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Protein Science (2001), 10:339-351.
Copyright © 2001 The Protein Society

Effects of i-propanol on the structural dynamics of Thermomyces lanuginosa lipase revealed by tryptophan fluorescence

Keng Zhu, Arimatti Jutila, Esa K.J. Tuominen and Paavo K.J. Kinnunen

Helsinki Biophysics and Biomembrane Group, Department of Medical Chemistry, Institute of Biomedicine, FIN-00014 University of Helsinki, Finland

Reprint requests to: Dr. Paavo K.J. Kinnunen, Helsinki Biophysics and Biomembrane Group, Department of Medical Chemistry, Institute of Biomedicine, FIN-00014 University of Helsinki, Finland; e-mail: Paavo. Kinnunen{at}Helsinki.Fi; fax: 358-9-1918276.

(RECEIVED May 30, 2000; FINAL REVISION November 15, 2000; ACCEPTED November 16, 2000)

Article and publication are at www.proteinscience.org/cgi/doi/10.1110/ps.21201.


   Abstract
TOP
 Abstract
Introduction
 Results
 Discussion
 Conclusion
 Materials and methods
 References
 
Influence of isopropanol (iPrOH) on the structural dynamics of Thermomyces lanuginosa lipase (TLL) was studied by steady-state, time-resolved, and stopped-flow fluorescence spectroscopy, monitoring the intrinsic emission of Trp residues. The fluorescence of the four Trps of the wild-type enzyme report on the global changes of the whole lipase molecule. To monitor the conformational changes in the so-called "lid," an {alpha}-helical surface loop, the single Trp mutant W89m (W117F, W221H, W260H) was employed. Circular dichroism (CD) spectra revealed that iPrOH does not cause major alterations in the secondary structures of the wild-type TLL and W89m. With increasing [iPrOH], judged by the ratio of emission intensities at 350 nm and 330 nm, the average microenvironment of the Trps in the wild-type TLL became more hydrophobic, whereas Trp89 of W89m moved into a more hydrophilic microenvironment. Time-resolved fluorescence measurements revealed no major changes to be induced by iPrOH neither in the shorter fluorescence lifetime component ({tau}1 = 0.5–1.2 ns) for the wild-type TLL nor in the longer fluorescence lifetime component ({tau}2 = 4.8–6.0 ns) in the wild-type TLL and the W89m mutant. Instead, for W89m on increasing iPrOH from 25% to 50% the value for {tau}1 increased significantly, from 0.43 to 1.5 ns. The shorter correlation time {phi}1 of W89m had a minimum of 0.08 ns in 25% iPrOH. Judged from the residual anisotropy r{infty} the amplitude of the local motion of Trp89 increased upon increasing [iPrOH] 10%. Stopped-flow fluorescence spectroscopy measurements suggested the lid to open within {approx}2 ms upon transfer of W89m into 25% iPrOH. Steady-state anisotropies and longer correlation times revealed increasing concentrations of iPrOH to result also in the formation of dimers as well as possibly also higher oligomers by TLL.

Keywords: Thermomyces lanuginosa lipase; conformational dynamics; fluorescence spectroscopy

Abbreviations: carboxy-DCFDA, carboxy-2',7'-dichlorofluorescein diacetate • CD, circular dichroism • DMSO, dimethylsulfoxide • EDTA, ethylenediaminetetraacetic acid • Hepes, N-(2-hydroxyethyl) piperazine-N'-2-ethanesulphonic acid • TLL, Thermomyces lanuginosa lipase • I350/I330, ratio of fluorescence intensities at 350 and 330 nm • iPrOH, isopropanol • R.A., relative activity • RFI, relative fluorescence intensity • W89m, single Trp TLL mutant with substitutions W117F, W221H, and W260H • {tau}, fluorescence lifetime • {phi}, rotational correlation time • r{infty}, residual anisotropy.


   Introduction
TOP
 Abstract
 Introduction
Results
 Discussion
 Conclusion
 Materials and methods
 References
 
Lipases (triacylglycerol hydrolases, E.C. 3.1.1.3) constitute a large family of enzymes and are widely distributed in many organisms. Their function is to cleave a fatty acid chain either in position 1 or 3 of a triglyceride. Lipases have also become popular biocatalysts allowing for organic reactions to be performed under mild conditions (Kim et al. 1997). True lipases are distinguished from esterases by the characteristic interfacial activation they exhibit at lipid–water interfaces (Sarda and Desnuelle 1958; Pieterson et al. 1974). Several mechanisms have been forwarded to explain this property (Volwerk and de Haas 1982; Simons et al. 1997). These include an increased substrate concentration at the interface (Brockman et al. 1973), a better orientation of the scissile ester bond (Wells 1974), a reduction in the water shell around the ester molecule in water (Brockerhoff 1968), as well as conformational changes of the enzyme leading to an optimized active site geometry and enhanced catalytic activity (Sarda and Desnuelle 1958; Entressangles and Desnuelle 1974).

The amino acid sequence of TLL consists of 269 residues, including four tryptophans (Martinelle et al. 1995). The crystal structure of TLL has been solved at 1.8 Å resolution (Derewenda et al. 1994a). Accordingly, TLL consists of a single, roughly spherical domain containing a central eight-stranded, predominately parallel ß-pleated sheet, with five interconnecting {alpha}-helices, compacted to a volume of ~9.7 x 103 Å3. The active site contains a Ser(146)–Asp(201)–His(258) catalytic triad, closely reminiscent to that seen in the serine proteases (Brady et al. 1990). The active site serine (S146A) seems to be involved in maintaining the structure of TLL and its substitution leads to substantial conformational alterations as well as in different substrate binding affinities (Peters et al. 1998). The {alpha}-helical surface loop (amino acids 86–93) constituting the "lid" lies directly over the active site S146 (Brady et al. 1990; Berg et al. 1998), and is highly mobile in the crystals (Derewenda et al. 1994b). The lid of TLL contains a single Trp at position 89. Site-directed mutagenesis of this residue (Holmquist et al. 1994, 1995) showed that it is important for efficient hydrolysis of tributyrin and that it plays a role in the catalytic steps after the absorption of the lipase to the substrate interface. Trp89 has been concluded to influence the binding of substrates into the active site in a qualitative and not a quantitative way (Holmquist et al. 1995).

