from a MaxwellBoltzmann distribution at 298 K. The temperature was kept constant by coupling solute and solvent separately to a thermal bath at 298 K with a coupling constant T = 0.1 ps. Pressure was kept constant by coupling to a pressure bath at 1.0 bar, using a coupling constant
P = 0.5 ps (Berendsen et al., 1984
). Bond lengths were constrained using the SHAKE method with a relative tolerance of 0.0001 (Ryckaert et al., 1977
). Non-bonded interactions were calculated using a twin cut-off radius: within a short cut-off radius of 8 Å Coulomb and LennardJones interactions were calculated every time step, whereas Coulomb interactions within a radius of 12 Å were calculated every 10 steps. The step size was 0.002 ps. The behavior of the secondary structure elements was calculated with the program DSSP (Kabsch and Sander, 1983
). B-factors derived from the trajectory were defined as:
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The simulation was performed using the GROMOS87 software package (van Gunsteren and Berendsen 1987). Non-bonded routines were rewritten from scratch for use on a Cray J90 parallel computer. The simulation took
200 h of real time per nanosecond. Analysis of secondary structure elements, root mean square deviation (RMSD) and B-factors was performed with GROMACS (van der Spoel et al., 1996a
).
Design of the disulfide bond
To select a suitable site for the introduction of a disulfide bond connecting the helixloophelix region and the main domain the disulfide bond prediction computer program SSBOND (Hazes and Dijkstra, 1988) was used. As a starting structure the X-ray structure at pH 8.2 and a resolution of 1.9 Å (Brookhaven Protein Data Bank entry 1EDE) was used (Verschueren et al., 1993a
). The program selects potential sites based upon a series of conformational and energy constraints derived from disulfide bonds in proteins for which a three-dimensional structure has been determined. The program allows user input to adjust the energy constraints, so disulfides that do not exactly match any of the known disulfide conformations can be generated. The DhlA mutant Asp16Cys/Ala201Cys was generated after allowing the Cß1S
1S
2Cß2 dihedral angle to deviate ±10°.
Mutagenesis
The single mutants Asp16Cys and Ala201Cys were obtained according to Kunkel (Kunkel, 1985). Escherichia coli strain BW313 was used for the production of single-stranded uracil containing DNA of plasmid pGELAF(+), an expression vector based on pET-3d (Studier et al., 1990
) with the dehalogenase gene under control of the T7 promoter (Schanstra et al., 1993
). The primer used for the Ala201Cys mutation was TGACCGAATGCGAGGC, with the mutated codon in bold and the introduced BsmI site underlined. For the Asp16Cys mutation the following primer was used: GCAATCTCTGCCAGTATCC. Successful mutations could be detected by the absence of a BsaBI site caused by this primer. After separate introduction of each mutation the double mutant was obtained by ligating the appropriate BclIPstI DNA fragments, which were isolated from E.coli strain KA1271, a dam- strain. The oligonucleotides for mutagenesis were supplied by Eurosequence BV (Groningen, The Netherlands). Sequences of mutated genes were established by the Center for Biomedical Technology, University of Groningen (The Netherlands).
Expression and purification
For the expression of the double mutant we used E.coli strain BL21(DE3) (Studier et al., 1990), a strain expressing the T7 RNA polymerase of bacteriophage T7. After overnight growth at 30°C, the transformed E.coli cells were collected from plates, resuspended in 1 ml of LB medium, and used to inoculate 100 ml of LB (containing 100 µl/ml ampicillin) to an OD600 of 0.050.1. This preculture was grown at 30°C until it reached an OD600 of
1.0 and was added to the 1 l main culture. At an OD600 of 1.0, IPTG (0.4 mM) was added and growth was continued for 15 h at 20°C. Crude extract was prepared as described earlier (Schanstra et al., 1993
). After passage through a 0.2 µm filter, the cell extract was applied to a ResourceTMQ column (Pharmacia Biotech) on a Pharmacia liquid chromatography controller LCC-500. The buffer used during purification was TEMAG [10 mM TrisSO4, pH 7.5, 1 mM EDTA, 1 mM ß-mercaptoethanol, 3 mM sodium azide and 10% (v/v) glycerol]. Bound protein was eluted using TEMAG buffer with a gradient of 00.5 M ammonium sulfate. Purified protein was stored at 4°C in TEMAG without ammonium sulfate.
