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Marasmius scorodonius extracellular dimeric peroxidase — Exploring its temperature and pressure stability
Biochimica et Biophysica Acta 1794 (2009) 1091–1098
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Biochimica et Biophysica Acta j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / b b a p a p
Marasmius scorodonius extracellular dimeric peroxidase — Exploring its temperature and pressure stability Matthias Pühse a, Renate T. Szweda b, Yingying Ma b, Christoph Jeworrek a, Roland Winter a, Holger Zorn b,⁎ a b
Physical Chemistry I – Biophysical Chemistry, Dortmund University of Technology, D-44227 Dortmund, Germany Institute of Food Chemsitry and Food Biotechnology, Justus Liebig University Giessen, D-35392 Giessen, Germany
a r t i c l e
i n f o
Article history: Received 19 December 2008 Received in revised form 13 March 2009 Accepted 16 March 2009 Available online 2 April 2009 Keywords: Basidiomycete Peroxidase High pressure FTIR SAXS
a b s t r a c t The temperature and pressure dependent stability and function of MsP1, an uncommon peroxidase from the basidiomycetous fungus Marasmius scorodonius were investigated. To this end, a series of biophysical techniques (DSC, fluorescence and FTIR spectroscopy, small-angle X-ray scattering) were combined with enzymatic studies of the enzyme. The dimeric MsP1 turned out to be not only rather thermostable, but also highly resistant to pressure, i.e., up to temperatures of about 65 °C and pressures as high as 8–10 kbar at ambient temperatures. Remarkably, the activity of MsP1 increased by a factor of two until ∼ 500 bar. At about 2 kbar, the enzymatic activity was still as high as under ambient pressure conditions. As revealed by the fluorescence and SAXS data, the increased activity of MsP1 at pressures around 500 bar may result from slight structural changes, which might stabilize the transition state of the enzymatic reaction. Owing to this marked high pressure stability of MsP1, it may represent a valuable tool for industrial high pressure applications. © 2009 Elsevier B.V. All rights reserved.
1. Introduction Growing on lignocellulosic biomasses, the basidiomycetous fungus Marasmius scorodonius (“garlic mushroom”) secretes the uncommon peroxidase MsP1 (accession number B0BK71). M. scorodonius (strain No.: CBS 137.83) is a small edible fungus, which typically forms fruiting bodies on grass, bark, twigs, and needle duff [1]. Due to its intense garlic-like flavor, it is appreciated as a spice in human nutrition. In a previous study, MsP1 has been purified from culture supernatants and cloned from a cDNA library. Size exclusion chromatography and SDS-PAGE analyses suggested a dimeric structure of native MsP1 [2]. Based on the observed amino acid sequence homology of about 48% to the enzymes DyP (Q8WZK8) and TAP (Q8NKF3) from Thanatephorus cucumeris and Termitomyces albuminosus, respectively, MsP1 was assigned to the group of so-called DyP-type peroxidases [3]. When cultured submerged in standard nutrition solution, the expression of MsP1 was significantly induced by the addition of lignin [4]. Representing an essential part of the fungus' secretome, it may contribute to the modification of lignified biopolymers [5]. It thus could become an interesting tool for the production of second generation biofuels or lignin based polymers in biorefinery approaches. Based on the enzyme's capability to degrade carotenes and xanthophylls [2], further potential industrial applications com-
prise the bleaching of food products, as well as the formation of norisoprenoid flavor compounds [6]. A high process stability of the biocatalyst is an essential prerequisite for the implementation of new biotechnological processes. Hence, the intention of the present study was to investigate the temperature and pressure stability of MsP1, as well as to prove the enzyme's uncommon dimeric occurrence in the culture supernatant. To this end, a series of biophysical techniques (DSC, fluorescence and FTIR spectroscopy, small-angle X-ray scattering) was combined with functional studies of the enzyme. In fact, interest in pressure as an additional thermodynamic and kinetic variable in physico-chemical studies of biological materials has been growing also in recent years [7–11]. The behavior of all systems under high pressure is governed by Le Châtelier's principle, which predicts that the application of pressure shifts the equilibrium towards the state that occupies a smaller volume, and accelerates processes for which the transition state has a smaller volume than the ground state. Besides the general physicochemical interest in using high pressure as a tool for understanding the structure, energetics and kinetic processes of biomolecules, high pressure is also of increasing biotechnological interest (e.g., for high pressure food processing, baroenzymology) [10–13]. 2. Materials and methods 2.1. General
⁎ Corresponding author. Tel.:+49 641 99 34 900; fax: +49 641 99 34 909. E-mail address: [email protected] (H. Zorn). 1570-9639/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.bbapap.2009.03.015
The M. scorodonius strain (CBS 137.86) was obtained from the Dutch “Centraalbureau voor Schimmelcultures”, Baarn. Enzyme production and purification were performed as described previously [2].
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2.2. FT-IR spectroscopy 1 mL of the purified enzyme solution in 50 mM aqueous phosphate buffer (pH 6.0) or 30 mM Bis–Tris-Buffer (pH 5.8; c = 9.6 mg/mL), respectively, was lyophilized overnight, and the resulting powder was suspended in 200 μL D2O. The final protein concentration (2.1% (w/v) and 4.4% (w/v), respectively) was determined by UV-spectroscopy at λ = 280 nm. Due to their different temperature and pressure sensitivities [14], phosphate buffer was used for the analysis of the temperature dependency, and Bis–Tris buffer for the high pressure measurements. For the temperature dependent experiments, a Nicolet 5700 FT-IR spectrometer (Thermo Fisher Scientific, Waltham, USA), equipped with a liquid nitrogen cooled MCT detector (HgCdTe), was used. Approximately 20 μL of the sample were filled between CaF2 windows, which were separated by a 50 μm Mylar spacer. The sample cell was placed in a thermostatted jacket, and an external water bath was used for temperature control (accuracy: ± 0.2 °C). During the measurements, the sample chamber was purged constantly with dry air. Spectral recording was performed between 4000 and 1100 cm− 1. To improve the signal to noise ratio, 256 spectra were summed up for each measuring point (resolution: 2 cm− 1). Apodization was done with a Happ–Genzel function. Data acquisition was performed between 10 and 80 °C with 5 °C increments; the equilibration time before each measurement was 15 min. The pressure dependent measurements were conducted at 30 and 60 °C with a Nicolet MAGNA 550 FT-IR spectrometer. Pressure generation was achieved with a thermostatted High Pressure Diamond Optics P-series diamond anvil cell with type IIa diamonds (High Pressure Diamond Optics Inc., Tucson, USA). A 50 μm thick gasket of stainless steel with a 0.45 mm drilling was placed between the two diamonds holding ∼10 nL of the sample. Pressure was determined with co-added BaSO4, which shows the characteristic pressure sensitive symmetric sulfate stretching mode around 983 cm− 1 that increases linearly with pressure (accuracy: ± 300 bar [15]). All other technical parameters were identical to the temperature dependent measurements. Spectral processing and deconvolution of the amide-I′-band was done with the GRAMS/AI 8.0 software package (Thermo Fisher Scientific, Waltham, USA). After subtraction of noise and the temperature-corresponding buffer background (D2O), all spectra were normalized to the same area to ensure comparability. Finally, the amide-I′-region was fitted with the number of subbands (which are typical for particular secondary structure elements) detected by the 2nd derivative using a Voigt-function. Peak assignment to the different secondary structures was done according to the literature [16]. As the transition dipole moments of the different conformers might be slightly different, we focus essentially on the relative changes of the peak areas as a function of temperature and pressure.