Two distinct conformations, "closed" and "open," inactive and active, respectively, have been proposed for TLL (Svendsen et al. 1997). Accordingly, in an aqueous medium access of the substrate to the catalytic triad is blocked by the lid and the conformation is closed. Instead, in complexes with substrate analogs or serine protease inhibitors the lid helix is displaced and the active site is exposed, representing the open conformation (Brzozowski et al. 1991; Derewenda et al. 1992; Lawson et al. 1994). Interfacial activation of TLL would thus be associated with a significant conformational change of the lid (Martinelle et al. 1995), its opening generating a large hydrophobic surface that also includes the active site. The ability of TLL to efficiently form the acyl-enzyme intermediate at the interface thus requires a number of separate steps: (1) The enzyme must adapt the active lid open conformation; (2) the lid has to attach to the interface; and (3) the active site has to be saturated with the substrate (Holmquist et al. 1994). This model is developed further in a recent work by Cajal et al. (2000) reporting that on binding to liposomes, the lipase can adopt two different conformations, catalytically active and inactive, depending on the composition and curvature of the surface. Similar ideas were forwarded by Berg et al. (1998) emphasizing the importance of electrostatics in the interaction between the lipase and the surface.

Not only lipids (Berg et al. 1998; Cajal et al. 2000) and detergents (van Tilbeurgh et al., 1993; Martinelle et al. 1995; Hermoso et al. 1997; Simons et al. 1997; Graupner et al. 1999) but also organic solvents induce the opening of the lid. Accordingly, when the lipase from Candida rugosa was crystallized in the presence of 2-methyl-2,4-pentanediol the protein remains in the open conformation (Ransac et al. 1995a).

Dissolving C. rugosa lipase in different concentrations of polar organic solvents yielded in enzyme preparations with varying activities and opening of the lid was proposed as the underlying mechanism (Chamorro et al. 1998). This is in accordance with the results from Poisson-Boltzmann calculations indicating displacement of the lid of Rhizomucor miehei lipase upon decrease in the dielectricity of the medium (Jääskeläinen et al. 1999). These data suggest that electrostatic interactions play an important role in this process. Also, there is a recent comparison of the changes in the conformation of three different microbial lipases induced by iPrOH (Graupner et al. 1999).

Studies on the conformation of lipases in solution are scarce. Yet, the use of these enzymes as catalysts in synthesis almost invariably involves their action in an organic solvent (Graupner et al. 1999). In the present study, we used fluorescence spectroscopy to analyze conformational changes of TLL induced by iPrOH. More specifically, steady-state, time resolved, and stopped-flow fluorescence spectroscopy were employed to monitor the changes caused by iPrOH in the intrinsic Trp fluorescence of the wild-type TLL as well as the single Trp mutant W89m. The functional integrity of the active site was assessed by measuring their catalytic activity towards a water-soluble fluorogenic substrate. Our data indicate that iPrOH induces the opening of the lid as well as the oligomerization of TLL.


   Results
TOP
 Abstract
 Introduction
 Results
Discussion
 Conclusion
 Materials and methods
 References
 
Effects of iPrOH on the catalytic activity
The relative catalytic activities of wild-type and W89m TLL towards the nonspecific esterase substrate carboxy-DCFDA as function [iPrOH] are depicted in Figure 1Go. Both enzymes revealed the same tendency, their activities first increasing at low concentrations of iPrOH (<10% and 15%, respectively), and then decreasing, upon exceeding 10% and 15% iPrOH for the wild type and W89m, respectively. The catalytic activity of the TLLs were significantly augmented in <=25% [iPrOH] when the concentration of substrate was increased from 50 O——O, •——• (o—o, •—•) to 100 {square}——{square}, {blacksquare}——{blacksquare} ({nu}—{nu}) µM.. This enhancement was more pronounced for the wild type than W89m. Both wild-type and W89m TLL were almost completely inactive in 50% iPrOH.



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  		Fig. 1. Relative catalytic activities (R.A.) of the wild-type (open symbols) and W89m TLL (solid symbols) towards carboxy-DCFDA measured as a function of [iPrOH]. The concentration of the substrate was 50 µM (open circle, solid circle) or 100 µM (open square, solid square). Relative activity values of wild-type and W89m are normalized to 1.0 at [substrate] = 100 µM in the absence of iPrOH. The concentration of the lipases was 1.0 µM in 20 mM Hepes, 0.1 mM EDTA at pH 7.0. Temperature was 25°C.

 
Effects of iPrOH on the secondary structure
CD spectra were recorded in the amide region (200–250 nm) to observe possible changes in the secondary structure of TLL. Both the wild-type and W89m TLL revealed CD spectra typical of a predominantly {alpha}-helical protein, with the characteristic double minima at 208 and 222 nm (Fig. 2Go). The {alpha}-helicities for the wild-type and W89m TLL calculated from CD are 33% and 25%, respectively, whereas 26 % was obtained from the X-ray structure of the wild-type TLL (Laskowski et al. 1997). Increasing [iPrOH] up to 50% caused no changes in the ellipticities at 215 nm and 222 nm, corresponding to {alpha}-helix and ß-sheet structures, respectively. These data thus verify a lack of major alterations due to iPrOH in the secondary structures of the wild-type and W89m TLLs.