Verifying the formation of the disulfide bond
Purified protein was analyzed for disulfide bond formation by comparing reduced and non-reduced samples with SDSpolyacrylamide gel electrophoresis (PAGE) using Coomassie brilliant blue staining. Prior to the gel electrophoresis the protein samples were incubated at room temperature for at least 15 min in denaturation buffer [15 mM TrisCl (pH 6.8) containing 5% SDS, 25% glycerol and 0.05% bromophenol blue] which was supplemented with 15% ß-mercaptoethanol if the reduced form of the protein was analyzed. Free sulfhydryl groups were quantitatively determined using 5,5'-dithiobis (2-nitrobenzoic acid) (DTNB) (Ellman, 1959). This compound reacts stoichiometrically with the free sulfhydryl groups of the enzyme and subsequent release of the aromatic thiolate can be followed quantitatively by measuring the absorbance at 412 nm. The reactions with DTNB were performed by mixing 900 µl of enzyme (0.25 mg/ml in TEAG) with 100 µl of 0.13% (w/w) DTNB in 50 mM Tris buffer, pH 8.2. Incubations at room temperature were continued until the absorbance at 412 nm had stabilized. The number of reacting sulfhydryl groups was calculated from the amount of released 2-nitro-5-thiobenzoic acid using a calibration curve with cysteine.
To establish the formation of the correct disulfide bond and for the identification of the DTNB-modified sulfhydryl groups, trypsin digestions were performed. Prior to the digestion, the protein was dialyzed against 0.2 M ammonium acetate/ammonium bicarbonate buffer (pH 8). Complete digestion of 1 nM protein was obtained after incubating for 4 h at 37°C using 1% (w/w) TCPK-grade trypsin (Worthington Biochemical Corp., Lakewood, NJ). Peptide fragments were analyzed by reversed-phase HPLC using a 250x4.6 mm Nucleosil 100 C854 column (Alltech, Deerfield, IL). For identification, peptide fragments containing the DTNB-modified sulfhydryl groups, liquid chromatographymass spectrometry (LCMS) was used as described earlier (Pries et al., 1995).
Stability determination
Protein stability was analyzed by determining the residual activity as a function of temperature as well as urea concentration. Thermal stability was assayed by following the loss of activity during controlled increase of temperature. Identical aliquots of 50 µg of enzyme in 100 µl of TrisSO4 (pH 8.2) were heated in a thermocycler (Progene, Techne; New Brunswick Scientific, Edisol, NJ) at a rate of 1°C/min. At definite temperatures a vial was removed and immediately cooled on ice, after which the residual activity was determined.
Urea denaturation was assayed by incubating 20 µg of protein in 200 µl of 50 mM TrisSO4 (pH 8.2) with a certain urea concentration. Residual activity was determined after an incubation period of 1.5 h at room temperature. For stability determination of the reduced mutant the TrisSO4 buffer was supplemented with 100 mM dithiothreitol (DTT).
Dehalogenase activities were determined by measuring the rate of alcohol production in time. Thermal and urea denaturation samples were incubated at 30°C with 5 mM DCE as substrate. After a fixed reaction period the alcohol product was extracted with diethylether containing 0.05 mM bromohexane as the internal standard. The ether layer containing alcohol and substrate was analyzed on a Chrompack 438S gas chromatograph using a CPWax 52CB column (Chrompack, Middelburg, The Netherlands), coupled to an FID detector (Schanstra et al., 1996). Residual activity was expressed as a percentage of the initial activity and the apparent denaturation transition temperature was obtained using a sigmoid function from a curve-fitting program (Sigma Plot; Jandel Scientific, San Rafael, CA).