buffer mixture (Bis–Tris; pH 5.8) was dissolved in 100 μL H2O, yielding a 1.9% (w/v) solution. 20 μL of the sample were placed in a high pressure X-ray sample cell. The cell was equipped with KaptonÒ polymer windows, and the sample detector distance was 0.905 m. The plots of the diffraction intensity versus reciprocal spacings (Q, Q = 4πsinΘ / λ; 2Θ is the scattering angle and λ the wavelength of the radiation) were recorded using MATLAB™ based software, written by the ESRF staff. The pressure dependent measurements were performed at 30 °C. The maximum pressure applied was 3000 bar (accuracy: ± 20 bar). Measurements were performed at steps of 100 bar in the upward and downward direction. Additional measurements under the same conditions were taken with a pure buffer sample for background correction. Temperature dependent SAXS measurements were conducted with a Kratky-camera (Model KKK, Anton Paar, Graz, Austria). The X-ray energy was 8.05 keV (l = 0.154 Å). After dissolving the lyophilized protein-buffer mixture (phosphate buffer; pH 6.0) in 100 μL water, yielding a 1.5% (w/v) protein solution, measurements were executed at 30, 60, and 70 °C. Temperature control was achieved via an external water bath (DT = ± 1.5 °C). 2.5. Fluorescence spectroscopy Temperature dependent fluorescence spectroscopic measurements were performed with a Perkin Elmer LS 55 fluorescence spectrometer (PerkinElmer Inc., Waltham, USA), whereas the pressure dependent measurements were performed on a K2 multifrequency phase and modulation fluorometer (ISS Inc., Champaign, USA). For the temperature dependent measurements, the sample (∼300 mL of a 0.5 mg/mL enzyme solution in 50 mM phosphate buffer (pH 6.0)) was filled in a 3 mm cuvette equipped with quartz windows. Temperature control was achieved via an external water bath. Spectra were recorded in 5 °C steps, and the temperature-difference between bath and cuvette was measured and corrected afterwards. Before each measurement, the sample was allowed to equilibrate for 5 min. The tryptophan residues were excited at 295 nm, and the resulting emission was recorded from 307 to 400 nm. For the pressure dependent measurements at 30 °C, a custom made high pressure cell, consisting of a beryllium–copper alloy, with ethanol as pressurizing medium was used. Here, 1 mL of a 1.3 mg/mL solution in 30 mM Bis–Tris buffer (pH 5.8) was filled in a pressure-resistant quartz flask, tightly sealed with heat and solvent resistant laboratory film. Pressure was generated via an external manometer (accuracy: ± 10 bar). Spectra were recorded every 100 bar up to 2200 bar, with an equilibration time of 5 min. After subtraction of buffer background, spectral changes were monitored by calculating the center of spectral mass [17]. 2.6. Enzyme assays
2.3. Differential scanning calorimetry (DSC) For the DSC-measurements, a 0.19% (w/v) enzyme solution in 50 mM phosphate buffer (pH 6.0) was used. Measurements were conducted in duplicate with a VP DSC calorimeter (MicroCal, Northampton, USA). The sample cell of the calorimeter was filled with 0.5 mL of the enzyme solution, whereas the reference cell was filled with the protein-free buffer. The differential heat flow was recorded between 10 and 100 °C with a heating rate of 40 °C/h. Data analysis was performed with a customized version of ORIGIN 7 (MicroCal). 2.4. Synchrotron small-angle X-ray scattering (SAXS) The SAXS experiments were performed at the ID2 high brilliance beamline at the European Synchrotron Radiation Facility (ESRF) in Grenoble, France. The medium X-ray energy was 12.46 keV, corresponding to a wavelength of 0.995 Å. The lyophilized protein-
2.6.1. Preparation of β-carotene solution 5 mg of β-carotene (Sigma-Aldrich, Germany) and 0.5 g of Tween 80 were dissolved in 20 mL dichloromethane. The solvent was evaporated under vacuum, and the residue was suspended immediately in 30 mL water. The suspension was filtered through a 0.22 μm syringe filter and filled up to 50 mL with distilled water. This solution was prepared freshly before each measurement. Temperature dependent enzymatic measurements were performed on a PerkinElmer Lambda 25 UV/VIS spectral photometer (PerkinElmer Inc., Waltham, USA), which was temperature-controlled by an external water bath (DT = ±1.5 °C). The activity was quantified by the ABTS-assay [18]. The reaction mixture contained 700 μL 50 mM sodium acetate buffer (pH 3.5), 100 μL 5 mM 2,2-azinobis-(3-ethyl benzthiazoline-6-sulphonate) (ABTS), 100 μL MsP1-solution (10 mUABTS, 30 °C, pH 5.8) and 100 μL 20 mM H2O2. The increasing absorbance was monitored at l = 420 nm from 30 to 70 °C in 5 °C steps
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over 360 s. One unit of enzyme activity was defined as the amount of enzyme oxidizing 1 μmol of substrate per minute. The high pressure measurements were performed on a K2 multifrequency phase and modulation fluorometer, which was adjusted to collect absorption spectra. All other technical details were identical to the high pressure fluorescence measurements (see above). For the ABTS-assay, 1.06 mL of the reaction mixture (930 μL 33 mM Bis–Tris buffer, pH 5.8, 50 μL 5 mM ABTS, 30 μL MsP1-solution (3 mUABTS, 30 °C, pH 5.8) and 50 μL 20 mM H2O2) were placed in a pressure-resistant and tightly sealed quartz flask. For every pressure step, the increase in absorbance was measured over 600 s at l = 420 nm and 30 °C in a pressure range from 1 to 2500 bar. The β-carotene assay was performed similarly with 1.06 mL reaction mixture (820 μL 33 mM Bis–Tris buffer, pH 5.8, 150 μL β-carotene solution, 50 μL MsP1-solution (0.5 mUβ-carotene, 30 °C, pH 5.8) and 40 μL 20 mM H2O2). For every pressure step, the decrease in
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absorbance was measured at l = 450 nm and 30 °C over 1000 s from 1 bar to 2500 bar. 2.7. Protein quantification The protein concentration was calculated using an absorption coefficient of MsP1 of 27,960 M− 1 cm− 1 at 280 nm (ProtParam, [19]). 3. Results 3.1. Temperature and pressure dependent FT-IR experiments To monitor the conformational changes of the protein, the amide-I′-band at ∼ 1700–1600 cm− 1 was analyzed. The relative area of each subband corresponds in good approximation to the fraction of the respective secondary structure [20–23]. A representative example
Fig. 1. (a) Deconvoluted and fitted amide-I′-FT-IR spectrum of MSP-1 at 10 °C. (b) Second derivative of the amide-I′-FT-IR spectrum of MSP-1 at 10 °C. Negative peaks correspond to the loci of chosen FT-IR subbands.
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Fig. 2. Temperature dependence of the relative infrared peak areas of the subbands of MSP-1, which are assigned to the following secondary structures: ▴ α-helices, ● loops and turns, □ intramolecular β-sheets, and Δ intermolecular β-sheets (commonly associated with aggregates).
of the deconvoluted amide-I′-band after peak fitting is depicted in Fig. 1. Four main subbands were deduced from the second derivative spectra, which appeared at ∼1672, ∼1657, ∼ 1642, and ∼1630 cm− 1. These subbands correspond to loops and turns, α-helices, random coil structures, and intramolecular β-sheets, respectively. The positions of these subbands do not change significantly upon heating. As depicted in Fig. 2, MsP1 reveals to be highly α-helical, with an approximately equal fraction of unordered structures. Also, minor contributions of intramolecular β-sheets and turns are present. With increasing temperature, no apparent change in the band areas are detected until ∼65 °C, where the α-helical content starts to decrease and the unordered fraction increases concomitantly (Fig. 2). Simultaneously, a new β-sheet fraction at ∼1617 cm− 1 appears at ∼ 70 °C, which can be ascribed to formation of intermolecular β-sheets in aggregated structures. This behavior is in good agreement with the temperature dependence of the enzyme activity, indicating an abrupt loss of enzyme activity above about 60 °C (see below). With increasing hydrostatic pressure at 60 °C, the subbands retain their position, and no significant change of the enzyme's secondary structure is observed (Fig. 3), indicating a high pressure stability of MsP1. Above ∼ 8 kbar, only a minor increase in unordered structures at
▪ random coil structure,
the expense of turns and α-helical conformations is observed. Similar results were obtained at 30 °C (data not shown). A small percentage of aggregates were present from the beginning, which may be ascribed to the high protein concentration necessary for the FTIR spectroscopic measurements in the diamond anvil cell. 3.2. Differential scanning calorimetry Up to ∼60 °C, the differential heat capacity Cp (with respect to pure buffer solution) increases almost linearly with temperature, peaks at ∼63 °C, and decreases abruptly above ∼78 °C (data now shown), where, according to the FTIR data, thermal aggregation of the protein takes place. In fact, protein aggregation is generally an exothermic process [24]. 3.3. Temperature and pressure dependent fluorescence spectroscopy The MSP-1 dimer contains 8 tryptophane residues (i.e., there are 4 Trp residues per monomer), which are located in different parts of the three-dimensional structure. The fluorescence spectrum of the protein thus gives an average over all contributing intrinsic
Fig. 3. Pressure dependence of the secondary structure elements of MSP-1 as revealed from the FTIR spectra at 60 °C. For symbol denotation, see Fig. 2.