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  		Fig. 2. Far UV CD spectra of the wild-type TLL (A) and W89m (B) in 0 (solid square), 25 (open circle) and 50% (x) iPrOH. The concentration of lipases was 0.2 mg/mL, corresponding to 6.7 µM, in 20 mM Hepes, 0.1 mM EDTA at pH 7.0. Temperature was 25°C.

 
Effects of iPrOH on the tertiary structure
Near-UV CD spectra were recorded in the 240–340 nm region where the spectrum reflects the environmental asymmetry of aromatic amino-acid residues (Fig. 3Go). TLL has a number of aromatic residues, and their specific contributions to the near-UV spectra cannot be assigned. A maximum at 268 nm is not seen in the mutants, thus probably resulting from the removal of one or several of the tryptophans. Different iPrOH concentrations do not induce significant changes in the wild-type TLL near-UV spectra, suggesting minimal conformational effect due to this solvent. Instead, for W89m the intensity of the negative band in the Phe range 240–260 nm decreases remarkably upon increasing iPrOH concentration from 25% to 50%.



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  		Fig. 3. Near UV CD spectra of the wild-type TLL (A) and W89m (B) in 0% (solid square), 25% (open circle) and 50% (x) iPrOH. The concentration of lipases was 1 mg/mL, corresponding to 33.3 µM, in 20 mM Hepes, 0.1 mM EDTA at pH 7.0. Temperature was 25°C.

 
Steady-state fluorescence measurements
The wavelength of the emission maximum {lambda}max for Trp depends on its microenvironment. A low polarity, hydrophobic microenvironment is characterized by {lambda}max {approx}331 nm, whereas for Trp in an aqueous phase {lambda}max is 350–353 nm (Brustein et al. 1973). When conventional fluorescence emission spectra of the wild type and W89m in buffer were measured with excitation wavelength at 295 nm, the maximum emission wavelengths for the wild type and W89m were 342 and 345 nm, respectively. In 50% iPrOH, the values of {lambda}max for the wild-type and W89m TLL were 341 and 346.5 nm, respectively. The increase in {lambda}max by 1.5 nm for W89m indicates the vicinity of this residue to become more polar in iPrOH. Changes in fluorescence intensities induced by iPrOH are shown in Figure 4Go. For the wild-type TLL, fluorescence intensity first increased up to 25% iPrOH and then decreased at higher concentrations. For W89m, similar tendency was evident, however maximum emission intensity was observed between 15% to 30% iPrOH. Notably, there are 11 Tyr residues in TLL (Derewenda et al. 1994c). A convenient way to exclude any interference of Tyr emission in the value for {lambda}max is to record synchronous fluorescence spectra (Miller 1979).



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  		Fig. 4. The maximum fluorescence intensity of the wild-type (solid circle) and W89m (open square) as a function of iPrOH concentration. The excitation wavelength was 295 nm, and the emission wavelengths for the wild type and W89m are 342 and 345 nm, respectively. Both excitation and emission bandpasses were 5 nm. Concentration of the lipase was 1 µM in 20 mM Hepes, 0.1 mM EDTA at pH 7.0. The temperature was 25°C. These data were retrieved from conventional emission spectra.

 
Accordingly, at {Delta}{lambda} = 60 nm (corresponding to the Stokes shift for Trp) only the fluorescence of Trp was observed. Synchronous spectrum for W89m in buffer revealed {lambda}max at 340 nm (Fig. 5Go, inset), suggesting Trp89 to reside in an environment of polarity intermediate between a hydrocarbon phase and water. Synchronous spectrum of the wild-type TLL in an aqueous buffer yielded {lambda}max = 340.5 nm, representing the average of its four Trp residues.



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  		Fig. 5. Effect of iPrOH on the fluorescence intensity ratio I350/I330 for the wild-type TLL (solid circle), W89m (open square), and free tryptophan (x). The excitation wavelength was 295 nm. Synchronous fluorescence spectra for W89m TLL (inset). Difference between the excitation and emission wavelengths {Delta}{lambda} was 60 nm. Concentration of the lipase was 1 µM in 20 mM Hepes, 0.1 mM EDTA at pH 7.0. The temperature was 25°C.

 
The ratio of emission intensities at 350 nm and at 330 nm (I350/I330) was used to further monitor the polarity of the microenvironments of Trp residues (Zhao and London 1986; Wang et al. 1997). Interestingly, opposite changes were induced by iPrOH in I350/I330 for the wild-type and W89m TLL (Fig. 5Go). Accordingly, for the wild-type TLL •——• (•—•) a minor, yet highly reproducible decrease in I350/I330 was evident. Instead, for W89m ({square}——{square} an increase was observed, in keeping with an increased polarity of the environment of its single Trp on transfer of the lipase into an iPrOH solution. For free Trp I350/I330 (x—x) decreased almost linearly upon increasing iPrOH concentration from 0% to 50%, in accordance with diminishing solvent polarity.

Rotational diffusion of fluorophores is the dominant cause of fluorescence depolarization. In proteins the mobility of fluorescent amino acid residues bears a close relationship with the overall state of a protein and any factor that affects its size, shape, or segmental flexibility will also affect the observed steady-state anisotropies (Lakowicz 1999). Accordingly, anisotropy reports on both conformational changes and the state of association of a protein. Fluorescence anisotropies of wild type and W89m as a function of [iPrOH] are shown in Figure 6Go and reveal similar overall behavior, the values for r first increasing steeply upon increasing [iPrOH] from 5% to 7.5%. Thereafter, exceeding 12% and 15% [iPrOH] for the wild-type TLL and W89m, respectively, values for the steady-state emission anisotropy decreased.



