Article pubs.acs.org/Langmuir
Activation Volumes of Enzymes Adsorbed on Silica Particles Vitor Schuabb and Claus Czeslik* Department of Chemistry and Chemical Biology, TU Dortmund University, D-44221 Dortmund, Germany S Supporting Information *
ABSTRACT: The immobilization of enzymes on carrier particles is useful in many biotechnological processes. In this way, enzymes can be separated from the reaction solution by filtering and can be reused in several cycles. On the other hand, there is a series of examples of free enzymes in solution that can be activated by the application of pressure. Thus, a potential loss of enzymatic activity upon immobilization on carrier particles might be compensated by pressure. In this study, we have determined the activation volumes of two enzymes, α-chymotrypsin (α-CT) and horseradish peroxidase (HRP), when they are adsorbed on silica particles and free in solution. The experiments have been carried out using fluorescence assays under pressures up to 2000 bar. In all cases, activation volumes were found to depend on the applied pressure, suggesting different compressions of the enzyme-substrate complex and the transition state. The volume profiles of free and adsorbed HRP are similar. For α-CT, larger activation volumes are found in the adsorbed state. However, up to about 500 bar, the enzymatic reaction of α-CT, which is adsorbed on silica particles, is characterized by a negative activation volume. This observation suggests that application of pressure might indeed be useful to enhance the activity of enzymes on carrier particles.
1. INTRODUCTION Enzymes are nature’s catalysts. Their function has been optimized in a perfect way in order to catalyze chemical reactions under mild conditions, such as neutral pH-values, ambient pressure, and almost ambient temperatures. Usually, they are highly specific for a particular reaction pathway, enabling yields with high enantiomeric excess. Therefore, enzymes are very attractive for use in biotechnological, pharmaceutical, and biomedical processes. The major drawbacks of using enzymes compared to chemical catalysts are their relatively low stability and their relatively high cost. Expensive enzymes are worth being recovered from the reaction media to be reused in many cycles. Removing an enzyme from a reaction mixture can be carried out easily by filtration or centrifugation, when the enzyme is immobilized on carrier particles large enough to be retained or sedimented.1,2 However, any deviation from the protein’s natural environment, such as an aqueous−solid interface, can cause a structural change and denaturation of the protein.3 Furthermore, interactions with a solid substrate can also slow down the dynamics of a protein.4 Additionally, overall, enzymatic activity at interfaces may also be decreased by an unfavorable orientation of the enzyme molecules because the active site can be blocked either by the material surface or by neighboring enzyme molecules. To preserve enzymatic activity at aqueous−solid interfaces, different strategies have been proposed and are applied.1,2 For example, the use of hydrophilic surfaces, the incorporation of an enzyme into porous materials, and genetic engineering can be helpful to overcome limited enzymatic activity at aqueous− solid interfaces. In this study, we will explore pressure as a new tool to control and potentially enhance enzymatic activity at © 2014 American Chemical Society
aqueous−solid interfaces. Generally, the effect of pressure on enzymes is two-fold, because enzymes can be regarded as a normal protein with a folded conformation, which can be destroyed under pressure, and as a biological catalyst, whose rate can be modified under pressure. Although a folded conformation of a protein represents an almost perfect dense packing, some void volumes and cavities remain, which are filled by water upon unfolding. Thus, there is a negative volume change, ΔV, associated with the unfolding of proteins (at least at ambient temperatures).5,6 Other contributions to ΔV are discussed in the literature, such as electrostriction upon exposure of charged and polar residues to the solvent.6 Due to this volume decrease, protein unfolding and subsequent loss of enzymatic activity is often observed at pressures of several 1000 bar. In addition, the catalytic properties of enzymes have been shown to sensitively depend on pressure below the pressure of unfolding in the folded state.7−9 A transition state is favored under pressure, when it has a smaller volume than the reactants. This is expressed as negative activation volume. The activation volume, ΔV‡, is related to the change of the rate constant, k, with pressure, p, according to8,9 ⎛ ∂ ln k ⎞ ΔV ‡ ⎜ ⎟ =− RT ⎝ ∂p ⎠T
(1)
Received: September 9, 2014 Revised: November 19, 2014 Published: December 5, 2014 15496