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fluorophores. Generally, a red-shift of this band indicates a more polar environment of the fluorescent tryptophane residues, which occurs upon protein unfolding and denaturation [17]. From 15 to 50 °C, the center of spectral mass (CSM) of the Trp-emission exhibits a linear blue-shift, which can be ascribed to an essentially hydrophobic environment, indicating that the protein is still in a compact state. Beyond 50 °C, a pronounced red-shift up to 70 °C is observed, where a change in slope of the temperature dependence of the emission wavelength, l (T), and a further red-shift is detected (Fig. 4). The redshift above 50 °C can be explained by a temperature-induced conformational change with subsequent increasing exposure of tryptophane residues. The midpoint of the transition occurs at around 63 °C. By comparing the final CSM-value of ∼357 nm at 85 °C with the one of fully hydrated tryptophane (∼ 362 nm, [25]), it can be concluded, that most Trp residues are largely, but not fully, unshielded at this temperature. The change in slope at ∼ 68 °C may be ascribed to the end of the thermally induced dimer dissociation process that has also been detected in the SAXS data described below, which is then followed by the unfolding reaction of the protein. This interpretation is in good agreement with the FTIR- and DSC-data. Notably, the corresponding loss of secondary structure up to 80 °C, as derived from the infrared spectroscopic data, is rather small (Fig. 2). At 30 °C, the pressure dependent CSM-values exhibit a small and almost linear red-shift with increasing pressure (Fig. 5). A discontinuity with a subsequent change in slope is observed between 500 and 600 bar, however, which might be due to a minor structural change, as corroborated by the SAXS measurements (see below). Surprisingly, the enzyme activity under high hydrostatic pressure conditions shows a maximum at about 500 bar as well at the same temperature (see below). Hence it is very likely that this minor structural change is responsible for the observed change in enzyme activity. The increasing red-shift at pressures above 700 bar may be ascribed to the intrinsic pressure effect on the tryptophane residue [26] and, possibly, increasing pressure-induced fluctuations of the protein structure, which generally increase upon pressurization due to a weakening of hydrophobic interactions [7–10]. 3.4. Small-angle X-ray scattering (SAXS) To reveal details of these pressure-induced small conformational changes of the protein in the lower pressure regime, SAXS measurements were performed as a function of pressure up to 3 kbar. Fig. 6 shows the small-angle X-ray scattering intensity as a function of momentum transfer, I(Q), for pressures from 1 bar to 3 kbar at T = 30 °C. The shape of the SAXS curve with a breakpoint in the slope
Fig. 4. Temperature dependence of the spectral center of mass (bλN) of the Trpfluorescence of MSP-1 from 15 to 86 °C.
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Fig. 5. Pressure dependence of the spectral center of mass (bλN) of the Trp-fluorescence of MSP-1 at 30 °C.
of I(Q) at about 1.2 nm− 1 is typical for dimeric protein structures [27]. Interestingly, upon pressurization, changes occur only in the intermediate momentum transfer region from 1.25 to 3 nm− 1, indicating only minor medium-range-order changes of the protein conformation in this pressure range. Besides, with increasing pressure, a decrease in scattering intensity is observed, only, which is due to a decrease of the electron density contrast between the protein and the surrounding water upon pressurization. The radius of gyration, Rg, was determined from the Guinier-plots (lnI versus Q2) in the low-Q regime as well as from the pair-distance distribution function [21]. Within the accuracy of the measurements, a constant value of Rg of 2. 41 ± 0.05 nm was found up to pressures of 3 kbar (data not shown). For further data analysis, the measured I(Q) data were subjected to indirect Fourier transformation using the program GNOM, which calculates the distance distribution function p(r) for monodisperse systems from the one-dimensional scattering curves (Fig. 7). Interestingly, p(r) exhibits two more or less broad maxima with a minimum around 1.5 nm, again indicating the presence of a dimeric structure. The first maximum at an r-value of about 0.5 nm represents distances with high electron densities, which are basically not changing with pressure. With increasing pressure, a minimum is emerging at intermediate distances of approx. 3 nm, however, which might be due to a decreasing electron density in the connection region of the two monomeric parts of the protein. The change of p(r)
Fig. 6. SAXS data of MSP-1 at selected pressures and T = 30 °C.
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Fig. 7. Pair-distance distribution function of MSP-1 at selected pressures and T = 30 °C.
between 2 and 5 nm thus originates from a pressure-induced small tertiary structural change in the pressure range starting below 1 kbar. After all, no dissociation and no significant unfolding or major changes in tertiary structure can be deduced from the analysis of the pressure dependent SAXS measurements up to 3 kbar. The Guinier-plots of the temperature dependent measurements at ambient pressure exhibit an Rg value of ∼2.4 nm at 30 and 60 °C. At 70 °C, this value first drops to ∼1.2 nm and subsequently increases drastically due to aggregation of the protein (data not shown as the Rg of such a large polydisperse aggregate cannot be determined). This clearly indicates a temperature-induced dissociation of the MSP-1 dimer into monomers, which manifests itself in a decrease of the enzyme activity as observed above ∼60 °C (see below), as well as in the aggregation of the protein as revealed by the DSC and FTIR spectroscopic data. Hence, the catalytic activity of the enzyme seems to be restricted to the integrity of the dimeric structure. 3.5. Temperature and pressure dependence of MsP1 activity Maximum activity of MsP1 was detected between 55 and 60 °C (Fig. 8). The decrease of activity above 60 °C is in agreement with the FTIR spectroscopy data, which indicate conformational changes beyond ∼65 °C. When the enzyme was incubated at 50 °C for 24 h,
the loss of enzyme activity amounted to less than 10%, only (data not shown). The catalytic properties of MsP1 were further characterized by steady-state kinetic analyses at 27 °C and a constant H2O2 concentration of 20 mM. Under these experimental conditions, Km and kcat values of 60 μM and 0.8 s− 1, respectively, were determined. For evaluation of the pressure stability of MsP1, two assays were employed: ABTS as a common peroxidase substrate, and additionally β-carotene, representing a potential industrial application of MsP1 [2]. A greenish blue radical cation is formed from ABTS, and βcarotene is degraded to norisoprenoids by peroxidase activity in the presence of hydrogen peroxide. The absorbance of the ABTS solution slightly depended on the pressure, which was considered in the calculation of the enzyme activity. No changes of the extinction coefficient were observed with β-carotene with increasing pressure (data not shown). As depicted in Fig. 9, for the oxidation of both substrates, activity optima at about 500 bar were determined. Compared to atmospheric pressure, the enzymatic activity of MsP1 is roughly doubled at 500 bar. Beyond 800 bar, the activity steadily decreases to reach the level of atmospheric pressure for β-carotene degradation at 2300 bar, and for ABTS oxidation at about 1100 bar. Compared to the activity at atmospheric pressure, only a negligible activity loss commenced even at 2500 bar. 4. Discussion and conclusions Compared to other members of the DyP-type enzyme family, MsP1 exhibits a remarkably high temperature optimum and a good thermostability. For the peroxidase DyP from T. cucumeris (formerly denominated as Geotrichum candidum), which is capable of decolorizing various synthetic dyes, an optimal temperature of 30 °C has been described [28]. Lignolytic enzymes like manganese peroxidases, laccases, and members of the DyP family represent interesting tools for the low energy and low cost alternatives for industrial processes in which lignin modification is of importance, such as paper and pulp processing and processes for valorization of agricultural residues. The polymeric carbohydrates of agricultural residues, cellulose and hemicellulose are typically hydrolyzed by carbohydrases up to temperatures of 50 °C. More recent developments of cellulases active beyond 50 °C is of interest, since reactions are energetically more favourable at higher temperatures and they give additional benefits in processes, such as decrease of viscosity [29,30]. Thus, simultaneous delignification and saccharification at higher temperatures requires thermostable delignifying enzymes.
Fig. 8. Temperature dependence of the enzyme activity of MsP1 (oxidation of ABTS).
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Fig. 9. Pressure dependence of the enzyme activity of MsP1 for the oxidation of ABTS and β-carotene (at 30 °C).