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  		Fig. 6. Steady-state fluorescence anisotropy for wild-type (solid circle) and W89m (open square) as a function of iPrOH. Concentration lipases was 1 µM in 20 mM Hepes, 0.1 mM EDTA at pH 7.0. The excitation wavelength was 295 nm, whereas emission was recorded at 342 and 345 nm for the wild-type and W89m TLL, respectively. Both excitation and emission bandpasses were 10 nm. Temperature was 25°C.

 
Time-resolved fluorescence measurements
Not only its spectral features but also the fluorescence lifetimes of Trp depend on its microenvironment, thus allowing to obtain further insight into the conformational changes of a protein. In <=10% [iPrOH], W89m could be fitted to a one-exponential decay with a lifetime of 5.4 ns and with a {chi}2 of 0.9–1.2. In >=15% [iPrOH], however, use of one-exponential decay for the analysis of the fluorescence decay curves resulted in {chi}2>2, indicating poor fit to the recorded data. Yet, for the latter data two-exponential decays gave values for varying 0.9 to 1.2 revealing good fitting. A representative fluorescence intensity decay curve is illustrated in Figure 7Go. Emission of the wild-type TLL decayed two-exponentially in all concentrations of iPrOH studied. The respective fluorescence lifetimes are compiled in Figure. 8GoA,B. The shorter lifetime component ({tau}1) of the wild type varied between 0.5 ns and 1.2 ns. For W89m the value of {tau}1 increased 3.4-fold at increasing concentrations of iPrOH, reaching 1.5 ns in 50% solvent. The longer fluorescence lifetime component ({tau}2) showed similar behavior for both wild-type and W89m TLL. In brief, the value of {tau}2 first increased at low iPrOH concentrations (<20% and 30% for the wild-type and W89m, respectively), and decreased when iPrOH concentration was further increased. The maximum values of {tau}2 of ~5.5 and 5.9 ns were observed in 20% and 30% iPrOH for the wild type and W89m, respectively. The fractional intensities of {tau}1 (Fig. 8CGo) were augmented with increasing [iPrOH]. However, the changes for the wild-type TLL were not as pronounced as for W89m. The most significant changes for W89m were observed between 25% to 50% iPrOH, with the fractional intensity of {tau}1 increasing from 3.5% to 23.7%.



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  		Fig. 7. Time-resolved fluorescence intensity decay of W89m in 25% iPrOH (B). The smooth lines 1 and 2 (inset) represent local fitted curves assuming either one- or two-exponential decays, respectively. (A) Residuals for the one-exponential (solid line) and two-exponential (dotted line) fitting of fluorescence intensity decay are shown. The respective values for {chi}2 were 1.13 and 4.35, respectively. The excitation wavelength was 298 nm, and the emission wavelength was 345 nm. The concentration of W89m was 1 µM.

 


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  		Fig. 8. Variation of the shorter ({tau}1, A) and longer ({tau}2, B) fluorescence lifetime components, and the fractional intensity of {tau}1 (C) for the wild-type (solid circle) and W89m (open square) TLL as a function of the concentration iPrOH. Conditions are as in Figure 1Go.

 
Time-dependent anisotropy decays provide information on the diffusive motions of a fluorophore during its excited lifetime. More specifically, these data can show whether a fluorophore is free to rotate over all angles, or if the environment surrounding the fluorophore restricts its angular Brownian motion. Moreover, these measurements allow to distinguish between decays of anisotropy due to a single process, and those involving multiple rotational motions (Lakowicz 1999). We measured fluorescence anisotropy decays of W89m (Fig. 9Go) as a function of [iPrOH] and calculated the rotational correlation times ({phi}1 and {phi}2) and residual anisotropy values (r{infty}) from these data. The correlation times of W89m are compiled in Figure 10GoA. The value of the shorter correlation time {phi}1 ({Delta}—{Delta}) started to decrease in 20% iPrOH, reaching a minimum of 0.08 ns in 25% iPrOH. Interestingly, the longer correlation time {phi}2 {blacksquare}——{blacksquare} ({gamma}—{gamma}) increased stepwise, obtaining values of about 20, 40, and 58 ns upon progressively increasing concentration of iPrOH from 0% to 30%. Thereafter, at [iPrOH] > 30%, the value for {phi}2 became too long to be fitted properly. Residual anisotropy (x—x) decreased upon increasing [iPrOH] and its overall values were {approx}0.17 in <=10% [iPrOH] and {approx}0.11 in >=15 % [iPrOH] (Fig. 10GoB).



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  		Fig. 9. Time-resolved anisotropy decay of W89m in 25% iPrOH (B). The .solid and open symbols correspond to I|| and I{bot}, respectively. The smooth lines represent fitted curves. (A) The residuals for the fitting of I{bot} (solid line) and I{bot} (dotted line) fluorescence intensity decay are shown. The excitation wavelength was 298 nm, and the emission wavelength was 345 nm. The concentration of W89m was 1 µM.

 


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  		Fig. 10. (A) The shorter ({Delta}) and longer (solid square) correlation time ({phi}1 and {phi}2, respectively) of W89m as a function of iPrOH concentration. (B) The residual anisotropy (x) for W89m as a function of concentration of iPrOH. The excitation wavelength was 298 nm, and the emission wavelength 345 nm. The concentration of W89m was 1 µM.