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somewhat larger diameters have been found from dynamic light scattering experiments.17 Whereas Ludox AM silica particles have a negative surface charge, Ludox CL silica particles are coated with alumina, converting their charge from negative to positive. Enzymatic Assays. α-CT was adsorbed on silica particles by mixing stock solutions of α-CT (250 μL of a 2 mg mL−1 solution) and Ludox AM (68 μL of a 1.21 g mL−1 solution) in 682 μL buffer solution (20 mM Tris-HCl, pH = 7.8) and incubating the mixed solution at 4 °C over 3 h. The calculated surface coverage of the silica particles by α-CT is on the order of 10−2 = 1% (calculation given in the Supporting Information). To probe the enzymatic activity of free or adsorbed α-CT (in the absence or presence of silica particles), samples contained 1.4 μg mL−1 of α-CT, 20 mM Tris-HCl buffer (pH = 7.8), and 1.57 mg mL−1 EnzCheck substrate. This substrate consists of a fluorophore and a quencher moiety that are covalently connected via an amino acid chain. Upon chain cleavage by α-CT, the quencher separates from the fluorophore that is then free to emit fluorescence.18 Experiments were carried out with a K2 fluorescence spectrometer from ISS (Champaign, Illinois, USA) with a high pressure sample cell from ISS. Fluorescence was excited at 502 nm with a Xe arc lamp and detected at 528 nm in photon counting mode. All measurements were performed at 25 °C. HRP was adsorbed on silica particles by mixing stock solutions of HRP (1.1 μL of a 23 μg mL−1 solution) and Ludox CL (100 μL of a 1.23 g mL−1 solution) in 1 mL buffer solution (50 mM sodium phosphate, pH = 7.4) and incubating the mixed solution at 4 °C over 3 h. The calculated surface coverage of the silica particles by HRP is on the order of 10−7 = 10−5 % (calculation given in Supporting Information). To probe the enzymatic activity of free or adsorbed HRP (in the absence or presence of silica particles), samples contained 1.16 μg mL−1 of HRP (0.026 μM), 50 mM sodium phosphate buffer (pH = 7.4), 50 μM Amplex Red substrate, and 1 mM H2O2. HRP catalyzes the oxidation of the Amplex Red substrate to the resorufin product by H2O2. Because resorufin is fluorescent, it can be quantified easily.18 Fluorescence was also measured using the K2 fluorescence spectrometer. Fluorescence of resorufin was excited at 530 nm and detected at 585 nm. All measurements were performed at 25 °C. Degree of Adsorption. Ludox CL silica particles have a positive surface charge due to alumina coating and interact with negatively charged amino acid residues of HRP. The net charge of HRP at neutral pH-values is very close to 0.19,20 α-CT, which has a small positive net charge at neutral pH-values (the isoelectric point is at pH = 9),21,22 can bind to the negatively charged Ludox AM silica particles. To determine the degree of enzyme adsorption on the silica particles, we have used Millipore Amicon Ultra centrifugal filters (50 kDa), which can only be passed by free, nonadsorbed enzyme molecules. Silica particles with adsorbed enzyme molecules cannot pass. The filters were loaded with the mixed solutions of enzymes and silica particles. After centrifugation at 14000g for 10 min at 4 °C, the filtrates were analyzed for free enzymes. No free HRP or α-CT could be detected by UV-spectroscopy. Moreover, no enzymatic activity of HRP or α-CT could be detected in the filtrate when carrying out the enzymatic assays described above. The fluorescence intensity did not change with time upon addition of the corresponding substrate (Figure S1, Supporting Information). Thus, within the sensitivity of the applied UV and fluorescence measurements, all enzyme molecules are adsorbed in the presence of the silica particles. Data Collection and Analysis. Fluorescence intensity is proportional to the fluorophore concentration, c. Thus, the rate of product formation, dc/dt, is proportional (not equal) to the slope of the fluorescence intensity that is plotted as a function of time (Figure 2). To remove all instrumental factors affecting the fluorescence intensity, the slope observed under pressure, S, has been normalized to the slope observed under ambient pressure, S0. Each fluorescence assay started with a few minutes of data collection at 1 bar followed by a 1−2 min period, during which the pressure has been increased, and a further time period under high pressure. The ratio of the slopes measured directly before and after the pressure change can be related to the ratio of the corresponding rate constants, because the concentrations are very similar before and after the pressure change:
Positive and negative activation volumes are illustrated in Figure 1. Depending on the sign of the activation volume, enzymatic reactions can proceed faster (negative sign) or slower (positive sign) under pressure.