Remarkably, the activity of MsP1 increases until ∼ 500 bar, before it starts to decrease again to about the ambient pressure activity level. Conversely, various enzymes are markedly inactivated upon high pressure treatment of about 2 kbar. Activity losses of 20 and 50% have been reported for soy lipoxygenase and horseradish peroxidase, respectively, after 15 min exposure to a hydrostatic pressure of 2 kbar at 55 °C. Total inactivation of the lipoxygenase was observed at 5.7 kbar [31]. On the other hand, some proteins are stabilized at high pressure. Up to 4 kbar were required to maintain a carboxypeptidase from the archaebacterium Sulfolobus solfataricus in its active state at higher temperatures [32], and L-galactosidases from E. coli and Aspergillus oryzae were stabilized in the range of 2–3 kbar at 50 and 65 °C, respectively [33]. The pressure level causing full dissociation of the MsP1 dimer could not be revealed in the present study. Other oligomeric enzymes such as glyceraldehyde-3-phosphate dehydrogenase, lactate dehydrogenase, and malate dehydrogenase were found to reversibly dissociate and re-associate upon application and release of hydrostatic pressure up to 2 kbar [33]. The increased activity of MsP1 at pressures around 500 bar may result from slight structural changes stabilizing the transition state of the enzymatic reaction. Generally, high pressure may influence enzyme-catalyzed reactions either by enzyme conformational changes, or by modification of the reaction mechanism [13]. In fact, high pressure has been used to enhance the activity or selectivity of a number of enzymes, paving the way for new applications. Most of the applications have been reported from the food processing industry, especially from dairy and juice production [34]. For example, the reduction of bitterness in citrus juices correlates directly with the concentration of naringin. The enzyme naringinase hydrolyses naringin to the tasteless naringenin, and the activity of the enzyme was shown to increase 3-fold under high pressure conditions (1.6 kbar) [35]. The use of microbial transglutaminase (MTG) under hydrostatic pressure of up to 6 kbar allowed for the cross-linking of proteins which could not be affected by MTG at atmospheric pressure [36]. However, at higher pressures the tertiary structure of the native enzyme was impaired, and α-helical secondary structures were unfolded [37]. No significant changes in the secondary structure of MsP1 occurred up to 10 kbar, minor changes – possibly a slight pressure-induced swelling, indicating the onset of the unfolding reaction – occur above about 8 kbar up to 10 kbar, the maximum pressure reached, only. Regarding this remarkable high pressure stability of MsP1, it may represent a valuable tool for industrial high pressure applications. Delignification of agricultural residues represents an interesting target. The common pretreatment of lignocellulosic substrates prior
to carbohydrate hydrolysis and fermentation to ethanol is performed by steam explosion [38,39], dilute acid [40,41], or an organosolv process [42]. However, these procedures come along with the formation of toxic by-products, which impede the downstream enzymatic hydrolysis and the fermentative conversion of the released sugars into ethanol [40,43–47]. An entirely enzymatic delignification process may overcome these obstacles. Acknowledgements Financial support from the Deutsche Forschungsgemeinschaft (DFG), the Deutsche Bundesstiftung Umwelt (DBU), and the Fonds der Chemischen Industrie is gratefully acknowledged. References [1] G.C. Ainsworth, F.K. Sparrow, A.S. Sussman, A taxonomic review with keys: basidiomycetes and lower fungi, in: G.C. Ainsworth, F.K. Sparrow, A.S. Sussman (Eds.), The Fungi, an Advanced Treatise, vol. 4B, Academic Press, Orlando, 1973. [2] M. Scheibner, B. Hülsdau, K. Zelena, M. Nimtz, L. de Boer, R.G. Berger, H. Zorn, Novel peroxidases of Marasmius scorodonius degrade β-carotene, Appl. Microbiol. Biotechnol. 27 (2008) 1241–1250. [3] V. Faraco, A. Piscitelli, G. Sannia, P. Giardina, Identification of a new member of the dye-decolorizing peroxidase family from Pleurotus ostreatus, World J. Microbiol. Biotechnol. 23 (2007) 889–893. [4] B. Hülsdau, Oxidativer Abbau von Carotinoiden durch Pilzenzyme, Dissertation, Gottfried Wilhelm Leibniz Universität, Hannover, 2007. [5] H. Bouws, A. Wattenberg, H. Zorn, Fungal secretomes—nature's toolbox for white biotechnology, Appl. Microbiol. Biotechnol. 80 (2008) 381–388. [6] H. Zorn, S. Langhoff, M. Scheibner, M. Nimtz, R.G. Berger, A Peroxidase from Lepista irina cleaves β-carotene to flavour compounds, Biol. Chem. 384 (2003) 1049–1056. [7] F. Meersman, L. Smeller, K. Heremans, Protein stability and dynamics in the pressure–temperature plane, Biochim. Biophys. Acta 1764 (2006) 346–354. [8] I. Daniel, P. Oger, R. Winter, Origins of life and biochemistry under high pressure conditions, Chem. Soc. Rev. 35 (2006) 858–875. [9] J.L. Silva, D. Foguel, C.A. Royer;, Pressure provides new insights into protein folding, dynamics and structure, Trends Biochem. Sci. 26 (2001) 612–618. [10] R. Winter (Ed.), High Pressure Bioscience and Biotechnology II, Springer-Verlag, Heidelberg, Germany, 2003. [11] F. Abe, Exploration of the effects of high hydrostatic pressure on microbial growth, physiology and survival: perspectives from piezophysiology, Biosci. Biotechnol. Biochem. 71 (2007) 2347–2357. [12] C. Balny, P. Masson, K. Heremans, High pressure effects on biological macromolecules: from structural changes to alteration of cellular process, Biochim. Biophys. Acta 1595 (2002) 3–10. [13] D.B. Northrop, Effects of high pressure on enzymatic activity, Biochim. Biophys. Acta 1595 (2002) 71–79. [14] R.C. Neumann, W. Kauzmann, A. Zipp, Pressure dependence of weak acid ionization in aqueous buffers, J. Phys. Chem. 77 (1973) 2687–2691. [15] P.T.T. Wong, D.J. Moffat, A new internal pressure calibrant for high-pressure infrared spectroscopy of aqueous systems, Appl. Spectr. 43 (1989) 1279–1281. [16] A. Barth, Infrared spectroscopy of proteins, Biochim. Biophys. Acta 1767 (2007) 1073–1101.