 
Stopped-flow fluorescence measurements
Time courses of the iPrOH-induced conformational changes in the >ms time domain were followed by recording the intrinsic Trp fluorescence as a function of time after rapid mixing. A kinetic trace measured in the 0–25 ms regime after the transfer of one µM W89m from the aqueous buffer into 25% iPrOH (final concentrations) showed a rapid increase in fluorescence intensity within {approx}2 ms (data not shown). A control experiment using buffer instead of iPrOH showed no changes in fluorescence intensity, thus confirming that the increase in fluorescence can be attributed to conformational changes induced by iPrOH.


   Discussion
TOP
 Abstract
 Introduction
 Results
 Discussion
Conclusion
 Materials and methods
 References
 
Studies on the C. rugosa lipase have indicated iPrOH to induce the opening of the lid (Chamorro et al. 1998). Accordingly, it was of interest to study the effects of this solvent on the structural dynamics of the closely related lipase from Thermomyces lanuginosa.

Wild-type TLL contains four Trps, two of which, W89 and W117, are located in two different {alpha}-helices and the other two, W221 and W260, in two different ß-pleated sheets. Changes in the intrinsic fluorescence of the wild-type TLL thus report on the global conformational changes of this protein. Accordingly, we were particularly interested in the single Trp mutant W89m, in which the fluorescent residue is contained in the lid, an {alpha}-helical structure undergoing a major reorientation upon the lid opening.

Decreasing peak intensity in the near-UV spectra of W89m at 50% iPrOH can be attributed to intramolecular fluctuations that lead to loss of environmental asymmetry of aromatic residues (Woody and Dunker 1996). Lack of simultaneous changes in the far-UV CD spectra indicates a native-like secondary structure (Fig. 2Go ).

Thus the CD data suggest partial unfolding of the protein tertiary structure in 50% iPrOH. Time resolved fluorescence of the single Trp in W89m demonstrated decreased residual anisotropy on increasing iPrOH concentration from 25% to 50%. Increase in Trp mobility together with the changes in near-UV CD suggest enhanced overall conformational fluctuations. This is in keeping with our results from the esterase activity measurements with decreased enzymatic activity for the W89m in 50% iPrOH. Our results thus demonstrate loosening of the tertiary structure while secondary structure is preserved, indicating a molten globule state (Dolgikh et al. 1985).

This is in keeping with the suggestion that the function of interfacially active enzymes can be significantly influenced by interactions with apolar molecules which may help to maintain and modulate rather than to destroy their three-dimensional structures (Ransac et al. 1995b; Graupner et al. 1999). Yet, fluorescence spectroscopy did reveal major alterations in the conformational dynamics of TLL to be induced by iPrOH, which had significant effects also on the catalytic activity of the lipase. The major conclusion is the opening of the lid in the presence of iPrOH. A complex pattern of alterations in the different parameters measured was evident as a function of iPrOH concentration.

Notably, the lack of coinciding changes with respect to [iPrOH] is not unexpected. This stems from the very nature of fluorescence, which allows to distinguish between changes in fluorophore mobility (r, r{infty}, and the derived rotational correlation times) and subtle changes in its electronic transitions (reflected by {lambda}max, intensity, and {tau}1 and {tau}2) which are further determined by different constraints imposed by the environment of the fluorophore. Accordingly, whereas the recorded fluorescence parameters obviously are connected, they also differ in their responses to the solvent-induced changes in the structure of the protein. Moreover, on increasing [iPrOH], it is expected that the solvent can induce more subtle changes in protein conformational dynamics, preceeding the closed -> open conformational transition. Also the presence of the soluble substrate is likely to influence the conformational equilibrium of the lipase, most likely shifting it towards the open conformation, thus representing another complication affecting the concentration dependence of the effects of iPrOH.

Steady-state conformational changes in TLLs
At [iPrOH] = 50%, there was a blue shift by 1 nm in the {lambda}max of the wild-type TLL. The opposite was true for W89m, with a small red shift (from 345 to 346.5 nm), suggesting an increment in the polarity of the microenvironment of the Trp in the lid. Fluorescence intensities changed dramatically on increasing [iPrOH] (Fig. 4Go). The quantum yield of Trp fluorescence increases when the polarity of microenvironment decreases (She et al. 1998). Accordingly, this indicates that iPrOH caused the Trps in wild-type TLL to move into less polar average microenvironments. Fluorescence intensity of W89m increased only slightly, by ~3% in 30% iPrOH where after it decreased when 40% iPrOH was exceeded, thus indicating the microenvironment of Trp89 to become more polar. The observed red shift revealed by an increase in I350/I330 for W89m in 50% iPrOH (Fig. 5Go) supports this conclusion (Zhao and London 1986; Wang et al. 1997). The initial, minor increase in emission intensity at [iPrOH] <= 30% suggests that Trp89 in the absence of iPrOH, (i.e., in the closed conformation) contacts a vicinal residue slightly quenching its fluorescence. As Trp89 contributes ~61% to the fluorescence of the wild-type TLL (Stobiecka et al. 1998), the change in the quantum yield of this residue has a significant effect on the Trp fluorescence of this protein.

Interestingly, the fluorescence intensity of the wild-type TLL started to decrease when [iPrOH] exceeded 25%. Accordingly, this can be concluded to result from the exposure of Trp89 to a more polar environment. Our stopped-flow measurements on the mixing of W89m with iPrOH revealed fluorescence intensity to reach a maximum in <2 ms (data not shown), indicating the opening of the lid to be rapid. Compared to the ~3% increase in fluorescence intensity by 25% iPrOH in steady-state measurements, the emission increased {approx}2.5-fold in the stopped-flow chamber under these conditions. A reason for this difference is likely to be a slow solvation of iPrOH into the buffer, resulting in incomplete solvent mixing within 2 ms. Accordingly, when Trp89 moves into the incompletely solvated iPrOH upon opening of the lid, its quantum yield should increase.