Figure 1. Schematic simplified volume diagrams of enzymatic reactions. The enzyme-substrate (ES) complex transforms into the transition state (TS), which finally dissociates into the product (P) and the enzyme (E). The transition state can have a higher or a lower volume than the ES complex. As a consequence, the enzymatic reaction is inhibited or accelerated under pressure, respectively. The volume change from the ES complex to the transition state is called the activation volume, ΔV‡.
Here, we have studied two enzymes, horseradish peroxidase (HRP) and α-chymotrypsin (α-CT). A peroxidase catalyzes the oxidation of a substrate by H2O2. In the literature, it is reported that carrot peroxidase loses activity up to 2000 bar, but shows increased activity at 5000 bar.10 However, for HRP, an inactivation has been observed under pressures up to 5000 bar, but this pressure effect seems to depend on the kind of the used substrate.11 Enhanced activity has been reported for HRP that is immobilized in a Langmuir−Blodgett film of a phospholipid.12 α-CT is hydrolyzing peptide bonds. The catalytic activity of α-CT is known to be enhanced at high pressures.13,14 For example, an increase in pressure to 4700 bar at 20 °C results in an acceleration of the hydrolysis by a factor of 6.5.13 Furthermore, α-CT activation can also be achieved by simply adding a polyelectrolyte that is oppositely charged as compared with the substrate.15 When α-CT adsorbs on oppositely charged particles, positively charged substrates are preferred demonstrating the importance of electrostatic interactions for the substrate binding and the release of product.16 Thus, HRP and α-CT appear to be interesting candidates to study the combined effects of an aqueous-solid interface and pressure. As we will show here, pressure has a pronounced effect on enzymatic activities at interfaces and might even be used to enhance activities of immobilized enzymes. Furthermore, we present activation volumes of adsorbed enzymes for the first time, which provide valuable information about the enzymatic mechanisms.
2. MATERIALS AND METHODS Enzymes and Substrates. HRP and Amplex Red substrate (10acetyl-3,7-dihydroxyphenoxazine) were purchased from Thermo Fisher Scientific (Assay Kit A22188). α-CT from bovine pancreas was purchased from Sigma-Aldrich (C7762). The EnzCheck Peptidase/Protease Assay Kit from Thermo Fisher Scientific (Assay Kit E33758) was used to probe the α-CT activity. Ludox AM and Ludox CL colloidal silica particles were purchased from Sigma-Aldrich and used for immobilization of the enzymes. They have an average diameter of about 12 nm according to the manufacturer, although 15497
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added to the sample as an internal pressure sensor. The shift of the 983 cm−1 band to higher wavenumbers is proportional to the applied pressure.23 All spectra are corrected for background measured with buffer solution. D2O was used as the solvent.
3. RESULTS AND DISCUSSION 3.1. Pressure Stability of HRP and α-CT. In this study, we have performed enzymatic assays under pressures up to 2000 bar at 25 °C. It is helpful to separate pressure effects on the enzyme kinetics from pressure effects on the protein conformation. Free HRP and α-CT do not unfold in solution in this pressure range.24−26 However, in the adsorbed state, the conformational stability of proteins is usually lowered. This means that the temperature and the pressure of protein unfolding are significantly lowered after adsorption at an aqueous−solid interface.17,27,28 Thus, we have analyzed the conformation of HRP and α-CT when they are adsorbed on silica particles and, for comparison, free in solution as a function of pressure. Using FTIR spectroscopy in combination with a diamond anvil cell, the amide I′ band (in D2O) has been recorded. This band is known to be sensitive for the secondary structure of a protein. Any change in the secondary structure leads to a change of the band shape, because the amide I′ band is a superposition of the infrared absorption bands of the various secondary structure elements, which have all individual wavenumbers.29 In Figure 3, the amide I′ bands of HRP and αFigure 2. Typical fluorescence intensity data as recorded to probe the enzymatic activities of α-CT and HRP. The fluorescence intensity has been measured at 1 bar and after pressure increase. The slope of the data is proportional to the rate of the enzymatic reaction. The slopes before and after the pressure increase have been determined by linear fits.