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[32] N. Bec, A. Villa, P. Tortora, V.V. Mozhaevt, C. Balny, R. Lange, Enhanced stability of carboxypeptidase from Sulfolobus solfataricus at high pressure, Biotechnol. Lett. 18 (1996) 483–488. [33] B.B. Boonyaratanakornkit, C.B. Park, D.S. Clark, Pressure effects on intra- and intermolecular interactions, Biochim. Biophys. Acta 1595 (2002) 235–249. [34] G.K. Kouassi, R.C. Anantheswaran, S.J. Knabel, J.D. Floros, Effect of highpressure processing on activity and structure of alkaline phosphatase and lactate dehydrogenase in buffer and milk, J. Agric. Food Chem. 55 (2007) 9520–9529. [35] L. Ferreira, C. Afonso, H. Vila-Real, A. Alfaia, M.H.L. Ribeiro, Evaluation of the effect of high pressure on naringin hydrolysis in grapefruit juice with naringinase immobilised in calcium alginate beads, Food Technol. Biotechnol. 46 (2008) 146–150. [36] S. Lauber, I. Krause, H. Klostermeyer, T. Henle, Microbial transglutaminase crosslinks b-casein and b-lactoglobulin to heterologous oligomers under high pressure, Eur. Food Res. Technol. 216 (2003) 15–17. [37] O. Mendez, H. Rawel, U. Schwarzenbolz, T. Henle, Structural changes of microbial transglutaminase during thermal and high-pressure treatment, J. Agric. Food Chem. 54 (2006) 1716–1721. [38] M. Galbe, G. Zacchi, Pretreatment of lignocellulosic materials for efficient bioethanol production, Adv. Biochem. Eng. Biotechnol. 108 (2007) 41–65. [39] R.P. Chandra, R. Bura, W.E. Mabee, A. Berlin, X. Pan, J.N. Saddler, Substrate pretreatment: the key to effective enzymatic hydrolysis of lignocellulosics? Adv. Biochem. Eng. Biotechnol. 108 (2007) 67–93. [40] X.B. Lu, Y.M. Zhang, J. Yang, Y. Liang, Enzymatic hydrolysis of corn stover after pretreatment with dilute sulfuric acid, Chem. Eng. Technol. 30 (2007) 938–944. [41] Y.Y. Lee, P. Iyer, R.W. Torget, Dilute-acid hydrolysis of lignocellulosic biomass, Adv. Biochem. Eng. Biotechnol. 65 (1999) 93–115. [42] F. Sun, H. Chen, Organosolv pretreatment by crude glycerol from oleochemicals industry for enzymatic hydrolysis of wheat straw, Bioresour. Technol. 99 (2008) 5474–5479. [43] M. Cantarella, L. Cantarella, A. Gallifuoco, A. Spera, F. Alfani, Effect of inhibitors released during steam-explosion treatment of poplar wood on subsequent enzymatic hydrolysis and SSF, Biotechnol. Prog. 20 (2004) 200–206. [44] J.P. Delgenes, R. Moletta, J.M. Navarro, Effects of lignocellulose degradation products on ethanol fermentations of glucose and xylose by Saccharomyces cerevisiae, Zymomonas mobilis, Pichia stipitis, and Candida shehatae, Enzyme. Microb. Technol. 19 (1996) 220–225. [45] S.I. Mussatto, M. Fernandes, A.M.F. Milagres, C. Roberto, Effect of hemicellulose and lignin on enzymatic hydrolysis of cellulose from brewer's spent grain, Enzyme. Microb. Technol. 43 (2008) 124–129. [46] E. Ruiz, C. Cara, P. Manzanares, M. Ballesteros, E. Castro, Evaluation of steam explosion pre-treatment for enzymatic hydrolysis of sunflower stalks, Enzyme, Microb. Technol. 42 (2007) 160–166. [47] S. Ando, I. Arai, K. Kiyoto, S. Hanai, Identification of aromatic monomers in steamexploded poplar and their influences on ethanol fermentation by Saccharomyces cerevisiae, J. Ferment. Technol. 64 (1989) 567–570.
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Biochimica et Biophysica Acta j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / b b a p a p
Marasmius scorodonius extracellular dimeric peroxidase — Exploring its temperature and pressure stability Matthias Pühse a, Renate T. Szweda b, Yingying Ma b, Christoph Jeworrek a, Roland Winter a, Holger Zorn b,⁎ a b
Physical Chemistry I – Biophysical Chemistry, Dortmund University of Technology, D-44227 Dortmund, Germany Institute of Food Chemsitry and Food Biotechnology, Justus Liebig University Giessen, D-35392 Giessen, Germany
a r t i c l e
i n f o
Article history: Received 19 December 2008 Received in revised form 13 March 2009 Accepted 16 March 2009 Available online 2 April 2009 Keywords: Basidiomycete Peroxidase High pressure FTIR SAXS
a b s t r a c t The temperature and pressure dependent stability and function of MsP1, an uncommon peroxidase from the basidiomycetous fungus Marasmius scorodonius were investigated. To this end, a series of biophysical techniques (DSC, fluorescence and FTIR spectroscopy, small-angle X-ray scattering) were combined with enzymatic studies of the enzyme. The dimeric MsP1 turned out to be not only rather thermostable, but also highly resistant to pressure, i.e., up to temperatures of about 65 °C and pressures as high as 8–10 kbar at ambient temperatures. Remarkably, the activity of MsP1 increased by a factor of two until ∼ 500 bar. At about 2 kbar, the enzymatic activity was still as high as under ambient pressure conditions. As revealed by the fluorescence and SAXS data, the increased activity of MsP1 at pressures around 500 bar may result from slight structural changes, which might stabilize the transition state of the enzymatic reaction. Owing to this marked high pressure stability of MsP1, it may represent a valuable tool for industrial high pressure applications. © 2009 Elsevier B.V. All rights reserved.
1. Introduction Growing on lignocellulosic biomasses, the basidiomycetous fungus Marasmius scorodonius (“garlic mushroom”) secretes the uncommon peroxidase MsP1 (accession number B0BK71). M. scorodonius (strain No.: CBS 137.83) is a small edible fungus, which typically forms fruiting bodies on grass, bark, twigs, and needle duff [1]. Due to its intense garlic-like flavor, it is appreciated as a spice in human nutrition. In a previous study, MsP1 has been purified from culture supernatants and cloned from a cDNA library. Size exclusion chromatography and SDS-PAGE analyses suggested a dimeric structure of native MsP1 [2]. Based on the observed amino acid sequence homology of about 48% to the enzymes DyP (Q8WZK8) and TAP (Q8NKF3) from Thanatephorus cucumeris and Termitomyces albuminosus, respectively, MsP1 was assigned to the group of so-called DyP-type peroxidases [3]. When cultured submerged in standard nutrition solution, the expression of MsP1 was significantly induced by the addition of lignin [4]. Representing an essential part of the fungus' secretome, it may contribute to the modification of lignified biopolymers [5]. It thus could become an interesting tool for the production of second generation biofuels or lignin based polymers in biorefinery approaches. Based on the enzyme's capability to degrade carotenes and xanthophylls [2], further potential industrial applications com-
prise the bleaching of food products, as well as the formation of norisoprenoid flavor compounds [6]. A high process stability of the biocatalyst is an essential prerequisite for the implementation of new biotechnological processes. Hence, the intention of the present study was to investigate the temperature and pressure stability of MsP1, as well as to prove the enzyme's uncommon dimeric occurrence in the culture supernatant. To this end, a series of biophysical techniques (DSC, fluorescence and FTIR spectroscopy, small-angle X-ray scattering) was combined with functional studies of the enzyme. In fact, interest in pressure as an additional thermodynamic and kinetic variable in physico-chemical studies of biological materials has been growing also in recent years [7–11]. The behavior of all systems under high pressure is governed by Le Châtelier's principle, which predicts that the application of pressure shifts the equilibrium towards the state that occupies a smaller volume, and accelerates processes for which the transition state has a smaller volume than the ground state. Besides the general physicochemical interest in using high pressure as a tool for understanding the structure, energetics and kinetic processes of biomolecules, high pressure is also of increasing biotechnological interest (e.g., for high pressure food processing, baroenzymology) [10–13]. 2. Materials and methods 2.1. General
⁎ Corresponding author. Tel.:+49 641 99 34 900; fax: +49 641 99 34 909. E-mail address: [email protected] (H. Zorn). 1570-9639/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.bbapap.2009.03.015
The M. scorodonius strain (CBS 137.86) was obtained from the Dutch “Centraalbureau voor Schimmelcultures”, Baarn. Enzyme production and purification were performed as described previously [2].
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2.2. FT-IR spectroscopy 1 mL of the purified enzyme solution in 50 mM aqueous phosphate buffer (pH 6.0) or 30 mM Bis–Tris-Buffer (pH 5.8; c = 9.6 mg/mL), respectively, was lyophilized overnight, and the resulting powder was suspended in 200 μL D2O. The final protein concentration (2.1% (w/v) and 4.4% (w/v), respectively) was determined by UV-spectroscopy at λ = 280 nm. Due to their different temperature and pressure sensitivities [14], phosphate buffer was used for the analysis of the temperature dependency, and Bis–Tris buffer for the high pressure measurements. For the temperature dependent experiments, a Nicolet 5700 FT-IR spectrometer (Thermo Fisher Scientific, Waltham, USA), equipped with a liquid nitrogen cooled MCT detector (HgCdTe), was used. Approximately 20 μL of the sample were filled between CaF2 windows, which were separated by a 50 μm Mylar spacer. The sample cell was placed in a thermostatted jacket, and an external water bath was used for temperature control (accuracy: ± 0.2 °C). During the measurements, the sample chamber was purged constantly with dry air. Spectral recording was performed between 4000 and 1100 cm− 1. To improve the signal to noise ratio, 256 spectra were summed up for each measuring point (resolution: 2 cm− 1). Apodization was done with a Happ–Genzel function. Data acquisition was performed between 10 and 80 °C with 5 °C increments; the equilibration time before each measurement was 15 min. The pressure dependent measurements were conducted at 30 and 60 °C with a Nicolet MAGNA 550 FT-IR spectrometer. Pressure generation was achieved with a thermostatted High Pressure Diamond Optics P-series diamond anvil cell with type IIa diamonds (High Pressure Diamond Optics Inc., Tucson, USA). A 50 μm thick gasket of stainless steel with a 0.45 mm drilling was placed between the two diamonds holding ∼10 nL of the sample. Pressure was determined with co-added BaSO4, which shows the characteristic pressure sensitive symmetric sulfate stretching mode around 983 cm− 1 that increases linearly with pressure (accuracy: ± 300 bar [15]). All other technical parameters were identical to the temperature dependent measurements. Spectral processing and deconvolution of the amide-I′-band was done with the GRAMS/AI 8.0 software package (Thermo Fisher Scientific, Waltham, USA). After subtraction of noise and the temperature-corresponding buffer background (D2O), all spectra were normalized to the same area to ensure comparability. Finally, the amide-I′-region was fitted with the number of subbands (which are typical for particular secondary structure elements) detected by the 2nd derivative using a Voigt-function. Peak assignment to the different secondary structures was done according to the literature [16]. As the transition dipole moments of the different conformers might be slightly different, we focus essentially on the relative changes of the peak areas as a function of temperature and pressure.
