Báo cáo khoa học: Role of conformational flexibility for enzymatic activity in NADH oxidase from Thermus thermophilus pptx

Role of conformational flexibility for enzymatic activity in NADH oxidase from Thermus thermophilus Gabriel Z ˇ olda ´ k 1 ,Ro ´ bert S ˇ ut’a ´ k 1,2 , Maria ´ n Antalı ´ k 1,3 , Mathias Sprinzl 4 and Erik Sedla ´ k 1 1 Department of Biochemistry, Faculty of Sciences P. J. S ˇ afa ´ rik University, Kos ˇ ice, Slovakia; 2 Department of Parasitology, Charles University, Prague, Czech Republic; 3 Department of Biophysics, Institute of Experimental Physics, Slovak Academy of Sciences, Kos ˇ ice, Slovakia; 4 Laboratorium fu ¨ r Biochemie, Universita ¨ t Bayreuth, Germany NADH oxidase from Thermus thermophilus is a homodimer with an unknown physiological function. As is typical for an enzyme isolated from a thermophile, the catalytic rate, k cat , is low at low temperatures and increases with temperature, achieving an optimum at the physiological temperature of the organism, i.e. at % 70 °CforT. thermophilus.Atlow temperatures, the k cat of several enzymes from thermophilic and mesophilic organisms can be increased by chaotropic agents. The catalytic rate of NADH oxidase increases in the presence of urea. At concentrations of 1.0–1.3 M urea it reaches 250% of the activity in the absence of urea, at 20 °C. At higher urea concentrations the enzyme activity is inhi- bited. The urea-dependent activity changes correlate with changes in the fluorescence intensity of Trp47, which is located in the active site of the enzyme. Both fluorescence and circular dichroism measurements indicate that the acti- vation by chaotropic agents involves local environmental changes accompanied by increased dynamics in the active site of the enzyme. This is not related to the global structure of NADH oxidase. The presence of an aromatic amino acid interacting with the flavin cofactor is common to numerous flavin-dependent oxidases. A comparison of the crystal structure with the activation thermodynamic parameters, DH*andTDS*, obtained from the temperature dependence of k cat, suggests that Trp47 interacts with a water molecule and the isoalloxazine flavin ring. The present investigation suggests a model that explains the role of the homodimeric structure of NADH oxidase. Keywords: NADH oxidase; conformational dynamics; flavo- proteins; fluorescence quenching; Thermus thermophilus. The activity and stability of an enzyme is a compromise between two opposing forces in the dynamics of the polypeptide chain. While the active site of an enzyme has to have a certain flexibility to fit the incoming substrate, the stability is related to the rigidity of the polypeptide chain [1–3]. The balance between the stability/rigidity and the flexibility of the protein structure is achieved in the native structure at physiological temperatures [4,5]. It was suggested nearly 50-years ago [6,7] that conformational flexibility in the active site is important for substrate binding, and for enzyme catalysis. The highly dynamic active site is more highly sensitive to perturbations of the environment than the rest of the polypeptide structure, which agrees with the observation that enzyme inactivation precedes global unfolding of the enzyme structure [8]. The extreme stability of enzymes from thermophilic organisms is an attractive feature for biotechnological applications [9]. On the other hand, these enzymes have low activity at temperatures below their physiological temperature. Find- ing conditions in which an enzyme is activated but not destabilized at low temperature is one way to increase the catalytic efficiency of the thermophilic enzymes. Another way would be to identify the rate-limiting step in enzyme catalysis. This information may indicate a suitable amino-acid residue in the active site as a target for protein engineering that could result in activation of the enzyme [2]. Here, we report the case of a thermophilic enzyme that is sensitive to the conformational flexibility of the active site. We have studied the effect of urea on NADH oxidase (EC 1.6.99.3) from Thermus thermophilus.NADHoxidase is a dimeric flavoprotein containing one molecule of FMN in each 25-kDa monomer, and it catalyzes hydride transfer from NADH to an acceptor such as FAD, ferricyanide, oxygen, and others [10]. It belongs to the flavin reductase/nitroreductase family that has similar broad substrate specificity, similar folding and similar quaternary structure [11,12]. The localization and potential physiological role of this ÔalternativeÕ dehydrogenase in this thermophile species is not known. In the course of the purification procedure, the major activity of NADH oxidase was found in the supernatant of the cell lysates. The main location of the NADH oxidase activity was found in the polar aqueous solution. This indicates a possible role in regulation of the cytoplasmic NADH/ NAD + moiety. The flavin cofactors, FMN and FAD, are tightly bound with dissociation constants of % 10 )7 M )1 and % 10 )5 M )1 , respectively. The low temperature factor determined from the crystal structure also indicates tight binding [10]. NADH oxidase is relatively rigid, however, the cofactor is located in Correspondence to E. Sedla ´ k, Department of Biochemistry, Faculty of Sciences, P. J. S ˇ afa ´ rik University, Moyzesova 11, 041 54 Kos ˇ ice, Slovakia. E-mail: sedlak_er@saske.sk Enzymes: NADH oxidase (EC 1.6.99.3). (Received 20 September 2003, accepted 22 October 2003) Eur. J. Biochem. 270, 4887–4897 (2003) Ó FEBS 2003 doi:10.1046/j.1432-1033.2003.03889.x the intermonomeric interface that is a region with relatively high dynamics [13]. Binding of substrate in homologous reductases is accompanied by the induced fit of the helix at the intermonomeric interface. The high temperature factor of the analogous helix in NADH oxidase indicates its high flexibility. Destabilization of this helix containing Trp47 would affect the interaction of the flavin cofactor with Trp47 in the active site of the enzyme. This could be a general mechanism of substrate/enzyme interaction in this flavoprotein family. In the work presented here, we have shown that NADH oxidase can be activated in low concentrations of chaotropic agents due to increased local dynamics in the active site. The rate-limiting step in NADH oxidase is proposed to include movement of Trp47. The observed correlation between activity and tryptophan fluorescence can occur only when the enzyme is in a dimeric form, indicating that NADH oxidase is a functional homodimer. Experimental procedures Analytical-grade biochemicals were obtained from Merck (Germany). Urea (high purity grade) was purchased from Sigma. Urea concentrations were determined from refract- ive index measurements using an Abbe Refractometer AR3- AR6. The pH values of the solutions were measured with a Sensorex glass electrode before and after measurement at room temperature. Only the measurements at which the pH change was less than 0.2 pH units were taken for further consideration. Protein expression and purification The NADH oxidase from T. thermophilus was overpro- duced in E. coli JM 108 using recombinant plasmid pTNADOX (amp R , tac promotor and nox gene) [14]. 