Fluorescence lifetimes provide further insight into the changes in conformational dynamics induced by iPrOH. Intensity decay of W89m is one-exponential in <=10% iPrOH, whereas, two-exponential fitting was required in >=15% iPrOH (Fig. 8Go). The two populations of Trp(s) with different fluorescence lifetimes have been ascribed to different Trp rotamer conformations (Chen et al. 1991).

In proteins, the shorter lifetime component {tau}1 has been further suggested to reflect interactions of the surface-exposed Trp(s) with the solvent (Graupner et al. 1999). The crystal structure of the open conformation of TLL revealed Trp89 to be exposed on the protein surface (A.M. Brzozowski, pers. comm.), in accordance with the pronounced increase in {tau}1 on increasing [iPrOH]. Notably, the fractional intensity of {tau}1 increased from 3.5% to 23.2% with [iPrOH], indicating a considerable shift in the conformational equilibrium of W89m. For the wild-type TLL with four Trps a similar change in the fractional intensity of {tau}1 could not be resolved.

Changes in conformational dynamics of TLL induced by isopropanol
Decrease in the steady-state anisotropy r of a fluorophore corresponds to its augmented mobility (Lakowicz 1999).

Judged from these data on W89m, the movements of Trp89 became more restricted in <=20% iPrOH, whereas the opposite is true above this iPrOH concentration. The same overall tendency, yet less pronounced, is observed for the wild-type TLL. It should be noted that steady-state anisotropy measurements as such do not allow one to distinguish between local segmental motions of Trp from changes in the rate of Brownian tumbling of the protein in solution. Accordingly, the increase in r may also report oligomerization of TLL, further supported by the time-resolved fluorescence anisotropy measurements.

More specifically, Trp fluorescence anisotropy decays reflect rapid local motions of the Trp side chains as well as the overall rotational diffusion of the entire protein (Steiner 1991). Two rotational correlation times were measured for W89m. The shorter correlation time {phi}1 can be assigned to the local segmental motions of Trp89 (Davidson et al. 1999). Movement of Trp89 became faster when [iPrOH] approaches 20%, with {phi}1 reaching a minimum of 0.08 ns in 25% iPrOH (Fig 10AGo). Residual anisotropy r{infty} has been interpreted as resulting from an energy barrier that prevents rotational diffusion of the fluorophore beyond a certain angle. Therefore r{infty} does not decay to zero when the angular range of the rotational motion of the fluorophore is restricted within the time regime of the fluorophore lifetime (Lakowicz 1999). The values for r{infty} decreased steeply when [iPrOH] exceeded 10%, indicating less restriction for the motion of Trp89 (Fig 10BGo), i.e., diminished constraints for its mobility. These data would be compatible with the location of Trp89 on the protein surface in the open conformation.

The longer correlation time {phi}2 reflects the motions of the entire lipase molecule, relating to the hydrodynamic volume by the Einstein-Stoke's equation


(1)

where {eta} is the viscosity of the medium at temperature T, M is the molecular mass of protein, v is the specific volume of protein, h is the hydration factor, and R is the gas constant. Our results show {phi}2 and thus also the hydrodynamic volume of the lipase to increase with increasing [iPrOH]. Moreover, values of {phi}2 for W89m increased in a stepwise manner and in multiples of the original value upon increasing [iPrOH] between 5% and 25% iPrOH. The value of {phi}2 in the absence of iPrOH, 20 ns, is consistent with that expected for a spherical 30 kD protein with a typical hydration of 0.2 g of H2O/g (Lakowicz 1983). Using a value of 0.89 cp for the viscosity of water at 25°C, this {phi}2 corresponds to a hydrodynamic radius of 27.8 Å. This radius is in excellent agreement with further value the radius of 28 Å (= the maximum distance from the center of mass) obtained from crystal structure (Swaminathan et al. 1996), and thus confirms W89m to be a monomer in the absence of iPrOH. Accordingly, it is possible that the stepwise increments in {phi}2 reflect the formation of dimers and trimers by W89m in the presence of iPrOH. Notably, steady-state anisotropies for wild-type and W89m TLL increased abruptly between 5% and 10% iPrOH. In the light of the {phi}1 values, the frequency of the movement of the lid did not change in this concentration range. Therefore, the decrease in the rotation of Trp89 should result from a decreased tumbling of the entire TLL molecule in solution, in keeping with protein oligomerization. TLL can be aggregated by an electrostatic attraction in crystals in the presence of the detergent pentaoxyethylene octylether (C8E5) and trioxyethylene decylether (C10E3) (A.M. Brzozowski, pers. comm.).

The reason that iPrOH can induce the aggregation of TLL is likely to be a partially open structure of lipase when hydrophobic interactions are attenuated. Aggregates are thus likely to form when the structure is partially opened so that the exposed hydrophobic regions from different molecules interact (Swaminathan et al. 1996). Another possibility is that the organic solvent promotes the formation of intermolecular hydrogen bonds.

Above 20% iPrOH however, the Trp89 rotates at a high frequency, evidenced by a decrease in {phi}1, while the oligomeric state of the protein is maintained, as evidenced by {phi}2. We interpret the decrease in steady-state anisotropy at iPrOH>15% to result from conformational changes accompanying the formation of higher oligomers. Changes in the conformational dynamics induced by iPrOH are further reflected in the measurements of the catalytic activity of the wild-type and W89m TLL as a function of [iPrOH]. For W89m, the catalytic activity towards the water-soluble substrate carboxy-DCFDA increased up to 15% iPrOH, whereas exceeding this concentration caused an inhibition. Activation of TLL by iPrOH is in accordance with the opening of the lid. Inhibition of the hydrolytic activity at iPrOH concentrations above 20%–30 % could result from a number of reasons. Accordingly, iPrOH could occupy the active site after the opening of the lid, thus impeding the access of the substrate as well as possibly also the mandatory water to the catalytic residues. Increasing the concentration of substrate should offset this effect, as observed (Fig. 1Go). Another mechanism is diminished water activity in general.