S k = S0 k0
(2)
The specific meaning of the rate constants is discussed below. At 1 bar, the slope, S0, was found to be absolutely constant over time in both enzyme assays, indicating that the enzymes were saturated with substrates and were working with maximum rates. Thus, S/S0 = 1 at 1 bar for both enzymes. It follows from eq 1 that a plot of ln(S/S0) as a function of pressure, p, yields a curve whose slope contains the activation volume:
d ln(S /S0) ΔV ‡ =− dp RT
(3)
This equation can be applied to all pressures, p, and does not require a constant activation volume. When the activation volume is pressure dependent, a change in the slope of ln(S/S0) versus p is observed. In addition, the raw slope, as taken from the fit under high pressure (Figure 2), has been corrected for the pressure dependence of the fluorescence quantum yield of the product. This dependence has been measured using an enzymatic assay with free HRP or α-CT (without silica particles) that was allowed to run for about 5 h until the fluorescence intensity of the product reached a constant value, and no additional product was formed anymore. Then, the steady-state fluorescence intensity of this assay was measured as a function of pressure. After normalization to 1 bar (Figure S2), the steady-state fluorescence intensity, I(p), was used to correct the raw slope, Sraw(p), by calculating S(p) = Sraw(p)/I(p). Fourier Transform Infrared (FTIR) Spectroscopy. Infrared absorption spectra of HRP and α-CT were recorded using a Nicolet 6700 Fourier transform infrared spectrometer from Thermo Fisher Scientific at a spectral resolution of 2 cm−1. Sample solutions were analyzed in a thermostated diamond anvil cell. Barium sulfate was
Figure 3. Amide I′ band in the FTIR spectra of free and adsorbed αCT and HRP (selected data). Each diagram contains three overlapping amide I′ bands in the pressure range 1−2300 bar indicating the pressure stability of the secondary structures. Data curves refer to 1 bar (solid line), 1000 bar (dashed line), and 2300 bar (dotted line).
CT both in the adsorbed state on silica particles and free in solution are plotted. Apparently, no significant change of the band shape of the amide I′ bands can be observed, when the pressure is increased up to 2 kbar, indicating stable secondary structures of the enzymes. Indeed, the pressure of unfolding has been reported to be 12 kbar in the case of HRP24 and about 4 kbar in the case of α-CT.25,26 In addition, we have also probed 15498
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the tertiary structure of α-CT by recording the Trp fluorescence band (unfortunately, the Trp fluorescence of HRP was too weak to observe due to intramolecular Trp heme energy transfer30). A red shift of the Trp fluorescence band indicates an exposure of buried Trp residues to water in the course of protein unfolding. As can be seen in Figure S3, the Trp bands of free and adsorbed α-CT do not shift with pressure, clearly indicating an intact tertiary structure up to 2000 bar. For comparison, the Trp band of α-CT has also been measured as a function of temperature (Figure S4). Here, a pronounced red-shift of this band is observed upon thermal unfolding. Thus, pressure-induced changes of the enzyme kinetics of HRP and α-CT (see below) are not related to any pressure-induced partial unfolding of these enzymes. 3.2. Pressure Effects on the Enzymatic Activity of Free and Adsorbed α-CT. α-CT from bovine pancreas has a molar mass of about 25 000 g mol−1 and an isoelectric point of about 9 giving a positive net charge to the enzyme at neutral pHvalues.21,22 However, α-CT binds to and is active on both negatively and positively charged surfaces of polyelectrolyte layers.,31 and it binds to silica particles in a quantitative way (see Materials and Methods section). α-CT cleaves peptide bonds, preferentially those formed by the aromatic amino acids Trp, Tyr, and Phe.32 It hydrolyzes a peptide bond via an initial nucleophilic attack of the residue Ser195, which is located in the active site. The mechanism is of Michaelis−Menten type:32 k1
Figure 4. Pressure dependence of the enzymatic activity of free and adsorbed α-CT. The slopes of the plots are determined by linear fits. They are proportional to the negative activation volume according to eq 3. Each error bar covers the results of two independent measurements. S and S0 are the slopes of the fluorescence intensity as a function of time under high and ambient pressure, respectively.