1m M IPTG (Gerbu) was added after the bacterial culture reached D 600 ¼ 0.9–1.0 and harvested after 4–5 h. The purification procedure for the overproduced NADH oxi- dase was described earlier [15] and used with only minor modifications. The heat treatment step was performed in the presence of a small amount of FAD (increases the thermal stability of the enzyme), dialyzed and loaded on a Blue Sepharose CL-6B affinity column. After the washing procedure NADH oxidase was eluted with 1 m M NAD + . The final product was a single band on a SDS/PAGE gel [16] stained with Coomassie Brilliant Blue. Before use, the protein was dialyzed in the absence of FAD in 50 m M phosphate buffer, pH 7.2. The final preparation yielded NADH oxidase with a specific activity of 11.32 unitsÆmg )1 at 20 °C. One unit is defined as 1 l M NADH oxidized per min. Determination of the protein concentration The extinction coefficient (e) of the protein at 280 nm was calculated from the number of tryptophan residues (4), tyrosine residues (7) and cysteine residues (0) per mono- mer using an equation in [17]. The predicted molar absorption coefficient for apoenzyme is e 280 ¼ 32 430 M )1 Æcm )1 . The noncovalently bound cofactor FAD also contributes to the extinction coefficient at 280 nm. The molar absorption coefficient for FAD dissolved in pH 7.2 phosphate buffer, is e 280 ¼ 20 600 M )1 Æcm )1 . Therefore, the protein concentration with the bound cofactor was determined using the extinction coefficient e 280 ¼ 52 030 M )1 Æcm )1 . The calcu- lated specific activity is very similar to previous data [15], provided the protein concentration was determined according to the method of Bradford. Steady-state kinetics All kinetic measurements were performed on a Shimadzu UV3000 spectrophotometer. The kinetic parameters were determined from the initial decrease in the absorbance of NADH at 340 nm (e 340 ¼ 6220 M )1 Æcm )1 ), at 20 °C. Measurements were performed after incubation (12 h) in 120 n M NADH oxidase holoenzyme, 50 m M sodium phosphate, pH 7.2, containing 0.120 m M FAD and differ- ent concentrations of urea. The reaction was started with the addition of NADH. The observed rate at 340 nm is a combination of the enzyme-mediated rate changes and other rates, e.g. the self-decay of NADH and the reduction of externally added FAD. The self-decay of NADH is insignificant in these conditions and needs to be taken into account only at high temperatures. The externally added FAD has an absorption maximum at 375 nm, and reduction of the flavin might affect the absorbance at 340 nm. To determine if the change in the redox state of exogenously added FAD contributes to the time-dependent changes in absorbance at 340 nm, related to oxidation of NADH, we have monitored the reduction/oxidation reac- tion of FAD. Because it is very complicated to follow this reaction in the presence of NADH at 340 nm we have monitored the reduction/oxidation of FAD at 450 nm. Our results indicated that equilibrium of the reaction has been achieved within the time (% 10 s) the instrument took to start collecting data, which is in accordance with a previously reported observation [18]. Therefore, this reac- tion does not contribute to time-dependent changes in absorbance at 340 nm during measurements. The oxidation rate of NADH depends on the initial flavin concentration, and saturation occurred at nearly 0.10 m M flavin. In the enzyme assay the concentration of FAD was always 0.120 m M . The data were fitted to the Michaelis–Menten equation where K M, app corresponds to the apparent Michaelis constant and the apparent V max is the maximum velocity for the catalytic reaction. The experimental data were also plotted according Lineweaver-Burk and analyzed by linear regression. Similar results were obtained using both methods. Temperature dependence of enzyme activity Enzyme activity measurements were performed in 50 m M phosphate buffer, 0.120 m M FAD and 120 n M holoenzyme. The reactions were started by the addition of NADH to achieve a final concentration of 0.180 m M NADH. The initial velocities were measured from 20 to 40 °C. The temperature during measurements was kept constant by temperature controlled water circulation around the cuvette. Temperature dependences were analyzed with a simple Arrhenius equation 4888 G. Z ˇ olda ´ k et al.(Eur. J. Biochem. 270) Ó FEBS 2003 lnk cat ¼À E a RT þ C 1 ð1Þ where R is the gas constant (8.314 JÆK )1 Æmol )1 ), E a is the activation energy for the observed reaction and C 1 is a temperature independent constant. Data (at least five points) were plotted as ln(k cat )vs.T )1 and analyzed by linear regression. Coefficients of linearity were typically higher than 0.98. From comparison of the Arrhenius equation and the transition state theory the enthalpy DH* and entropy DS* of activation were calculated DH à ¼ E a À RT ð2Þ T ln k cat T  ¼ TDS à R þ C 2 ð3Þ C 2 is the temperature independent constant. This approach avoided any extrapolation connected with large errors in the estimation of the activation entropy [19]. ThefreeenergyofactivationDG* was calculated from the equation: DG à ¼ DH à À TDS à ð4Þ Fluorescence emission spectroscopy The fluorescence steady-state measurements were per- formed on a Shimadzu RF5000 spectrofluorophotometer. Using different excitation wavelengths, i.e. 280, 290 and 450 nm, we were able to follow changes in the environment close to different internal chromophores, i.e. Tyr, Trp and FAD, respectively. The cuvette contained 50 m M sodium phosphate, pH 7.2, with various concentrations of urea and 2.4 l M dimeric protein in a total volume of 2.5 mL. To avoid the inner filter effect the absorbance of protein samples was always lower than 0.1. Samples were incubated 12 h at room temperature. The data from all fluorescence and quenching experiments were collected at 25 °C. The quantum yields were calculated by a comparative method using the integrated areas of fluorescence intensity for protein samples and for free L -tryptophan [20,21]. The quantum yield of free L -tryptophan was used as a standard (F L-Trp ¼ 0.14) [22]. A similar approach was also used for FAD in solution (F FAD ¼ 0.05). Fluorescence quenching Quenching experiments were performed with acrylamide (Carl Roth GmbH & Co., Germany). A fresh 2 M acryl- amide (14.2%) solution was dissolved in 50 m M sodium phosphate buffer, pH 7.2. Protein concentrations of 5–10 l M were used in 50 m M sodium phosphate buffer, pH 7.2, and various concentrations of urea in a total volume of 2.5 mL. The acrylamide was added to the cuvette in 5, 10 and 20 lL aliquots. After 30 s incubation the emission spectra after excitation at 290 nm were recorded. Longer incubation times were not necessary. No significant changes occurred in the emission band even after 1 h of incubation. Therefore, a 30 s incubation interval was used for all measurements and samples were assumed to reach equilibrium. Analysis of the experimental data was performed using several models. The Stern–Volmer equation (Eqn 5) assumes a homogenous population of fluorophores: F 0 F ¼ 1 þ k 0 s 0 ½Qð5Þ where k 0 s 0 ¼ K SV which is the quenching constant. k 0 is the bimolecular quenching constant describing collisional quenching, and s 0 is the fluorescence lifetime of the tryptophan residues. In some cases, quenching of the tryptophan moiety could be described with a model of a single fluorophore population [23]. This model was success- fully used for N-acetyl- L -tryptophanamide and