Conformational changes induced by high [iPrOH] leading to an unfavorable active site geometry cannot be excluded. The above mechanism is in alignment with the strongly correlating changes in steady-state anisotropy (Fig. 6Go) and catalytic activity (Fig. 1Go) for both the wild-type and W89m TLL.

This correlation is very clear when the relative activities for the lipases are illustrated as a function of r (Fig. 11Go). Accordingly, on increasing [iPrOH] up to 15% and 10% respectively, a change in the conformation of lipase caused the steady-state anisotropy r to increase for both the wild-type and W89m TLL. This change in protein conformation also resulted in an ~2- to 6-fold enhancement in the catalytic activity and is likely to represent the opening of the lid. On increasing [iPrOH] up to 50%, a further change in the conformational dynamics occurs, accompanied by a decrease in r and inhibition of the catalytic activity. The latter would be compatible with formation of higher oligomers by TLL in the open conformation at high concentrations of iPrOH, perhaps involving the burying of the active site in the protein–protein contact site.



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  		Fig. 11. Relative catalytic activities of the wild-type (A) and W89m (B) TLL as a function of steady-state anisotropy r. The concentration of the fluorogenic substrate was either 50 µM (solid circle) or 100 µM (open square). Data were taken from Figures 3 and 11GoGo. Arrows indicate the direction of increasing iPrOH.

 

   Conclusion
TOP
 Abstract
 Introduction
 Results
 Discussion
 Conclusion
Materials and methods
 References
 
The data reported here demonstrate significant changes in the conformational dynamics of TLL to be induced by iPrOH. Although the information obtained about the global changes in the lipase is limited, an interesting view is gained into the dynamics of the lid, as follows. In the presence of iPrOH, Trp89 became accommodated in a more hydrophilic microenvironment, fluorescence decay of W89m became two-exponential, and the fractional intensity of the shorter lifetime component {tau}1 increased. Also the motions of Trp89 became faster and the amplitude of these motions increased. All the above changes are compatible with iPrOH promoting the closed {Leftrightarrow} open conformational equilibrium of TLL towards the latter. The opening of the lid by iPrOH can be attributed to the reduction of hydrophobic interactions by two mechanisms. First, iPrOH disrupts water structure so as to diminish the penalty of contacts of hydrophobic surfaces with the aqueous environment, offering alternative solvation of hydrophobic groups by the nonpolar portions of the organic solvent (Rosell et al. 1995). As the contact between the lid and the rest of the lipase in the closed conformation is mainly hydrophobic in nature (Svendsen et al. 1997), the lid opens due to attenuated hydrophobic interactions. Second, the addition of iPrOH also reduces the dielectric constant of the aqueous phase. Theoretical considerations (Norin et al. 1993; Jääskeläinen et al. 1999) have indicated the latter change to promote the open conformation of the lid. Although iPrOH reduces the hydrophobic interactions between the lid and active site containing cleft, it does not disturb the secondary structure of TLL. Finally, the derived longer rotational correlation time {phi}2 revealed TLL to be a monomer in aqueous solution. Increasing concentrations of iPrOH caused stepwise increases in {phi}2, compatible with the formation of dimers and trimers, and possibly also higher oligomers.


   Materials and methods
TOP
 Abstract
 Introduction
 Results
 Discussion
 Conclusion
 Materials and methods
References
 
Materials
The wild-type and W89m (W117F, W221H, W260H) TLL were obtained from Novo Nordisk (Bagsværd, Denmark). Their concentrations were determined as described previously (Bradford 1976). Activities of wild-type TLL and W89m mutant were 2158 and 1130 µmoles/mg/min, respectively (Svendsen et al. 1997). Analytical grade iPrOH was purchased from BDH Chemical Ltd. (Toronto, Canada). In all experiments the buffer was 20 mM Hepes, 0.1 mM EDTA at pH 7.0, made in water deionized in a Milli RO/Milli Q (Millipore, Bedford, MA, USA) filtering system.

Assay for catalytic activity
TLL activity measurements were carried out with a Perkin Elmer LS 50B spectrofluorometer. The excitation and emission wavelengths were 511 and 526 nm, respectively, and both excitation and emission bandpasses were set at 5 nm. Carboxy-DCFDA (Molecular Probes, Inc., OR, USA) was dissolved in DMSO (Merck, Darmstadt, Germany) to 10 mM concentration and stored at -70°C. For assay, the concentration of carboxy-DCFDA was either 0.05 or 0.1 mM in iPrOH, as indicated. The reactions were started by the addition of TLL to a concentration of 1 µM. On the hydrolysis of carboxy-DCFDA, an intensively fluorescent product is formed (Thomas et al. 1979). The increase in fluorescence intensity with time was recorded and the slopes of the kinetic curves were determined so as to obtain the changes in relative activity. These data were corrected for the hydrolysis of the substrate in the absence of enzyme as well as for the progressive enhancement in the quantum yield of the fluorescent product observed at increasing concentrations of iPrOH. All experiments were carried out at 25°C.