buffer mixture (Bis–Tris; pH 5.8) was dissolved in 100 μL H2O, yielding a 1.9% (w/v) solution. 20 μL of the sample were placed in a high pressure X-ray sample cell. The cell was equipped with KaptonÒ polymer windows, and the sample detector distance was 0.905 m. The plots of the diffraction intensity versus reciprocal spacings (Q, Q = 4πsinΘ / λ; 2Θ is the scattering angle and λ the wavelength of the radiation) were recorded using MATLAB™ based software, written by the ESRF staff. The pressure dependent measurements were performed at 30 °C. The maximum pressure applied was 3000 bar (accuracy: ± 20 bar). Measurements were performed at steps of 100 bar in the upward and downward direction. Additional measurements under the same conditions were taken with a pure buffer sample for background correction. Temperature dependent SAXS measurements were conducted with a Kratky-camera (Model KKK, Anton Paar, Graz, Austria). The X-ray energy was 8.05 keV (l = 0.154 Å). After dissolving the lyophilized protein-buffer mixture (phosphate buffer; pH 6.0) in 100 μL water, yielding a 1.5% (w/v) protein solution, measurements were executed at 30, 60, and 70 °C. Temperature control was achieved via an external water bath (DT = ± 1.5 °C). 2.5. Fluorescence spectroscopy Temperature dependent fluorescence spectroscopic measurements were performed with a Perkin Elmer LS 55 fluorescence spectrometer (PerkinElmer Inc., Waltham, USA), whereas the pressure dependent measurements were performed on a K2 multifrequency phase and modulation fluorometer (ISS Inc., Champaign, USA). For the temperature dependent measurements, the sample (∼300 mL of a 0.5 mg/mL enzyme solution in 50 mM phosphate buffer (pH 6.0)) was filled in a 3 mm cuvette equipped with quartz windows. Temperature control was achieved via an external water bath. Spectra were recorded in 5 °C steps, and the temperature-difference between bath and cuvette was measured and corrected afterwards. Before each measurement, the sample was allowed to equilibrate for 5 min. The tryptophan residues were excited at 295 nm, and the resulting emission was recorded from 307 to 400 nm. For the pressure dependent measurements at 30 °C, a custom made high pressure cell, consisting of a beryllium–copper alloy, with ethanol as pressurizing medium was used. Here, 1 mL of a 1.3 mg/mL solution in 30 mM Bis–Tris buffer (pH 5.8) was filled in a pressure-resistant quartz flask, tightly sealed with heat and solvent resistant laboratory film. Pressure was generated via an external manometer (accuracy: ± 10 bar). Spectra were recorded every 100 bar up to 2200 bar, with an equilibration time of 5 min. After subtraction of buffer background, spectral changes were monitored by calculating the center of spectral mass [17]. 2.6. Enzyme assays
2.3. Differential scanning calorimetry (DSC) For the DSC-measurements, a 0.19% (w/v) enzyme solution in 50 mM phosphate buffer (pH 6.0) was used. Measurements were conducted in duplicate with a VP DSC calorimeter (MicroCal, Northampton, USA). The sample cell of the calorimeter was filled with 0.5 mL of the enzyme solution, whereas the reference cell was filled with the protein-free buffer. The differential heat flow was recorded between 10 and 100 °C with a heating rate of 40 °C/h. Data analysis was performed with a customized version of ORIGIN 7 (MicroCal). 2.4. Synchrotron small-angle X-ray scattering (SAXS) The SAXS experiments were performed at the ID2 high brilliance beamline at the European Synchrotron Radiation Facility (ESRF) in Grenoble, France. The medium X-ray energy was 12.46 keV, corresponding to a wavelength of 0.995 Å. The lyophilized protein-
2.6.1. Preparation of β-carotene solution 5 mg of β-carotene (Sigma-Aldrich, Germany) and 0.5 g of Tween 80 were dissolved in 20 mL dichloromethane. The solvent was evaporated under vacuum, and the residue was suspended immediately in 30 mL water. The suspension was filtered through a 0.22 μm syringe filter and filled up to 50 mL with distilled water. This solution was prepared freshly before each measurement. Temperature dependent enzymatic measurements were performed on a PerkinElmer Lambda 25 UV/VIS spectral photometer (PerkinElmer Inc., Waltham, USA), which was temperature-controlled by an external water bath (DT = ±1.5 °C). The activity was quantified by the ABTS-assay [18]. The reaction mixture contained 700 μL 50 mM sodium acetate buffer (pH 3.5), 100 μL 5 mM 2,2-azinobis-(3-ethyl benzthiazoline-6-sulphonate) (ABTS), 100 μL MsP1-solution (10 mUABTS, 30 °C, pH 5.8) and 100 μL 20 mM H2O2. The increasing absorbance was monitored at l = 420 nm from 30 to 70 °C in 5 °C steps
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over 360 s. One unit of enzyme activity was defined as the amount of enzyme oxidizing 1 μmol of substrate per minute. The high pressure measurements were performed on a K2 multifrequency phase and modulation fluorometer, which was adjusted to collect absorption spectra. All other technical details were identical to the high pressure fluorescence measurements (see above). For the ABTS-assay, 1.06 mL of the reaction mixture (930 μL 33 mM Bis–Tris buffer, pH 5.8, 50 μL 5 mM ABTS, 30 μL MsP1-solution (3 mUABTS, 30 °C, pH 5.8) and 50 μL 20 mM H2O2) were placed in a pressure-resistant and tightly sealed quartz flask. For every pressure step, the increase in absorbance was measured over 600 s at l = 420 nm and 30 °C in a pressure range from 1 to 2500 bar. The β-carotene assay was performed similarly with 1.06 mL reaction mixture (820 μL 33 mM Bis–Tris buffer, pH 5.8, 150 μL β-carotene solution, 50 μL MsP1-solution (0.5 mUβ-carotene, 30 °C, pH 5.8) and 40 μL 20 mM H2O2). For every pressure step, the decrease in
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absorbance was measured at l = 450 nm and 30 °C over 1000 s from 1 bar to 2500 bar. 2.7. Protein quantification The protein concentration was calculated using an absorption coefficient of MsP1 of 27,960 M− 1 cm− 1 at 280 nm (ProtParam, [19]). 3. Results 3.1. Temperature and pressure dependent FT-IR experiments To monitor the conformational changes of the protein, the amide-I′-band at ∼ 1700–1600 cm− 1 was analyzed. The relative area of each subband corresponds in good approximation to the fraction of the respective secondary structure [20–23]. A representative example
Fig. 1. (a) Deconvoluted and fitted amide-I′-FT-IR spectrum of MSP-1 at 10 °C. (b) Second derivative of the amide-I′-FT-IR spectrum of MSP-1 at 10 °C. Negative peaks correspond to the loci of chosen FT-IR subbands.