also for NADH oxidase in 9 M urea. Equation 5 does not include static quenching, i.e. the formation of a fluorophore complex with the quencher before excitation. In the case of static quenching, the dependence of F 0 /F on Q, as plotted, has an upward curvature due to factor e [Q]V where V is the static constant [24]. Data obtained from the quenching of NADH oxidase by acrylamide were impossible to fit to a simple Stern–Volmer equation due to a downward curva- ture of F 0 /F vs. Q. This is typical for heterogeneous populations of fluorophores. This is not surprising because NADH oxidase contains four tryptophan residues, each with a different extent of accessibility to the quencher. Quenching of the tryptophan moieties of NADH oxidase could be described in terms of accessible and nonaccessible populations using a modified Stern–Volmer equation [25]: F 0 F 0 À F ¼ 1 f a þ 1 f a Á K c ½Q ð6Þ where f a is the fraction of accessible fluorophore and K c is the effective collisional quenching constant. This modified equation assumes that the population is heterogeneous and that there is a difference in the quenching behavior of the different tryptophan moieties. A linear regression, F 0 F 0 ÀF vs 1 ½Q whose slope ¼ 1 f a K c and intercept ¼ 1 f a was used for data analysis. Data processing was performed using GRAFIT 3.00 (Erithacus Software Ltd, Cambridge, UK). Circular dichroism measurements CD measurements were performed on a Jasco J-600 (Tokyo, Japan) spectropolarimeter at 20 °Cwith29.3l M NADH oxidase in 10 m M sodium phosphate, pH 7.2, and urea. A 0.1 cm path-length cuvette was used for the peptide region and a 1 cm cuvette for the aromatic region. Each spectrum was an accumulation of 4–6 consecutive scans. The thermal transitions were recorded at 222 nm with a constant scan rate of 1 KÆmin )1 . The temperature was measured with a PTC)348 WI Peltier block inside the cuvette. The temperature calibration was performed with a Brand (Wertheim, Germany) precision thermometer. Results Enzyme activity The catalytic mechanism of NADH oxidase is not under- stood. The enzyme kinetics of NADH oxidase from T. thermophilus were analyzed using a simple Michaelis– Menten model where FAD is at a saturation level. Figure 1 Ó FEBS 2003 Flexibility of the NADH oxidase active site (Eur. J. Biochem. 270) 4889 shows the time-dependence of NADH oxidation monitored at 340 nm in the absence and in the presence of 1.25 M and 4 M urea. Surprisingly, the activity of NADH oxidase is increased in the presence of urea and reached its optimum at 1.25 M urea. The Lineweaver–Burk plot (Fig. 1, inset) indicates that the presence of low urea concentrations affects both the apparent maximal velocity of the reaction and the apparent Michaelis constant for NADH’s interaction with the enzyme. In the absence of urea the apparent steady-state parameters were k cat ¼ 6.6 ± 0.1 s )1 and K M,app ¼ 5.2 ± 0.2 l M , and the catalytic efficiency was k cat /K M,app ¼ 1.3 ± 0.1 · 10 6 M )1 Æs )1 . These values are similar to those published previously [15]. All parameters consist of multiple kinetic terms and could not be associated directly with any one step in the catalytic reaction. The effect of urea was studied in detail, and the results are shown in Fig. 2. The measured parameters are summarized in Table 1. The velocity of NADH oxidation is 2.5 fold higher at 1.0–1.3 M urea compared to the control. The catalytic rate of NADH oxidase is also increased in the presence of ionic chaotropic reagents such as guanidine hydrochloride (% 0.5 M ) and sodium perchlorate (% 0.25 M ) (data not shown). In all experiments, externally added flavin was the electron acceptor that recovered the reduced internal flavin. However, a similar activation of NADH oxidase was also observed in the presence of alternative acceptors such as ferricyanide (data not shown). The presence of urea has a similar effect on both k cat and K M , i.e. the increase in k cat is associated with an increase in K M . This results in nearly constant values of k cat /K M at different urea concentrations (Table 1). At higher concen- trations of urea (> 2 M ) k cat sequentially decreases and, at 6.0 M urea, the enzyme is essentially inactive. Fluorescence NADH oxidase from T. thermophilus contains many fluoro- phore groups: seven tyrosine residues, four tryptophan residues and the flavin cofactor per monomer. The trypto- phan residues emission spectra were followed after excitation at 290 nm. The maximum of the emission spectrum was 336 nm, i.e. the maximum shifted to lower wavelengths compared to the emission spectrum of solvent exposed L - tryptophan (352 nm) (Fig. 2, inset). This indicates that tryptophan residues in the NADH oxidase dimer are buried in nonpolar regions of the protein [26]. The emission band is a convoluted contribution of all tryptophan residues in the enzyme; therefore, it is difficult to determine separate quantum yields. The averaged quantum yield is low (F av ¼ 0.07). The quantum yield of solvent accessible L - tryptophan is 0.14 and it increases if the tryptophan residues are buried. The low quantum yield of tryptophan fluorescence in NADH oxidase shows efficient quenching of the trypto- phan residues in the protein. Such quenching can be the result of interactions with the flavin cofactor, the imidazole ring of histidine residues, negatively charged carboxylic groups and/ or by the highly mobile indole group of the tryptophan residues [27]. Steady-state analysis of the FAD fluorescence in NADH oxidase has shown that its emission maximum after excitation at 450 nm is centered at 522 nm. This is very similar to the value of the emission maximum characteristic for free FAD in aqueous solution (emission at 525 nm). This finding is in agreement with the location of the cofactor in the crystal structure of NADH oxidase [10]. The quantum yield of the flavin cofactor in NADH oxidase (F ¼ 0.02) is smaller than that of free FAD in solution (F ¼ 0.05). The position of the tryptophans and the flavin cofactor in the crystal structure of NADH oxidase is depicted in Fig. 3. It should be noted that the structure shown contains FMN as the cofactor. However, the exchange of FMN for FAD results in essentially an identical structure with only Fig. 1. Enzymatic oxidation of NADH by NADH oxidase from T. thermophilus monitored by absorbance at 340 nm at 0 M , 1.25 M , and 4 M urea. Changes in absorbance were normalized. The curve is not based on a theoretical analysis, it serves only to lead eyes. Inset: Line- weaver–Burk plot for NADH oxidation in the absence of urea (s)and in the presence of 1.25 M urea (d). Assays were performed at 20 °C. Fig. 2. The effect of urea concentration on the activity (d) and intrinsic fluorescence (n)ofNADHoxidasefromT. thermophilus. Values of fluorescence intensities are shown as the ratio F/F 0 , where F 0 corres- ponds to fluorescence at 0 M urea, and similarly A/A 0 is the ratio of the activity (A) in the presence of urea and A 0 corresponds to the enzyme activity at 0 M urea. Inset: Fluorescence emission spectra of NADH oxidase in the absence (solid line) and in the presence of 1.0 M urea (dashed line). Decrease in the fluorescence and the slight red-shift of the fluorescence maximum was observed at the low urea concentra- tion. Activity was determined from the initial linear decrease of the absorbance at 340 nm. The fluorescence measurements were per- formed with 5 l M protein using an excitation wavelength of 290 nm for tryptophan residues. All experiments were performed at 20 °C. 4890 G. Z ˇ olda ´ k et al.