CD measurements
CD spectra were collected in an N2 atmosphere with a CD spectrophotometer (Olis RSF 1000F, On-line Instrument Systems Inc., Bogart, GA, USA). Far-UV CD spectra in the 200–250 nm region were recorded with 1.0 mm path-length cells containing 0.2 mg/mL (corresponding to 6.7 µM) of either the wild-type or W89m TLL at different concentrations of iPrOH. Near-UV CD spectra (240–340 nm) were obtained with a 10 mm cuvette containing 33.3 µM lipase. Final spectra, representing the average of at least three tracings, were corrected for the background. Protein {alpha}-helical content was analyzed from the far-UV CD spectra using a computer program SELCON3 (Sreerama et al. 1999) for secondary structure analysis. Value presented for helicity is a sum of values for regular and distorted {alpha}-helix from calculations with a 29 protein reference set.

Steady-state fluorescence measurements
Steady-state fluorescence measurements were carried out with a Perkin Elmer LS 50B spectrofluorometer equipped with a magnetically stirred, thermostated cuvette compartment. The excitation wavelength for Trp was 295 nm (Das and Mazumdar 1995; Jennifer and Kathleen 1997) with both excitation and emission bandpasses set at 5 nm. When recording synchronous fluorescence spectra the difference {Delta}{lambda} between emission and excitation wavelength was set at 60 nm. In steady-state anisotropy measurements, excitation and emission bandpasses were 10 nm, and the emission wavelengths were 342 nm and 345 nm for the wild-type and W89m TLL, respectively.

Time-resolved fluorescence measurements
Commercial laserspectrometer (Photon Technology International, Ontario, Canada) was used to measure fluorescence lifetimes, rotational correlation times, and residual anisotropies. A train of 500 ps excitation pulses at 337 nm at a repetition rate of 10 Hz was produced by a nitrogen laser, pumping a dye (rhodamine 6G, Merck, Darmstadt, Germany, 5 mM solution in methanol) laser. Pulses from the dye laser at 596 nm were frequency doubled.

Excitation wavelength was 298 nm, and emission wavelengths were 342 nm and 345 nm for the wild-type and W89m TLL, respectively.

For the determination of the fluorescence lifetimes, the averages of five emission decay curves were analyzed using the software provided by the instrument manufacturer. Instrument response functions were measured separately using aqueous glycogen solution. The validity of the fit was judged by the value of the reduced chi-square, {chi}2 (Gratton et al. 1984; Jameson and Hazlett 1991), which varied in the range of 0.9–1.2. The minimum lifetime accessible to the instrument is 200 ps.

Fractional intensities I(t) were calculated according to the equation:


(2)

where {alpha}i is amplitude and {tau}i is lifetime. For polarized light, the decay of fluorescence intensity, F(t) was calculated from the raw data according to:


(3)

where I{bot}(t) is the intensity of light detected with both the excitation and emission polarizer being vertical (i.e., parallel polarizers) and I{bot}(t) is the intensity of light detected with vertical excitation polarizer and horizontal emission polarizer (i.e., perpendicular polarizers), and G is the correction term for the relative throughput of the respective polarized light component through the emission optics.

Time resolved anisotropy r(t) is defined as:


(4)

where G, I{bot} and I{bot} are as above.

The fluorescence and anisotropy decays are described by a sum of exponentials, as follows:


(5)


(6)

In the above equations, Ai and ri are the normalized preexponential initial fluorescence intensities and initial anisotropies, respectively, and {tau}i and {phi}i are the corresponding fluorescence lifetimes and rotational correlation times. When the angular range of the rotational motion of fluorophore is limited,


(7)

where r0 stands for the anisotropy in the absence of rotational diffusion. Equation 7 was used only to fit the data so as to obtain the value of the residual anisotropy.

Rotational correlation times ({phi}i) and residual anisotropies (r{infty}) were derived from the fitted curves. The decays I(t)|| and I(t){bot} were fitted before calculating the respective r(t) decays from these data. Accordingly, r(t) contains no experimental noise, and, therefore criteria such as the {chi}2 value do not apply. For the determination of rotational correlation times and residual anisotropies, each emission decay was measured 10 times.

Stopped-flow fluorescence measurements
Fluorescence changes in the millisecond time regime were monitored using a stopped-flow spectrofluorometer (Olis RSF 1000F, On-line Instrument Systems Inc.) equipped with a quartz glass fluorescence observation chamber (20 mm pathlength, 1.5 mm diameter) and a rapid scanning emission monochromator. The dead time of the instrument is ~2 min. When changes in lipase fluorescence due to mixing with iPrOH were measured, one of the syringes was filled with 2 µM lipase in buffer solution and the other syringe with 50% iPrOH. The final concentrations of the reactants in the observation chamber were thus 1 µM protein and 25% iPrOH. Excitation wavelength was 295 nm and fluorescence emission spectra were recorded in the range of 285 to 425 nm. In the analysis of the data, only emission at >310 nm was used to exclude interference from the excitation. Temperature in the reaction chamber and in the syringes was regulated by a computer-controlled circulating waterbath (Neslab, Portsmouth, NH, USA). Fluorescence intensities were detected using scanning rate of 1000 spectra/sec, with a data collection time of 0.16 sec. Each experiment was repeated at least three times and averaged values were used for further analysis.


   Acknowledgments
 
We thank Drs. Allan Svendsen and Shamkant Anant Patkar (Novo Nordisk, Bagsværd, Denmark) for providing the wild-type and W89m lipase and Drs. Marek Brzozowski (York University, York, UK) and Sanne Glad (Novo Nordisk) for their helpful comments. This study was supported by Biocentrum Helsinki, Finnish Medical Research Council, and the EU Structural Biology Programme (BIO4 CT972365).

The publication costs of this article were defrayed in part by payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 USC section 1734 solely to indicate this fact.


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