Kinetic stability was measured by following fluorescence changes after rapidly mixing the enzyme with denaturing urea concentrations. The experiments were performed on an Applied Photophysics model !SX17MV stopped-flow spectrofluorometer. The excitation wavelength was 290 nm and subsequent fluorescence emission was detected through a 320 nm cut-off filter. Reactions were performed at 30°C and initiated by mixing one volume of protein (1 mg/ml) with 10 volumes of a concentrated urea solution in TrisSO4 (pH 8.2). All fluorescence traces could be fitted to a single exponential from which the rate constant of unfolding ku was obtained.
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Results |
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To identify potentially flexible regions in DhlA, an MD simulation of the enzyme was performed. During the first 1 ns of the simulation the RMSD from the crystal structure showed a steady increase. Since this might suggest that the structure was starting to unfold, the simulation was continued for another 500 ps. After 1 ns the RMSD stabilized around a value of 0.35 nm, to increase again after
1.4 ns (Figure 1
). Close inspection of the trajectory revealed that after
1.25 ns some (charged) residues from the protein in the central periodic box started to interact with residues from one of the protein's images. The stabilization after 1 ns indicated that the structure remained intact, but we only considered the trajectory until 1.2 ns as being reliable and restricted further analysis to this time interval.
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To study the effect of rigidifying the mobile part of the enzyme, a disulfide bond was introduced between this most flexible helixloophelix region and the main domain. Wild-type Dhla contains four cysteine residues, located at positions 52, 150, 233 and 278, but none of them is involved in a disulfide bond. Using the computer program SSBOND (Hazes and Dijkstra, 1988) residue pairs were identified that might be capable of forming a disulfide bond when they would be replaced by a cysteine. A primary search using standard values for the structural parameters of the various torsion angles and SS bond length revealed 59 possible candidates. Eight pairs had one or two residues within the mobile helixloophelix region, but none of them would result in a cap domain/main domain connection. This result reflects the small number of H bonds and electrostatic interactions between the two domains: in wild-type enzyme only 13 hydrogen bonds and four salt bridges are found (Verschueren et al., 1993a
). The main force to keep the domains together is probably of hydrophobic nature. A second search, in which the Cß1S
1S
2Cß2 dihedral angle was allowed to deviate 10° from its standard value of ±90°, revealed 19 additional pairs. Two of them satisfied the desired positional criteria: the Asp16/Ala201 and the Ala195/Phe290 pair. Since Phe290 is flanking the conserved catalytic residue His289, changing this amino acid into a cysteine residue participating in a disulfide bond would probably affect the catalytic properties of the enzyme. Therefore, we chose to introduce a disulfide bond between the residue positions 16 and 201 (Figure 5
).
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Disulfide bond formation
After purification, the presence of a disulfide bond between the two mutated residues was confirmed by analyzing the mobility differences in non-reducing and reducing SDSPAGE. The introduction of a disulfide bond is known to reduce the radius of gyration of the protein, causing the oxidized sample to move faster through the gel (Pollitt and Zalkin, 1983). Whereas under reducing conditions the bands of wild-type enzyme and mutant were indistinguishable, the non-reduced mutant enzyme moved faster through the gel, indicating the presence of a disulfide bond. Under these non-reducing conditions the wild-type reference sometimes showed multiple bands, indicating adventitious disulfide bond formation. Additional HPLC analysis of trypsin digests of the oxidized mutant enzyme showed the appearance of a large peak upon the loss of peak T3 and T18, which have been assigned to the fragments containing amino acids 1235 and 194220 (Krooshof, 2000
). This indicated that the disulfide bond in the mutant was indeed formed between Cys16 and Cys201. The identity of the adventitious bands in wild-type enzyme could not be established. SDSPAGE analysis of mutant protein samples incubated with different reductant concentrations revealed that at least 100 mM ß-mercaptoethanol was needed for complete reduction of the disulfide bond. Because disulfide bonds are rarely formed in E.coli cytosol due to the strongly reducing environment (Ziegler and Poulsen, 1977
), we assume that the disulfide bond forms spontaneously during cell lysis and purification, since the ß-mercaptoethanol concentration in the purification buffer is only 1 mM.