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Fig. 2. Temperature dependence of the relative infrared peak areas of the subbands of MSP-1, which are assigned to the following secondary structures: ▴ α-helices, ● loops and turns, □ intramolecular β-sheets, and Δ intermolecular β-sheets (commonly associated with aggregates).
of the deconvoluted amide-I′-band after peak fitting is depicted in Fig. 1. Four main subbands were deduced from the second derivative spectra, which appeared at ∼1672, ∼1657, ∼ 1642, and ∼1630 cm− 1. These subbands correspond to loops and turns, α-helices, random coil structures, and intramolecular β-sheets, respectively. The positions of these subbands do not change significantly upon heating. As depicted in Fig. 2, MsP1 reveals to be highly α-helical, with an approximately equal fraction of unordered structures. Also, minor contributions of intramolecular β-sheets and turns are present. With increasing temperature, no apparent change in the band areas are detected until ∼65 °C, where the α-helical content starts to decrease and the unordered fraction increases concomitantly (Fig. 2). Simultaneously, a new β-sheet fraction at ∼1617 cm− 1 appears at ∼ 70 °C, which can be ascribed to formation of intermolecular β-sheets in aggregated structures. This behavior is in good agreement with the temperature dependence of the enzyme activity, indicating an abrupt loss of enzyme activity above about 60 °C (see below). With increasing hydrostatic pressure at 60 °C, the subbands retain their position, and no significant change of the enzyme's secondary structure is observed (Fig. 3), indicating a high pressure stability of MsP1. Above ∼ 8 kbar, only a minor increase in unordered structures at
▪ random coil structure,
the expense of turns and α-helical conformations is observed. Similar results were obtained at 30 °C (data not shown). A small percentage of aggregates were present from the beginning, which may be ascribed to the high protein concentration necessary for the FTIR spectroscopic measurements in the diamond anvil cell. 3.2. Differential scanning calorimetry Up to ∼60 °C, the differential heat capacity Cp (with respect to pure buffer solution) increases almost linearly with temperature, peaks at ∼63 °C, and decreases abruptly above ∼78 °C (data now shown), where, according to the FTIR data, thermal aggregation of the protein takes place. In fact, protein aggregation is generally an exothermic process [24]. 3.3. Temperature and pressure dependent fluorescence spectroscopy The MSP-1 dimer contains 8 tryptophane residues (i.e., there are 4 Trp residues per monomer), which are located in different parts of the three-dimensional structure. The fluorescence spectrum of the protein thus gives an average over all contributing intrinsic
Fig. 3. Pressure dependence of the secondary structure elements of MSP-1 as revealed from the FTIR spectra at 60 °C. For symbol denotation, see Fig. 2.
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fluorophores. Generally, a red-shift of this band indicates a more polar environment of the fluorescent tryptophane residues, which occurs upon protein unfolding and denaturation [17]. From 15 to 50 °C, the center of spectral mass (CSM) of the Trp-emission exhibits a linear blue-shift, which can be ascribed to an essentially hydrophobic environment, indicating that the protein is still in a compact state. Beyond 50 °C, a pronounced red-shift up to 70 °C is observed, where a change in slope of the temperature dependence of the emission wavelength, l (T), and a further red-shift is detected (Fig. 4). The redshift above 50 °C can be explained by a temperature-induced conformational change with subsequent increasing exposure of tryptophane residues. The midpoint of the transition occurs at around 63 °C. By comparing the final CSM-value of ∼357 nm at 85 °C with the one of fully hydrated tryptophane (∼ 362 nm, [25]), it can be concluded, that most Trp residues are largely, but not fully, unshielded at this temperature. The change in slope at ∼ 68 °C may be ascribed to the end of the thermally induced dimer dissociation process that has also been detected in the SAXS data described below, which is then followed by the unfolding reaction of the protein. This interpretation is in good agreement with the FTIR- and DSC-data. Notably, the corresponding loss of secondary structure up to 80 °C, as derived from the infrared spectroscopic data, is rather small (Fig. 2). At 30 °C, the pressure dependent CSM-values exhibit a small and almost linear red-shift with increasing pressure (Fig. 5). A discontinuity with a subsequent change in slope is observed between 500 and 600 bar, however, which might be due to a minor structural change, as corroborated by the SAXS measurements (see below). Surprisingly, the enzyme activity under high hydrostatic pressure conditions shows a maximum at about 500 bar as well at the same temperature (see below). Hence it is very likely that this minor structural change is responsible for the observed change in enzyme activity. The increasing red-shift at pressures above 700 bar may be ascribed to the intrinsic pressure effect on the tryptophane residue [26] and, possibly, increasing pressure-induced fluctuations of the protein structure, which generally increase upon pressurization due to a weakening of hydrophobic interactions [7–10]. 3.4. Small-angle X-ray scattering (SAXS) To reveal details of these pressure-induced small conformational changes of the protein in the lower pressure regime, SAXS measurements were performed as a function of pressure up to 3 kbar. Fig. 6 shows the small-angle X-ray scattering intensity as a function of momentum transfer, I(Q), for pressures from 1 bar to 3 kbar at T = 30 °C. The shape of the SAXS curve with a breakpoint in the slope
Fig. 4. Temperature dependence of the spectral center of mass (bλN) of the Trpfluorescence of MSP-1 from 15 to 86 °C.
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Fig. 5. Pressure dependence of the spectral center of mass (bλN) of the Trp-fluorescence of MSP-1 at 30 °C.
of I(Q) at about 1.2 nm− 1 is typical for dimeric protein structures [27]. Interestingly, upon pressurization, changes occur only in the intermediate momentum transfer region from 1.25 to 3 nm− 1, indicating only minor medium-range-order changes of the protein conformation in this pressure range. Besides, with increasing pressure, a decrease in scattering intensity is observed, only, which is due to a decrease of the electron density contrast between the protein and the surrounding water upon pressurization. The radius of gyration, Rg, was determined from the Guinier-plots (lnI versus Q2) in the low-Q regime as well as from the pair-distance distribution function [21]. Within the accuracy of the measurements, a constant value of Rg of 2. 41 ± 0.05 nm was found up to pressures of 3 kbar (data not shown). For further data analysis, the measured I(Q) data were subjected to indirect Fourier transformation using the program GNOM, which calculates the distance distribution function p(r) for monodisperse systems from the one-dimensional scattering curves (Fig. 7). Interestingly, p(r) exhibits two more or less broad maxima with a minimum around 1.5 nm, again indicating the presence of a dimeric structure. The first maximum at an r-value of about 0.5 nm represents distances with high electron densities, which are basically not changing with pressure. With increasing pressure, a minimum is emerging at intermediate distances of approx. 3 nm, however, which might be due to a decreasing electron density in the connection region of the two monomeric parts of the protein. The change of p(r)
Fig. 6. SAXS data of MSP-1 at selected pressures and T = 30 °C.
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Fig. 7. Pair-distance distribution function of MSP-1 at selected pressures and T = 30 °C.
between 2 and 5 nm thus originates from a pressure-induced small tertiary structural change in the pressure range starting below 1 kbar. After all, no dissociation and no significant unfolding or major changes in tertiary structure can be deduced from the analysis of the pressure dependent SAXS measurements up to 3 kbar. The Guinier-plots of the temperature dependent measurements at ambient pressure exhibit an Rg value of ∼2.4 nm at 30 and 60 °C. At 70 °C, this value first drops to ∼1.2 nm and subsequently increases drastically due to aggregation of the protein (data not shown as the Rg of such a large polydisperse aggregate cannot be determined). This clearly indicates a temperature-induced dissociation of the MSP-1 dimer into monomers, which manifests itself in a decrease of the enzyme activity as observed above ∼60 °C (see below), as well as in the aggregation of the protein as revealed by the DSC and FTIR spectroscopic data. Hence, the catalytic activity of the enzyme seems to be restricted to the integrity of the dimeric structure. 3.5. Temperature and pressure dependence of MsP1 activity Maximum activity of MsP1 was detected between 55 and 60 °C (Fig. 8). The decrease of activity above 60 °C is in agreement with the FTIR spectroscopy data, which indicate conformational changes beyond ∼65 °C. When the enzyme was incubated at 50 °C for 24 h,
the loss of enzyme activity amounted to less than 10%, only (data not shown). The catalytic properties of MsP1 were further characterized by steady-state kinetic analyses at 27 °C and a constant H2O2 concentration of 20 mM. Under these experimental conditions, Km and kcat values of 60 μM and 0.8 s− 1, respectively, were determined. For evaluation of the pressure stability of MsP1, two assays were employed: ABTS as a common peroxidase substrate, and additionally β-carotene, representing a potential industrial application of MsP1 [2]. A greenish blue radical cation is formed from ABTS, and βcarotene is degraded to norisoprenoids by peroxidase activity in the presence of hydrogen peroxide. The absorbance of the ABTS solution slightly depended on the pressure, which was considered in the calculation of the enzyme activity. No changes of the extinction coefficient were observed with β-carotene with increasing pressure (data not shown). As depicted in Fig. 9, for the oxidation of both substrates, activity optima at about 500 bar were determined. Compared to atmospheric pressure, the enzymatic activity of MsP1 is roughly doubled at 500 bar. Beyond 800 bar, the activity steadily decreases to reach the level of atmospheric pressure for β-carotene degradation at 2300 bar, and for ABTS oxidation at about 1100 bar. Compared to the activity at atmospheric pressure, only a negligible activity loss commenced even at 2500 bar. 4. Discussion and conclusions Compared to other members of the DyP-type enzyme family, MsP1 exhibits a remarkably high temperature optimum and a good thermostability. For the peroxidase DyP from T. cucumeris (formerly denominated as Geotrichum candidum), which is capable of decolorizing various synthetic dyes, an optimal temperature of 30 °C has been described [28]. Lignolytic enzymes like manganese peroxidases, laccases, and members of the DyP family represent interesting tools for the low energy and low cost alternatives for industrial processes in which lignin modification is of importance, such as paper and pulp processing and processes for valorization of agricultural residues. The polymeric carbohydrates of agricultural residues, cellulose and hemicellulose are typically hydrolyzed by carbohydrases up to temperatures of 50 °C. More recent developments of cellulases active beyond 50 °C is of interest, since reactions are energetically more favourable at higher temperatures and they give additional benefits in processes, such as decrease of viscosity [29,30]. Thus, simultaneous delignification and saccharification at higher temperatures requires thermostable delignifying enzymes.