(Eur. J. Biochem. 270) Ó FEBS 2003 slight conformational changes of the C-terminal end between Glu189 and His194 to accommodate the second phosphate group of FAD [10]. Tryptophan residues in the monomeric form of NADH oxidase are spatially separated from the location of the flavin cofactor. The distance between N1 of the flavin cofactor and N e1 of tryptophans 47, 52, 131, 204 are 33.6 A ˚ , 22.7 A ˚ , 25.1 A ˚ , 29.5 A ˚ , respectively. In the dimeric form, the distances of trypto- phan residues 47, 52, 131, 204 from the flavin cofactor are 7.7 A ˚ , 16.3 A ˚ , 12.5 A ˚ , 25.3 A ˚ , respectively. Interestingly, changes in the enzyme activity correlate with changes in the fluorescence intensity of the tryptophan residues. Fluores- cence probes the properties of the local environment of the dipole–dipole interaction rather than global structural changes in proteins. As the dipole–dipole interaction decreases very steeply with distance (as 1/distance 6 ), the relative position of Trp47 and the flavin cofactor is especially notable. Moreover, the crystal structure indicates that contact between Trp47 and the cofactor is mediated through a tightly bound structural water molecule [10]. This strongly indicates that the fluorescence of Trp47 is respon- sible for the observed correlation between activity and tryptophan fluorescence (Fig. 2). It is not possible to exclude allosteric effects that could affect the distance between the cofactor and the other tryptophan residues. This is probably not the case because we could not see significant changes in the circular dichroism spectra that would accompany such a significant conformational change (see below). In the presence of 1.0–1.3M urea, there is nearly a 60% decrease in tryptophan fluorescence simultaneous with a slight red shift (% 5 nm) of the tryptophan emission maximum (Fig. 2, inset). Tryptophan residues 131 and 204 are completely exposed to solvent while Trp52 is rigidly embedded at a distant location in the protein matrix. In the case of NADH oxidase the perturbation of a microenvi- ronment, probably that of Trp47, is interrelated with the changes in activity at a narrow concentration range of urea. At higher concentrations of urea (> 7 M ) the fluorescence intensity sharply increases due to unfolding of the protein and dissociation of the flavin cofactor (data not shown). At 9 M urea the tryptophan residues of NADH oxidase possess characteristics very similar to free L -tryptophan F % 0.19 and k em ¼ 350 nm. The flavin fluorescence is not changed significantly in the presence of 0–7 M urea (data not shown). The urea-induced changes in enzyme activity and urea- induced protein unfolding, as monitored by fluorescence, show that inactivation of the enzyme takes place before the global unfolding of the protein. Circular dichroism The global structure of the protein may be efficiently monitored by CD spectroscopy. The effect of urea on the Table 1. Steady-state kinetic parameters (k cat and apparent K M ) at various concentrations of urea and temperature. Acrylamide quenching constants and the fraction of the accessible tryptophans for NADH oxidase at various concentrations of urea. Activity and quenching experiments were performed at 20 °C (see Experimental procedures). Kinetic parameters were obtained by the nonlinear regression analyses of a simple Michaelis– Menten equation. Quenching parameters were obtained from fitting by a modified Stern–Volmer equation (Eqn 6). Standard deviations (±) represent possible errors in the estimated parameters for straight line. Urea ( M ) 0 0.5 1.0 1.5 2.0 Activity K M, app (l M ) 5.2 ± 0.2 8.8 ± 0.2 9.2 ± 0.4 13.8 ± 0.9 9.1 ± 0.4 k cat (s )1 ) 6.6 ± 0.1 9.9 ± 0.1 14.9 ± 0.2 15.3 ± 0.4 9.9 ± 0.2 k cat /K M, app ( M )1 Æs )1 ) 1.27 · 10 6 1.12 · 10 6 1.64 · 10 6 1.10 · 10 6 1.10 · 10 6 T opt (°C) 70 ± 3 47 ± 3 50 ± 3 53 ± 3 52 ± 3 Fluorescence quenching K c ( M )1 ) 20 ± 1 7.6 ± 0.6 6.8 ± 0.4 10.0 ± 0.1 14.9 ± 0.1 f a 0.42 ± 0.03 0.59 ± 0.17 0.71 ± 0.08 0.61 ± 0.03 0.48 ± 0.03 r a (0.9900) (0.9720) (0.9900) (0.9968) (0.9891) a Coefficients obtained by linear regression. Fig. 3. Dimeric structure of NADH oxidase from T. thermophilus. Monomers are drawn in different greyscale. All tryptophan residues and FMN cofactors are shown. Noteworthy, Trp47 is located close to the environment of the FMN cofactor. The structure was drawn using VIEWER LITE 42 (1NOX.pdb). Ó FEBS 2003 Flexibility of the NADH oxidase active site (Eur. J. Biochem. 270) 4891 activity of NADH oxidase was therefore investigated by circular dichroism at various concentrations of urea (Fig. 4). The shape of the spectra in the far UV region is typical for a mixture of a-helix and b-sheet elements in the secondary structure. No apparent differences were observed in ellipticity in the peptide region in the absence or in the presence of 6.7 M urea. The secondary structure of the enzyme is unaffected even at high concentrations of urea indicating an extreme resistance to urea-induced perturbations. The ellipticity in the near UV region is characteristic for aromatic residues – tryptophan, tyrosine, and the flavin cofactor. Contrary to the situation in the peptide region, the aromatic region is sensitive to urea (Fig. 4B). The proximity of Trp47 to the flavin cofactor induces asymmetry in the tryptophan environment that is likely to result in a strong positive signal in the aromatic region. The addition of urea causes gradual changes in the near UV spectrum (Fig. 4B, inset) accompanied by a decrease in ellipticity at 265 nm and a slight shift to longer wavelengths. The isodichroic point at about 270 nm indicates that the conformational transition has a two- state character. As observed by fluorescence measurements, the circular dichroism results confirm that the inactivation of the enzyme at high concentrations of urea (> 6 M )is not accompanied by the global unfolding reaction. The gradual changes in ellipticity in the aromatic region indicate local conformational changes and/or changes in the tertiary structural dynamics in the environment of the flavin cofactor. Thermal stability of the active site and global structure of the enzyme The temperature dependence of the enzyme activity was measured as an indicator of stability of the active site. Unfolding of secondary structure is related to global unfolding of protein structure. The ellipticity at 220 nm was therefore measured to assess the stability of the global structure (Fig. 5). In the absence of urea, the enzyme achieves its maximal activity at % 70 °C which is close to the optimal temperature of T. thermophilus [15]. In the condi- tions where the maximal activity of the enzyme at room temperature was achieved, i.e. in the presence of 1.25 M urea, the stability of the active site of the enzyme is significantly perturbed. The optimal temperature for enzyme activity at 1.25 M urea was shifted by % )25 °C from the optimal temperature of NADH oxidase in the absence of urea (Fig. 5). An additional increase in the urea concentration (> 2.0 M ) had no significant effect on the optimal temperature of the enzyme but reduced the maximal enzyme activity (Fig. 5, Table 1). The transition temperature, T trs , of unfolding of the secondary structure, is represented by the position of the peak maximum of the first derivative of ellipticity at 220 nm dQ/dT. Global stability, characterized by this transition temperature, is significantly higher than the thermal stability of the active site of the enzyme (Fig. 5). In the absence of urea, T trs ¼ 88.6 °C , about 15 °C higher than the temperature of the physiolo- gical milieu of T. thermophilus. In the presence of low concentrations of urea, i.e. 2.5 M , the transition temperature decreases only by about 3 °C. Even in the conditions where the enzyme is completely inactive, i.e. at 6.7 M urea, the thermal transition of the protein secondary structure has a sigmoidal shape with T trs ¼ 71.6 °C (data not shown). In summary, the active site of the enzyme is considerably more sensitive to temperature-induced perturbation than the global structure of NADH oxidase. This is most pro- nounced at low concentrations of denaturant where the optimal activity of the enzyme is achieved at the expense of the flexibility/stability of the active site of the enzyme. Fig. 4. Circular dichroism spectra of NADH oxidase from T. thermo- philus in the peptide (A) and aromatic (B) regions in the absence and presence of urea. (A) 0 M urea (solid line), 6.7 M urea (dashed-double dotted line). (B) 0 M urea (solid line), 0.9 M (dashed line), 1.7 M (dotted line), 3.5 M (dash-dotted line), and 6.7 M urea (dash-double dotted line). Measurements were performed on a Jasco J-600 spectropola- rimeter with 29.3 l M NADH oxidase. A 0.1 cm path-length cuvette was used for far UV and a 1 cm cuvette for the aromatic region. Inset: dependence of difference in the CD spectra at 250–320 nm on con- centration of urea. The curve in the inset is not based on a theoretical analysis, it serves only to lead the eyes. Fig. 5. Normalized activity and the first derivative of the melting curve of NADH oxidase monitored by ellipticity at 220 nm. Activity was measured in 50 m M phosphate buffer, pH 7.2, in the absence of urea (d), 1.25 M urea (n)and2.0 M urea (h), respectively. Heat denatur- ation was measured in 10 m M phosphate buffer, pH 7.2, in 0 M urea (solid line), 1.3 M urea (dotted line) and at 2.6 M urea (dashed line). The lines are not based on a theoretical curve but only serve to lead the eyes. Transition temperature at 0 M urea T trs ¼ 88.6 °C, 1.3 M urea T trs ¼ 88.4 °C and at 2.6 M urea T trs ¼ 85.6 °C. 4892 G. Z ˇ olda ´ k et al.(Eur. J. Biochem. 270) Ó FEBS 2003 To address quantitatively the activation of NADH oxidase at low concentrations of denaturant, the activation parameters DH*andDS* of the reaction were determined (Fig. 6). The activation parameters were calculated accord- ing equations 1–3. The urea effect on DH*andTDS* indicates that both parameters reach a minimum at the urea concentration at which enhancement of activity is observed (Fig. 2). The Arrhenius plot in the absence of urea is linear from 20 to 65 °C without any apparent curvature or break (Fig. 6, inset) as was reported for some other enzymes from thermophiles [28]. Interestingly, the Arrhenius plots in the absence and in the presence of 1.0 M urea intersect at % 60 °C. This indicates that the activity of NADH oxidase is the same in both conditions at this temperature. Consequently, the enzyme was not activated by the addition of urea at 60 °C (data not shown). Fluorescence quenching In an effort to monitor the accessibility of the tryptophan residues as an indicator of the dynamics of the active site, quenching experiments were performed. Acrylamide was used to quench the fluorescence of the tryptophan residues. It is known to be an effective quencher of the fluorescence of tryptophan residues that are completely or partially exposed to solvent [29]. Quenching experiments enabled us to determine the effective quenching constants K c , and the number of accessible tryptophan residues f a .NADH oxidase contains four tryptophan residues in different locations within the enzyme that prevents a detailed analysis. A comparison of the parameters determined by quenching, however, enabled us to extract information about changes in the accessibility of the tryptophan residues in NADH oxidase (Table 1). Quenching of N-acetyltrypto- phanamide, as a model compound that is fully accessible to the solvent, is characterized by K c ¼ 35.7 ± 0.3 M )1 and f a ¼ 1.02 ± 0.01 (Eqn 6). If the simple Stern–Volmer model is used we obtained K SV ¼ 37 ± 2 M )1 (Eqn 5). In the absence of urea, the effective quenching constant for NADH oxidase was K c ¼ 20 ± 1 M )1 and the fraction of accessible fluorophore f a ¼ 0.42 ± 0.03, indicating that % 2 tryptophan residues are accessible to solvent (Fig. 7) which agrees with the crystal structure that shows that Trp131 and Trp204 are completely exposed to the solvent [10]. Iodide anions are another type of quencher that, due to the negative charges, is only accessible to protein surfaces. The fraction of fluorophores accessible to iodide anions was f a ¼ 0.41 ± 0.02. The activity and the conformation of the active site of NADH oxidase are very sensitive to both the ionic strength and the type of anions (G. Z ˇ olda ´ k, M. Sprinzl and E. Sedla ´ k, unpublished results). Therefore, for further quenching experiments we have used only uncharged acrylamide. Table 1 shows that the fraction of exposed tryptophan residues of NADH oxidase increased in 1.0 M urea to the value f a ¼ 0.71 ± 0.08. This value corresponds to the exposure of three tryptophan residues to solvent. To be certain that acrylamide does not affect the conformation of the enzyme active site, we have determined the enzyme activity in the presence of up to 180 m M acrylamide. At such relatively high concentrations of acrylamide the enzyme activity was only slightly diminished, % 10–15%, compared to the sample without acrylamide. This indicates that acrylamide does not significantly affect the conformation of the enzyme active site. This agrees with previously published results that acrylamide does not seriously perturb the native Fig. 6. Activation parameters DH*(d) and TDS*(s) as a function of urea concentration. Values were obtained from kinetic experiments described in Experimental procedures using Eqns 1–3. Arrhenius plots were analyzed at various concentrations of urea and temperatures in the range from 20 to 40 °C. Errors were calculated according to the deviation from linearity of the Arrhenius plots. The lines are not based on a theoretical curve, but only serve to lead the eyes. Inset: Arrhenius plots of NADH oxidase in the absence and in the presence of 1.0 M urea, respectively. The plots intersect at % 60 °C, i.e. the activity is equal at this temperature, indicating that the dynamics of the active site are comparable. Fig. 7. Modified Stern–Volmer plots (Eqn 6) for acrylamide quenching of tryptophan fluorescence at 0 M urea (d)and1.0 M urea (s). Lines were obtained from the linear regression analysis. F 0 , F is fluorescence in the absence and presence of acrylamide, DF represents the difference F 0 –F.For0 M urea: y ¼ 0.121Æx +2.41(r ¼ 0.990). For 1.0 M urea: y ¼ 0.210Æx +1.40 (r ¼ 0.990). For comparison, the acrylamide quenching of N-acetyl- L -tryptophanamide (NATA) fluorescence is shown (dashed line). NATA is a tryptophan analogue completely exposed to molecules of solvent and the quencher. Ó FEBS 2003 Flexibility of the NADH oxidase active site (Eur. J. Biochem. 