Detection of free thiol groups with DTNB (Ellman, 1959) revealed unexpected long-range structural consequences of the mutations. Although none of the native cysteine residues is located in the vicinity of the mutated residues, quantitative analysis showed that in the oxidized form of Asp16Cys/Ala201Cys only three out of four free sulfhydryl residues were modified with DTNB, whereas in the wild-type enzyme all four cysteine residues were reactive. DTNB reacts with the conjugate base of a thiol, so the rate of reaction varies with the pH and the pKa of the thiol. Apart from the presence of a disulfide bond, non-stoichiometric reactivity can also be caused by inaccessibility of the sulfhydryl group, for example, caused by a buried position inside the protein or complex formation with metal ligands (Chu et al., 1996
; Pregel and Storer, 1997
). To determine which of the cysteine residues was no longer capable of reacting with DTNB, we analyzed trypsin digests of the DTNB-modified mutant enzyme with LCMS. Only the DTNB-modified peptide fragments T4 and T25, containing Cys52 and Cys278, respectively, could be detected. This implies that either Cys150 or Cys233 is in the oxidized form of the mutant enzyme no longer accessible for DTNB modification. This non-reacting sulfhydryl residue could very well be one of the residues involved in the adventitious disulfide bond formation observed in the wild-type enzyme, since this never occurred in the oxidized mutant.
Thermostability
The thermal denaturation of DhlA is not a reversible process (Krooshof, 2000). This irreversibility precludes a formal and detailed study of the thermodynamics of unfolding. Therefore, thermal denaturation of wild-type enzyme and both forms of the mutant enzyme was analyzed by measuring the residual activity of identical samples that were subjected to a gradual temperature increase. To be able to compare the oxidized and reduced form of the mutant with wild-type enzyme, the apparent denaturation transition value Tm,app was defined as the temperature at which the residual activity had decreased to 50% of the initial value.
The denaturation curves show that the introduction of the disulfide bond leads to a substantially increased thermostability (Figure 6). The wild-type enzyme shows an apparent transition temperature of 47.5 ± 0.5°C, whereas the Tm,app of the oxidized mutant has increased to 52.7 ± 0.5°C. These results are in good agreement with temperature-induced denaturation monitored by circular dichroism (not shown), indicating that no renaturation takes place during the assay process. The reduced mutant enzyme is obviously less stable than the wild-type enzyme and has a Tm,app of 39.0 ± 0.5°C. The slope of the curve, which is a measure of the cooperativity of the unfolding process, is significantly lower for the reduced mutant enzyme. For a reversible process the slope at Tm is proportional to the unfolding enthalpy, but since the denaturation of Dhla is not reversible, the thermodynamic parameters of the unfolding process cannot be determined.
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Since products of thermal denaturation are often less extensively unfolded than those of urea unfolding (Pace, 1975), the stability of DhlA was further examined by analysis of the residual activity of enzyme samples exposed to various concentrations of urea (Figure 7
). Under the applied assay conditions, wild-type enzyme showed a rather sharp transition between 0.75 and 1.5 M urea, whereas the reduced form of the Asp16Cys/Ala201Cys mutant already started unfolding at very low urea concentrations. However, due to a much more gradual transition, complete unfolding occurred at only slightly lower urea concentrations than for wild-type enzyme. The oxidized form of the mutant was considerably more stable. This enzyme started unfolding at 1.25 M and for complete unfolding a concentration of 2 M urea was required.
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Kinetic stability
The mobility that was observed in the MD simulation of DhlA indicates that helices 6 and 7 of the cap domain may be involved in the early stages of unfolding or even may represent the area where unfolding is initiated. Engineered disulfide bonds can be used as probes to follow unfolding and refolding and they are able to decrease the rate of unfolding by stabilizing early-unfolding elements (Clarke and Fersht, 1993). Therefore, we investigated the kinetic stability of wild-type enzyme and both forms of the Asp16Cys/Ala201Cys mutant.