Fig. 8. Temperature dependence of the enzyme activity of MsP1 (oxidation of ABTS).
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Fig. 9. Pressure dependence of the enzyme activity of MsP1 for the oxidation of ABTS and β-carotene (at 30 °C).
Remarkably, the activity of MsP1 increases until ∼ 500 bar, before it starts to decrease again to about the ambient pressure activity level. Conversely, various enzymes are markedly inactivated upon high pressure treatment of about 2 kbar. Activity losses of 20 and 50% have been reported for soy lipoxygenase and horseradish peroxidase, respectively, after 15 min exposure to a hydrostatic pressure of 2 kbar at 55 °C. Total inactivation of the lipoxygenase was observed at 5.7 kbar [31]. On the other hand, some proteins are stabilized at high pressure. Up to 4 kbar were required to maintain a carboxypeptidase from the archaebacterium Sulfolobus solfataricus in its active state at higher temperatures [32], and L-galactosidases from E. coli and Aspergillus oryzae were stabilized in the range of 2–3 kbar at 50 and 65 °C, respectively [33]. The pressure level causing full dissociation of the MsP1 dimer could not be revealed in the present study. Other oligomeric enzymes such as glyceraldehyde-3-phosphate dehydrogenase, lactate dehydrogenase, and malate dehydrogenase were found to reversibly dissociate and re-associate upon application and release of hydrostatic pressure up to 2 kbar [33]. The increased activity of MsP1 at pressures around 500 bar may result from slight structural changes stabilizing the transition state of the enzymatic reaction. Generally, high pressure may influence enzyme-catalyzed reactions either by enzyme conformational changes, or by modification of the reaction mechanism [13]. In fact, high pressure has been used to enhance the activity or selectivity of a number of enzymes, paving the way for new applications. Most of the applications have been reported from the food processing industry, especially from dairy and juice production [34]. For example, the reduction of bitterness in citrus juices correlates directly with the concentration of naringin. The enzyme naringinase hydrolyses naringin to the tasteless naringenin, and the activity of the enzyme was shown to increase 3-fold under high pressure conditions (1.6 kbar) [35]. The use of microbial transglutaminase (MTG) under hydrostatic pressure of up to 6 kbar allowed for the cross-linking of proteins which could not be affected by MTG at atmospheric pressure [36]. However, at higher pressures the tertiary structure of the native enzyme was impaired, and α-helical secondary structures were unfolded [37]. No significant changes in the secondary structure of MsP1 occurred up to 10 kbar, minor changes – possibly a slight pressure-induced swelling, indicating the onset of the unfolding reaction – occur above about 8 kbar up to 10 kbar, the maximum pressure reached, only. Regarding this remarkable high pressure stability of MsP1, it may represent a valuable tool for industrial high pressure applications. Delignification of agricultural residues represents an interesting target. The common pretreatment of lignocellulosic substrates prior
to carbohydrate hydrolysis and fermentation to ethanol is performed by steam explosion [38,39], dilute acid [40,41], or an organosolv process [42]. However, these procedures come along with the formation of toxic by-products, which impede the downstream enzymatic hydrolysis and the fermentative conversion of the released sugars into ethanol [40,43–47]. An entirely enzymatic delignification process may overcome these obstacles. Acknowledgements Financial support from the Deutsche Forschungsgemeinschaft (DFG), the Deutsche Bundesstiftung Umwelt (DBU), and the Fonds der Chemischen Industrie is gratefully acknowledged. References [1] G.C. Ainsworth, F.K. Sparrow, A.S. Sussman, A taxonomic review with keys: basidiomycetes and lower fungi, in: G.C. Ainsworth, F.K. Sparrow, A.S. Sussman (Eds.), The Fungi, an Advanced Treatise, vol. 4B, Academic Press, Orlando, 1973. [2] M. Scheibner, B. Hülsdau, K. Zelena, M. Nimtz, L. de Boer, R.G. Berger, H. Zorn, Novel peroxidases of Marasmius scorodonius degrade β-carotene, Appl. Microbiol. Biotechnol. 27 (2008) 1241–1250. [3] V. Faraco, A. Piscitelli, G. Sannia, P. Giardina, Identification of a new member of the dye-decolorizing peroxidase family from Pleurotus ostreatus, World J. Microbiol. Biotechnol. 23 (2007) 889–893. [4] B. Hülsdau, Oxidativer Abbau von Carotinoiden durch Pilzenzyme, Dissertation, Gottfried Wilhelm Leibniz Universität, Hannover, 2007. [5] H. Bouws, A. Wattenberg, H. Zorn, Fungal secretomes—nature's toolbox for white biotechnology, Appl. Microbiol. Biotechnol. 80 (2008) 381–388. [6] H. Zorn, S. Langhoff, M. Scheibner, M. Nimtz, R.G. Berger, A Peroxidase from Lepista irina cleaves β-carotene to flavour compounds, Biol. Chem. 384 (2003) 1049–1056. [7] F. Meersman, L. Smeller, K. Heremans, Protein stability and dynamics in the pressure–temperature plane, Biochim. Biophys. Acta 1764 (2006) 346–354. [8] I. Daniel, P. Oger, R. Winter, Origins of life and biochemistry under high pressure conditions, Chem. Soc. Rev. 35 (2006) 858–875. [9] J.L. Silva, D. Foguel, C.A. Royer;, Pressure provides new insights into protein folding, dynamics and structure, Trends Biochem. Sci. 26 (2001) 612–618. [10] R. Winter (Ed.), High Pressure Bioscience and Biotechnology II, Springer-Verlag, Heidelberg, Germany, 2003. [11] F. Abe, Exploration of the effects of high hydrostatic pressure on microbial growth, physiology and survival: perspectives from piezophysiology, Biosci. Biotechnol. Biochem. 71 (2007) 2347–2357. [12] C. Balny, P. Masson, K. Heremans, High pressure effects on biological macromolecules: from structural changes to alteration of cellular process, Biochim. Biophys. Acta 1595 (2002) 3–10. [13] D.B. Northrop, Effects of high pressure on enzymatic activity, Biochim. Biophys. Acta 1595 (2002) 71–79. [14] R.C. Neumann, W. Kauzmann, A. Zipp, Pressure dependence of weak acid ionization in aqueous buffers, J. Phys. Chem. 77 (1973) 2687–2691. [15] P.T.T. Wong, D.J. Moffat, A new internal pressure calibrant for high-pressure infrared spectroscopy of aqueous systems, Appl. Spectr. 43 (1989) 1279–1281. [16] A. Barth, Infrared spectroscopy of proteins, Biochim. Biophys. Acta 1767 (2007) 1073–1101.
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