270) 4893 conformation of proteins because the enzyme activity of a number of proteins is unaffected in the presence of acrylamide [29]. Discussion Activation of NADH oxidase is caused by an increase in the conformational dynamics of the enzyme active site NADH oxidase from T. thermophilus, has diminished activity at low temperatures, similar to many enzymes from thermophilic organisms [4,30]. The activation of NADH oxidase at low concentrations of chaotropic agents may result from: (a) a conformational change and/or (b) increased dynamics in the enzyme active site or (c) destabilization of the enzyme-product complex. Several types of electron-acceptors used (FMN, FAD, oxygen, ferricyanide) gave similar results. This indicates that desta- bilization of the enzyme-product complex is not the rate- limiting step in NADH oxidase catalysis. It also indirectly indicates that the kinetic mechanism of NADH oxidase is the ping-pong reaction found in other homologue oxidases [31]. The activation is due to conformational and/or dynamic changes in the enzyme active site. This agrees with previously published observations that activation of differ- ent enzymes at low concentrations of chaotropic agents was associated with conformational changes in the tertiary [32–37], and secondary [38] structure of the enzymes, or the dynamics of the enzyme active site [39,40]. NADH oxidase activation correlates with changes in tryptophan fluorescence. Although the enzyme contains 4 tryptophan residues, only one tryptophan, Trp47, has a suitable spatial location. The distance and the orientation of the dipole moment (1L a ) towards the isoalloxazine ring of the flavin cofactor enables us to monitor static or dynamic conformational changes in the active site (Figs 3 and 8). It should be emphasized that the proper position of a tryptophan residue towards the flavin cofactor is possible only in the dimeric structure of NADH oxidase. None of the tryptophan residues is close to the binding site of the flavin cofactor in the monomeric form of the enzyme (Fig. 3). The correlation of the decrease of tryptophan fluorescence and enzyme activity at low concentrations of urea strongly indicates that NADH oxidase forms functional dimers. Although the dimeric structure is obvious from the crystal structure [10], it was not apparent from the properties of the enzyme in solution [15]. The enzyme concentration (80-fold difference) had no effect on the urea–induced activity changes. This implies that the quaternary structure of NADH oxidase is intact in the range of urea concentrations at which enzyme activation occurred. The effect of urea on the level of the tertiary but not the secondary structure of the enzyme active site is indicated by: (a) a significant shift of the optimal temperature/activity profile to lower values and unaffected thermal stability of global folding (Fig. 5), (b) ellipticity changes in the aromatic region (Fig. 4B) and unperturbed ellipticity in the peptide region (Fig. 4A), and (c) a change in intensity and a slight shift in the maximum position of tryptophan fluorescence without any other apparent conformational changes. There is a well-known relationship between protein stability and protein flexibility/rigidity that assumes the rigidity of the polypeptide chain is a prerequisite for global protein stability. This relationship is supported both experimentally [1,4,41,42] and theoretically [3]. On the other hand, an inverse relationship exists between rigidity and enzymatic catalysis [43,44]. Urea-induced perturbation of the environment of the active site of NADH oxidase results in a partial exposure of buried Trp47 to solvent due to the increased dynamics of the active site. This is supported by (a) the slight shift in the maximum of tryptophan fluores- cence to higher wavelengths (Fig. 2), (b) insignificant changes in the ellipticity in the aromatic region, and, importantly (c) an increased fraction of tryptophan residues made accessible by the addition of a quencher such as acrylamide (Fig. 7). The higher dynamics of the active site is also indirectly reflected by (a) a )25 °C shift of the optimal temperature and (b) an increase in the apparent K M values for NADH binding to the enzyme (Table 1). The pro- nounced local destabilization of the active site and the unaffected global stability (Fig. 5) agrees with observations that the active site is the most flexible and thus the most labile part of the enzyme [2]. Therefore, the enzyme active site undergoes inactivation at milder conditions than is necessary for global unfolding [8]. Protein dynamics are characterized by motion on a broad spectrum of time scales. Not all time scales are relevant or significant for the stability or activity of an enzyme [45]. Because the lifetime of the excitation state of tryptophan is several nanoseconds [24], the activation of NADH oxidase must occur on a nanosecond time scale. This matches the observed rate constants for the process of stacking and unstacking bases in nucleic acids that is in the range of 10 6 )10 7 s )1 [46,47]. Fig. 8. Localization of Trp47, Wat42 and FMN in the dimeric interface of NADH oxidase from T. thermophilus. The individual monomers are designated by the letters A and B. The side chain of Trp47 is almost parallel to the flavin system. The water molecule forms hydrogen bonds to the backbone nitrogen of Trp47 and to O2 of the ribityl chain and is located in the middle of the flavin system with distances of 3.3 A ˚ toN10and3.7A ˚ to N5. Arrows indicate the possible flipping move- ment of the Trp47 side chain induced by low concentrations of urea (1.0–1.5 m) or by incoming substrate – the nicotinamide ring. The structure was drawn using VIEWER LITE 42. 4894 G. Z ˇ olda ´ k et al.(Eur. J. Biochem. 270) Ó FEBS 2003 The determination of the activation parameters shows (a) positive values of DH*andTDS* in the absence of urea and (b) a decrease of both parameters by % 15–20 kJÆmol )1 in the range of urea concentrations where the enzyme activa- tion occurs (Fig. 6). Because urea effects both of the activation parameters, the so-called enthalpy/entropy com- pensation [19,48,49], tells us that DH*andTDS*have opposite effects on the activity of the enzyme. The decrease in DH* diminishes the energy barrier of the reaction, whereas the decrease in DS* decelerates the catalytic action. The decrease in both parameters indicates that the ground and transition states are similar. The positive entropy seems surprising, at first glance, for a process that involves a tight interaction of NADH and FAD in the active site of the enzyme. This is probably due to the structural role of the water molecule in the enzyme active site [50]. The observed decrease in both activation parameters at % 1.5 M urea (Fig. 6) indicates that the difference between the ground and the transition state was reduced. The effect of a low urea concentration (1.25 M ) might be interpreted as a breakage of a noncovalent bond in the active site that reduces its rigidity resulting in the negative affects on the activity of the enzyme at low temperature. The value of the decrease in DH* % 15–20 kJÆmol )1 corresponds to the loss of about one hydrogen bond and the location of