Unfolding rates were measured using stopped-flow fluorescence. Rapid mixing of the enzyme with denaturing urea concentrations revealed single exponential fluorescence traces from which the rate constant of unfolding (ku) could be obtained. The dependence of the rate constant for unfolding on urea concentrations can be described as (Clarke and Fersht, 1993):
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The increase in activation free energy G
-F caused by the introduction of the disulfide bond is 0.43 kcal mol1 compared to the wild-type enzyme and 0.88 kcal mol1 compared to the reduced form of the mutant.
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Discussion |
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Although crystallographic B-factors can also provide information on flexibility, the mobility of helices 6 and 7 detected in the MD simulations could not be observed from these B-factors (Figure 2). This is probably caused by the structural arrangement of the molecules in the crystal lattice (Verschueren et al., 1993a
). The great majority of crystal contacts involve cap domain residues interacting with main domain amino acids of a neighboring molecule. This results in B-factors similar to those of the ß-sheet; the average B-factors of the ß-sheet and of the cap domain helices are, respectively, 11.5 and 10.0 Å2, whereas the helices of the main domain have a mean B-value of 15.3 Å2. Solvent accessibility data of the cap domain already gave an indication that the observed rigidity was at least partly artificial, since these values did not show the usual correlation with crystallographic B-factors (Verschueren et al., 1993a
). The current results clearly emphasize the limitations of X-ray crystallography in determining flexibility of proteins in solution. The need for a more dynamical approach was also nicely illustrated by the results of a hydrogen exchange study on a disulfide bond mutant of barnase (Clarke et al., 1995b
). Although the crystallographic B-factors in this region indicated an increased mobility in the mutated area, the increased lifetime of the hydrogen bonds showed that the local stability had actually increased due to the introduction of the cross-link.
Theoretically, the reduction in entropy resulting from a cross-link in a random-coil peptide can be calculated using Equation (3) (Pace et al., 1988
):
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Our results show that introduction of the disulfide cross-link in DhlA enhanced the kinetic stability of the enzyme, suggesting that the cap domain region encompassing helices 6 and 7 is probably involved early in the unfolding process. Theoretically, the cross-linking of flexible regions could have a destabilizing effect on the native state, since it lowers the entropy of the protein. However, in the case where the regions connected by the disulfide are far apart from each other in the transition state, this state of the protein will be destabilized even more, resulting in a net stabilization of the native form. The results indicate that MD simulations may be a valuable tool for identifying flexible regions involved in the early stages of the unfolding process. The introduction of a disulfide bond in such a region can hamper or block the pathway to the transition state, slowing down the overall unfolding process. Therefore, MD simulations combined with the introduction of disulfide cross-links provide a promising tool for enhancing the kinetic stability of proteins.
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Notes |
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Acknowledgments |
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References |
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Applequist,J., Carl,J.R. and Fung,K. (1972) J. Am. Chem. Soc., 94, 29522960.[ISI]
Berendsen,H.J.C., Postma,J.P.M., van Gunsteren,W.F., DiNola,A. and Haak,J.R. (1984) J. Chem. Phys., 81, 36843690.[CrossRef][ISI]
Chu,Y., Lee,E.Y., Reimann,E.M., Wilson,S.E. and Schlender,K.K. (1996) Arch. Biochem. Biophys., 334, 8388.[CrossRef][ISI][Medline]
Clarke,J. and Fersht,A.R. (1993) Biochemistry, 32, 43224329.[ISI][Medline]
Clarke,J., Henrick,K. and Fersht,A.R. (1995a) J. Mol. Biol., 253, 493504.[CrossRef][ISI][Medline]
Clarke,J., Hounslow,A.M. and Fersht,A.R. (1995b) J. Mol. Biol., 253, 505513.[CrossRef][ISI][Medline]
CRC Handbook of Chemistry and Physics (1986) 67th edn. Lide,D.R. ed. CRC Press Inc., Boca Raton, FL.