the structural water molecule in the active site. The crystal structure of the enzyme supports the inclusion of the structural water molecule in catalysis. Implications of the observed changes for enzyme catalysis and conformational changes Generally speaking, proteins have more mobility in the side chains than in the peptide backbone. This is especially true for side chains that participate in enzy- matic catalysis [51]. Analysis of the cofactor in the binding site from the crystal structure shows an inter- action between the flavin cofactor, the water molecule and Trp47 (Fig. 8) [10]. The side chain of Trp47 is almost parallel to the isoalloxazine ring; however, the elevated temperature factor and the weaker electron density indicate it is flexible. The 6–7 A ˚ gap between this side chain and the flavin ring contains some diffused electron density and one well defined and tightly bound water molecule. This water molecule forms hydrogen bonds with the backbone nitrogen of Trp47 and the O2 of the ribityl chain, and it is located in the middle of the flavin system with distances of 3–4 A ˚ to the flavin nitrogens [10]. NADH oxidase and three homologous flavoproteins form a novel flavoprotein family in which all members contain an aromatic amino-acid residue (Phe, Trp) that interacts with the isoalloxazine ring of the flavin cofactor [31]. This conserved interaction may play an important role in the catalytic mechanism of the flavoproteins. In fact, the crystal structure of the nonhomologous enzyme NADPH-cytochrome P450 oxidoreductase, with NADP + shows nicotinamide access to FAD is blocked by a tryptophan residue that stacks against the isoalloxazine in the flavin ring [52]. It has been proposed that the tryptophan residue acts as a ÔswitchÕ – when reduced substrate, NADPH, enters the active site an interaction between the isoalloxazine and nicotinamide rings is able to displace the tryptophan residue. After the substrate is oxidized to NADP + the interaction between the nicotin- amide ring and the flavin cofactor weakens and the indole ring of the tryptophan displaces the oxidized substrate from the binding site. This mechanism may be common in the catalytic actions of flavoproteins. A similar movement of aromatic amino-acid residues in active sites of enzymes after interaction with substrate has been proposed for other flavoproteins [53]. In NADH oxidase, the binding site of the incoming substrate (NADH) is blocked by a water molecule tightly bound to the flavin cofactor and Trp47. Thus, the nicotin- amide ring has to displace the water molecule to achieve the proper position for hydride transfer to the cofactor. Breaking of the hydrogen bond(s) which displaces the water molecule, and the concomitant local conformational change in the enzyme active site might be the rate-limiting step in NADH oxidation. We speculate that this is the mechanism by which urea at low concentrations activates NADH oxidase at room temperature. It perturbs hydrogen bond(s) in the active site between the flavin cofactor, the water molecule and Trp47 (decrease in DH*by% 15–20 kJÆ mol )1 ). This leads to release of strain in the active site and an increase in the dynamics of the Trp47 side chain (decrease in the tryptophan fluorescence and a slight red shift of the fluorescence maximum), weakens the water molecule’s interaction with the flavin cofactor and opens the active site (increased f a value from quenching experiment). The isodichroic point in the aromatic region of the circular dichroism spectrum (Fig. 4B) indicates a two state character for the conformational change. The flavin-aromatic amino acids probably move from closed (buried and rigid) to open (exposed and dynamic) in the active site. In fact, it has been shown that the equilibrium between solvent-exposed and ÔburiedÕ forms of the flavin cofactor may be important in the catalytic mechanism of flavoproteins [54]. Although our results strongly suggest that Trp47 has a role in enzymatic catalysis, they are not conclusive. The current work; however, identifies Trp47 as a good candidate for site directed mutagenesis to elucidate the rate-limiting step in the NADH oxidase catalysis. The increased dynamics due to urea-induced perturbation of hydrogen bonds decreases the energy needed to go from the ground state to the transition state in the active site of NADH oxidase from T. thermophilus at room temperature. The changes in the dynamics of the active site of NADH oxidase at room temperature caused by changes in solvent properties, pH, and the presence of chaotropic anions further indicate an important role for dynamics/plasticity in the enzyme catalysis. Acknowledgements The authors would like to thank the Fonds der Chemischen Industrie for financial support. We are also grateful for support through grants No. D/01/02768 from the Deutsche Akademische Austauschdienst (DAAD), no. 1/8047/01 and 1/0432/03 from the Slovak Grant Agency, and an internal grant from the UPJS Faculty of Sciences (VVGS 2002) for E.S and G.Z ˇ . We thank Norbert Grillenbeck for his technical assistance. The authors wish to thank Linda Sowdal for her invaluable editorial help in preparing the manuscript. Ó FEBS 2003 Flexibility of the NADH oxidase active site (Eur. J. Biochem. 270) 4895 References 1. Vihinen, M. (1987) Relationship of protein flexibility to thermo- stability. Protein Eng. 1, 477–480. 2. Shoichet, B.K., Baase, W.A., Kuroki, R. & Matthews, B.W. (1995) A relationship between protein stability and protein func- tion. Proc. Natl Acad. Sci. USA 92, 452–456. 3. Tang, K.E. & Dill, K.A. (1998) Native protein fluctuations: the conformational-motion temperature and the inverse correlation of protein flexibility with protein stability. J. Biomol. Struct. Dyn. 16, 397–411. 4. Za ´ vodszky, P., Kardos, J., Svingor, A. & Petsko, G.A. (1998) Adjustment of conformational flexibility is a key event in the thermal adaptation of proteins. Proc. Natl Acad. Sci. USA 95, 7406–7411. 5. Thomas, T. & Cavicchioli, R. (2000) Effect of temperature on stability and activity of elongation factor 2 proteins from Antarctic and thermophilic methanogens. J. Bacteriol. 182, 1328–1332. 6. Lindstrøm-Lang, K.M. & Schellman, J.A. (1959) In Enzymes (Boyer, P., ed.) pp. 413–510. Academic, New York. 7. Koshland, D.E. (1958) Application of a theory of enzyme spe- cificity to protein synthesis. Proc. Natl Acad. Sci. USA 44, 98–104. 8. Tsou, C L. (1995) Inactivation precedes overall molecular con- formation changes during enzyme denaturation. Biochim. Biophys. Acta 1253, 151–162. 9. Vieille, C. & Zeikus, G.J. (2001) Hyperthermophilic enzymes: sources, uses, and molecular mechanisms for thermostability. Microbiol. Mol. Biol. Rev. 65, 1–43. 10. Hecht, H.J., Erdmann, H., Park, H.J., Sprinzl, M. & Schmid, R.D. (1995) Crystal structure of NADH oxidase from Thermus thermophilus. Nat. Struct. Biol. 2, 1109–1114. 11. Lovering, A.L., Hyde, E.I., Searle, P.F. & White, S.A. (2001) The structure of Escherichia coli nitroreductase complexed with nico- tinic acid: three crystal forms at 1.7 A ˚ , 1.8 A ˚ and 2.4 A ˚ resolution. J. Mol. Biol. 309, 203–213. 12. Haynes, C.A., Koder, R.L., Miller, A F. & Rodgers, D.W. (2002) Structures of nitroreductase in three states: effects of inhibitor binding and reduction. J. Biol. Chem. 227, 11513–11520. 