Doig,A.J. and Williams,D.H. (1991) J. Mol. Biol., 217, 389398.[ISI][Medline]
Ellman,G.L. (1959) Arch. Biochem. Biophys., 82, 7077.[ISI][Medline]
Flory,P.J. (1956) J. Am. Chem. Soc., 78, 52225235.[ISI]
Franken,S.M., Rozeboom,H.J., Kalk,K.H. and Dijkstra,B.W. (1991) EMBO J., 10, 12971302.[Abstract]
Greene,R.F.,Jr and Pace,C.N. (1974) J. Biol. Chem., 249, 53885393.
Hazes,B. and Dijkstra,B.W. (1988) Protein Eng., 2, 119125.[Abstract]
Kabsch,W. and Sander,C. (1983) Biopolymers, 22, 25772637.[ISI][Medline]
Keuning,S., Janssen,D.B. and Witholt,B. (1985) J. Bacteriol., 163, 635639.[ISI][Medline]
Krooshof,G.H., Floris,R., Tepper,A.W.J.W. and Janssen,D.B. (1999) Protein Sci., 8, 355360.
Krooshof,G.H. (2000) Structurefunction Relationships in Haloalkane Dehalogenase. Thesis, University of Groningen, The Netherlands.
Kunkel,T.A. (1985) Proc. Natl Acad. Sci. USA, 82, 488492.[Abstract]
Kuroki,R., Inaka,K., Taniyama,Y., Kidokoro,S., Matsushima,M., Kikuchi,M. and Yutani,K. (1992) Biochemistry, 31, 83238328.[ISI][Medline]
Li,Y., Coutinho,P.M. and Ford,C. (1998) Protein Eng., 11, 661667.[Abstract]
Mansfeld,J., Vriend,G., Dijkstra,B.W., Veltman,O.R., Van den Burg,B., Venema,G., Ulbrich-Hofmann,R. and Eijsink,V.G. (1997) J. Biol. Chem., 272, 1115211156.
Matsumura,M., Becktel,W.J., Levitt,M. and Matthews,B.W. (1989) Proc. Natl Acad. Sci. USA, 86, 65626566.[Abstract]
Ollis,D.L., Cheah,E., Cygler,M., Dijkstra,B., Frolow,F., Franken,S.M., Harel,M., Remington,S.J., Silman,I., Schrag,J. et al. (1992) Protein Eng., 5, 197211.[Abstract]
Pabo,C.O. and Suchanek,E.G. (1986) Biochemistry, 25, 59875991.[ISI][Medline]
Pace,C.N. (1975) CRC Crit. Rev. Biochem., 3, 143.[Medline]
Pace,C.N., Grimsley,G.R., Thomson,J.A. and Barnett,B.J. (1988) J. Biol. Chem., 263, 1182011825.
Pantoliano,M.W., Ladner,R.C., Bryan,P.N., Rollence,M.L., Wood,J.F. and Poulos,T.L. (1987) Biochemistry, 26, 20772082.[ISI][Medline]
Patkar,S., Vind,J., Kelstrup,E., Christensen,M.W., Svendsen,A., Borch,K. and Kirk,O. (1998) Chem. Phys. Lipids, 93, 95101.[CrossRef][ISI][Medline]
Perry,L.J. and Wetzel,R. (1984) Science, 226, 555557.[ISI][Medline]
Pjura,P.E., Matsumura,M., Wozniak,J.A. and Matthews,B.W. (1990) Biochemistry, 29, 25922598.[ISI][Medline]
Pollitt,S. and Zalkin,H. (1983) J. Bacteriol., 153, 2732.[ISI][Medline]
Pregel,M.J. and Storer,A.C. (1997) J. Biol. Chem., 272, 2355223558.
Pries,F., van den Wijngaard,A.J., Bos,R., Pentenga,M. and Janssen,D.B. (1994) J. Biol. Chem., 269, 1749017494.
Ryckaert,J.P., Cicotti,G. and Berendsen,H.J.C. (1977) J. Comp. Phys., 23, 327341.[ISI]
Schanstra,J.P. and Janssen,D.B. (1996) Biochemistry, 35, 56245632.[CrossRef][ISI][Medline]
Schanstra,J.P., Rink,R., Pries,F. and Janssen,D.B. (1993) Protein Expr. Purif., 4, 479489.[CrossRef][ISI][Medline]
Schanstra,J.P., Ridder,I.S., Heimeriks,G.J., Rink,R., Poelarends,G.J., Kalk,K.H., Dijkstra,B.W. and Janssen,D.B. (1996) Biochemistry, 35, 1318613195.[CrossRef][ISI][Medline]
Schanstra,J.P., Ridder,A., Kingma,J. and Janssen,D.B. (1997) Protein Eng., 10, 5361.[Abstract]
Shinkai,A., Hirano,A. and Aisaka,K. (1996) J. Biochem. (Tokyo), 120, 915921.[Abstract]
Sowdhamini,R., Srinivasan,N., Shoichet,B., Santi,D.V., Ramakrishnan,C. and Balaram,P. (1989) Protein Eng., 3, 95103.[Abstract]
Studier,F.W., Rosenberg,A.H., Dunn,J.J. and Dubendorff,J.W. (1990) Methods Enzymol., 185, 6089.[Medline]
Takagi,H., Takahashi,T., Momose,H., Inouye,M., Maeda,Y., Matsuzawa,H. and Ohta,T. (1990) J. Biol. Chem., 265, 68746878.
Thornton,J.M. (1981) J. Mol. Biol., 151, 261287.[ISI][Medline]
Tidor,B. and Karplus,M. (1993) Proteins, 15, 7179.[ISI][Medline]
van Buuren,A.R., Marrink,S.J. and Berendsen H.J.C. (1993) J. Phys. Chem., 97, 92069212.[ISI]
van der Spoel,D., Berendsen,H.J.C., Apol,E., Meulenhoff,P.J., Sijbers,A.L.T.M. and van Drunen,R. (1996a) Gromacs User Manual. Laboratory of Physical Chemistry, University of Groningen, The Netherlands. Internet: http://rugmd0.chem.rug.nl/~gmx.
van der Spoel,D., van Buuren,A.R., Tieleman,D.P. and Berendsen,H.J.C. (1996b) J. Biomol. NMR, 8, 229238.[ISI][Medline]
van Gunsteren,W.F and Berendsen,H.J.C. (1987) Gromos Manual. BIOMOS, Biomolecular Software. Laboratory of Physical Chemistry, University of Groningen, The Netherlands.
Verschueren,K.H., Franken,S.M., Rozeboom,H.J., Kalk,K.H. and Dijkstra,B.W. (1993a) J. Mol. Biol., 232, 856872.[CrossRef][ISI][Medline]
Verschueren,K.H., Seljee,F., Rozeboom,H.J., Kalk,K.H. and Dijkstra,B.W. (1993b) Nature, 363, 693698.[CrossRef][ISI][Medline]
Villafranca,J.E., Howell,E.E., Voet,D.H., Strobel,M.S., Ogden,R.C., Abelson,J.N. and Kraut,J. (1983) Science, 222, 782788.[ISI][Medline]
Vogl,T., Brengelmann,R., Hinz,H.J., Scharf,M., Lotzbeyer,M. and Engels,J.W. (1995) J. Mol. Biol., 254, 481496.[CrossRef][ISI][Medline]
Weber,P.C., Sheriff,S., Ohlendorf,D.H., Finzel,B.C. and Salemme,F.R. (1985) Proc. Natl Acad. Sci. USA, 82, 84738477.[Abstract]
Yamaguchi,S., Takeuchi,K., Mase,T., Oikawa,K., McMullen,T., Derewenda,U., McElhaney,R.N., Kay,C.M. and Derewenda,Z.S. (1996) Protein Eng., 9, 789795.[Abstract]
Ziegler,D.M. and Poulsen L.L. (1977) TIBS, 2, 7981.
Received May 15, 2001; revised November 23, 2001; accepted December 7, 2001.