13. Gerstein, M., Lesk, A.M. & Clothia, C. (1994) Structural mechanisms for domain movements in proteins. Biochemistry 33, 6739–6743. 14. Park, H.J., Kreutzer, R., Reiser, C.O.A. & Sprinzl, M. (1992) Molecular cloning and nucleotide sequence of the gene encoding a H 2 O 2 -forming NADH oxidase from the extreme thermophilic Thermus thermophilus HB8 and its expression in Escherichia coli. Eur. J. Biochem. 205, 875–879. 15.Park,H.J.,Reiser,C.O.A.,Kondruweit,S.,Erdmann,H., Schmid, R.D. & Sprinzl, M. (1992) Purification and character- ization of a NADH oxidase from the thermophile Thermus thermophilus HB8. Eur. J. Biochem. 205, 881–885. 16. Laemmli, U.K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680– 685. 17. Pace, C.N., Vajdos, F., Fee, L., Grimsley, G. & Gray, T. (1995) How to measure and predict the molar absorption coefficient of a protein. Protein Sci. 4, 2411–2423. 18. Schmidt, H L., Sto ¨ cklein, W., Danzer, J., Kirch, P. & Limbach, B. (1986) Isolation and properties of H 2 O-forming NADH oxidase from Streptococcus faecalis. Eur. J. Biochem. 156, 149–155. 19. Cornish-Bowden, A. (2002) Enthalpy-entropy compensation: a phantom phenomenon. J. Biosci. 27, 121–126. 20. Parker, C.A. & Reeses, W.T. (1960) Correction of fluorescence spectra and measurement of fluorescence quantum efficiency. Analyst (London) 85, 587–600. 21. Yengo, C.M., Fagnant, L., Chrin, L., Rovner, A.S. & Berger, C.L. (1998) Smooth muscle myosin mutants containing a single tryptophan reveal molecular interactions at the actin–binding interface. Proc.NatlAcad.Sci.USA95, 12944–12949. 22. Valeur, B. & Weber, G. (1977) Resolution of the fluorescence excitation spectrum of indole into the 1La and 1Lb excitation bands. Photochem. Photobiol. 25, 441–444. 23. Chang, W.I. & Matthews, K.S. (1995) Role of Asp274 in lac repressor: diminished sugar binding and altered conformational effects in mutants. Biochemistry 34, 9227–9234. 24. Lakowicz, J.R. (2000) Principles of Fluorescence Spectroscopy, 2nd edn. Plenum Press, New York. 25. Lehrer, S.S.(1971) Solute perturbation of protein fluorescence. The quenching of the tryptophanyl fluorescence of model compounds and of lysozyme by iodide ion. Biochemistry 10, 3254–3263. 26. Burstein, E.A., Vedenkina, N.S. & Ivkova, M.N. (1973) Fluores- cence and the location of tryptophan residues in protein molecules. Photochem. Photobiol. 18, 263–279. 27. Elodi, P. & Lakatos, S. (1973) The anomalous absorption of tryptophan. Eur. J.Biochem. 36, 45–52. 28. Truhlar, G.D. & Kohen, A. (2001) Convex Arrhenius plots and their interpretation. Proc.NatlAcad.Sci.USA98, 848–851. 29. Eftink, M.R. & Ghiron, C.A. (1981) Fluorescence quenching studies with proteins. Anal. Biochem. 114, 199–227. 30. Thomas, T.M. & Scopes, R.K. (1998) The effects of temperature on the kinetics and stability of mesophilic and thermophilic 3-phosphoglycerate kinases. Biochem. J. 330, 1087–1095. 31. Parkinson, G.N., Skelly, J.V. & Neidle, S. (2000) Crystal structure of FMN-dependent nitroreductase from Escherichia coli B: a prodrug-activating enzyme. J. Med. Chem. 43, 3624–3631. 32. Wolosiuk, R.A. & Stein, M. (1990) Modulation of spinach chloroplast NADP-glyceraldehyde-3-phosphate dehydrogenase by chaotropic anions. Arch. Biochem. Biophys. 279, 70–77. 33. Fan, Y X., Ju, M., Zhou, J M. & Tsou, C L. (1996) Activation of chicken liver dihydrofolate reductase by urea and guanidine hydrochloride is accompanied by conformational change at the active site. Biochem. J. 315, 97–102. 34. Narayanasami, R., Nishimura, J.S., McMillan, K., Roman, L.J., Shea, T.M., Robida, A.M., Horowitz, P.M. & Masters, B.S.S. (1997) The influence of chaotropic reagents on neuronal nitric oxide synthase and its flavoprotein module. Urea and guanidine hydrochloride stimulate NADPH-cytochrome c reductase activity of both proteins. Nitric Oxide 1, 39–49. 35. Zhang, H J., Sheng, X R., Pan, X M. & Zhou, J M. (1997) Activation of adenylate kinase by denaturants is due to the increasing conformational flexibility at its active sites. Biochem. Biophys. Res. Commun. 238, 382–386. 36. Das, M. & Dasgupta, D. (1998) Enhancement of transcriptional activity of T7 RNA polymerase by guanidine hydrochloride. FEBS Lett. 427, 337–340. 37. Deshpande, R.A., Kumar, A.R., Khan, M.I. & Shankar, V. (2001) Ribonuclease Rs from Rhizopus stolonifer: lowering of optimum temperature in the presence of urea. Biochim. Biophys. Acta 1545, 13–19. 38. Inui, T., Ohkubo, T., Urade, Y. & Hayaishi, O. (1999) Enhance- ment of lipocalin-type prostaglandin D synthase enzyme activity by guanidine hydrochloride. Biochem. Biophys. Res. Commun. 266, 641–646. 39. Kujo, C. & Oshima, T. (1998) Enzymological characteristics of the hyperthermostable NAD-dependent glutamate dehydrogenase from the archaeon Pyrobaculum islandicum and effects of dena- turants and organic solvents. Appl. Envir. Microbiol. 64, 2152– 2157. 40. Jaenicke, R. & Bo ¨ hm, G. (2001) Thermostability of proteins from Thermotoga maritima. Methods Enzymol. 334, 438–469. 4896 G. Z ˇ olda ´ k et al.(Eur. J. Biochem. 270) Ó FEBS 2003 [...]... Struct Biol 3, 821–832 44 Alper, K.O., Singla, M., Stone, J.L & Bagdassarian, C.K (2001) Correlated conformational fluctuations during enzymatic catalysis: Implications for catalytic rate enhancement Protein Sci 10, 1319–1330 45 Hernandez, G., Jenney, F.E.J., Adams, M.W & Le Master, D.M (2000) Millisecond time scale conformational flexibility in a hyperthermophile protein at ambient temperature Proc Natl... (1998) 1.8 A crystal structure of the major NAD(P)H: FMN oxidoreductase of a bioluminescent bacterium, Vibrio fischeri: overall structure, cofactor and substrate-analog binding, and comparison with related flavoproteins J Mol Biol 280, 259–273 Zheng, Y., Dong, J., Palfey, B.A & Carey, P.R (1999) Using Raman spectroscopy to monitor the solvent-exposed and ÔburiedÕ forms of flavin in p-hydroxybenzoate hydroxylase... Protein Sci 10, 661–667 Wolfenden,R.&Snider,M.J.(2001)The depth of chemical time and the power of enzymes as catalysts Acc Chem Res 34, 938–945 Sitnitsky, A.E (1994) Conformational dynamics of protein side chains and enzyme–substrate interaction J Biomol Struct Dyn 12, 475–486 Hubbard, P.A., Shen, A.L., Paschke, R., Kasper, C.B & Kim, J.-J.P (2001) NADPH-cytochrome P450 oxidoreductase Structural basis for. .. (1979) Laser temperature-jump study of stacking in adenylic acid polymers Biochemistry 18, 5757–5762 47 Hammes, G.G (2002) Multiple conformational changes in enzyme catalysis Biochemistry 41, 8221–8228 48 Lumry, R & Rajender, S (1970) Enthalpy-entropy compensation phenomena in water solutions of proteins and small 49 50 51 52 53 54 molecules: a ubiquitous property of water Biopolymers 9, 1125– 1227... Flexibility of the NADH oxidase active site (Eur J Biochem 270) 4897 41 Daniel, R.M., Cowan, D.A., Morgan, H.W & Curran, M.P (1982) A correlation between protein thermostability and resistance to proteolysis Biochem J 207, 641–644 42 Tsai, A.M., Udovic, T.J & Neumann, D.A (2001) The inverse relationship between protein dynamics and thermal stability Biophys J 81, 2339–2343 43 Cannon, W.R., Singleton, . Role of conformational flexibility for enzymatic activity in NADH oxidase from Thermus thermophilus Gabriel Z ˇ olda ´ k 1 ,Ro ´ bert. explains the role of the homodimeric structure of NADH oxidase. Keywords: NADH oxidase; conformational dynamics; flavo- proteins; fluorescence quenching; Thermus