α ‐CT + S ⇀ α ‐CT·S ↽⎯⎯⎯ k−1
k2
→ P2−α‐CT + P1 k3
⎯⎯⎯→ α ‐CT + P1 + P2 H 2O
Here, S is the substrate, which is hydrolyzed into the products P1 and P2. As described in the Materials and Methods section, we used a large excess of substrate (a few mg mL−1) that saturates the enzyme (a few μg mL−1). In this case, the rate of the enzymatic reaction is determined by step 2 and step 3 with a rate constant of kcat = k2k3/(k2 + k3).9 It has been proposed that k2 is small and rate-limiting for a peptide hydrolysis (this study), whereas k3 is small and rate-limiting for an ester hydrolysis.13,14,32 A back reaction is also avoided by a high substrate concentration. Therefore, the ΔV‡-values of α-CT, as observed in this study, probably refer to step 2 in the reaction scheme, where the enzyme is acylated (P2−α-CT) and an amine (P1) is released. In Figure 4a, logarithmic plots of the normalized fluorescence slopes, S/S0, are shown for α-CT free in solution. According to eq 3, the change of ln(S/S0) with pressure, p, is proportional to the negative activation volume, −ΔV‡. Apparently, the logarithmic plot is not consistent with a constant activation volume over the whole pressure range studied. Indeed, pressure-dependent activation volumes are described as the more usual case.33 The activation volume is the difference of the volumes of the transition state and the enzyme−substrate (ES) complex, ΔV‡ = V‡ − VES. If these volumes, V‡ and VES, are compressed to different extents, a change of ΔV‡ with pressure is the result. In our case, the curved data in Figure 4a can be approximated with two linear ranges from 1 to 500 bar and from 500 to 2000 bar. These linear ranges yield activation volumes of −67 mL mol−1 (1−500 bar) and −15 mL mol−1 (500−2000 bar) using eq 3.
A negative activation volume indicates that the volume of the transition state is smaller than the volume of the ES complex. In this way, high pressure favors the formation of the transition state, because the activation free energy is lowered by pΔV‡. The outcome is a faster enzymatic reaction. Negative activation volumes of α-CT have also been found in the literature.13,14 Taniguchi et al. report activation volumes in the range from −4.4 to −35 mL mol−1 for the enzyme deacylation (step 3 in the reaction scheme).14 Mozhaev et al. studied the acylation of the enzyme (step 2 in the reaction scheme) and observed an activation volume of about −10 mL mol−1 at 20 °C and −25 mL mol−1 at 50 °C.13 These values are in good agreement with our value of −15 mL mol−1 at 25 °C above 500 bar. However, the strongly negative activation volume observed in this study below 500 bar has not been resolved so far. The corresponding data of the α-CT catalysis in the adsorbed state on silica particles are shown in Figure 4b. Remarkably, as found for free α-CT, α-CT that is adsorbed on silica particles has a nonconstant activation volume in the pressure range of 1−2000 bar. There is a slight increase of the enzymatic rate up to 500 bar reaching a maximum at about 750 bar. At higher pressures, the enzymatic rate is decreasing with increasing pressure. From linear fits, activation volumes of −4 mL mol−1 (1−500 bar) and +5 mL mol−1 (1000−2000 bar) can be determined (Figure 4b). When comparing the data of free α-CT (Figure 4a) with those of adsorbed α-CT (Figure 4b), similar pressure profiles can be observed except for an overall reduction of the slopes and a corresponding increase of the activation volumes upon adsorption of α-CT on silica 15499
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enzyme surface enclosing the active site is binding to the silica surface, volume changes occurring during the enzymatic reaction are likely to be distorted. This aspect must also be considered in the interpretation of ΔV‡, and a single pathway might not be enough in the adsorbed state of the enzyme. On the other hand, conformational changes and a lowered thermodynamic stability of the folded state of the adsorbed enzyme do not play a significant role, as can be concluded from the measured FTIR (Figure 3) and fluorescence spectra (Figure S3), which are discussed in section 3.1. However, a clear molecular picture of the immobilization and pressure effects on the α-CT catalysis is not possible at this point. The effect of pressure on the activity of free and adsorbed αCT was found to be partially reversible only. After pressure release from 1000 to 1 bar, the fluorescence intensity was not linear with time, but was curved downward. This suggests a reduced activity due to substrate depletion. Recovered activities are ranging around 73% (free α-CT) and 92% (adsorbed αCT). 3.3. Pressure Effects on the Enzymatic Activity of Free and Adsorbed HRP. For comparison, we have also studied the effect of pressure on the catalytic activity of free and adsorbed HRP. HRP has a molar mass of about 44 000 g mol−1, is mainly α-helical, and binds heme as prosthetic group.36,37 HRP catalyzes the oxidation of a substrate by H2O2. The mechanism is based on the initial binding of an oxygen atom from H2O2 to the iron atom of heme. Then, two electrons are transferred successively from the substrate to the heme, and a water molecule is finally released after uptake of two protons:37,38
particles. Apparently, adsorption adds a positive contribution (in a mathematical sense) to the activation volume. A volume diagram of α-CT catalysis is given in Figure 5. Activation volumes are illustrated as arrows pointing from the
Figure 5. Activation volumes of free and adsorbed α-CT (not to scale, ΔV‡ given in mL mol−1). In both cases, the activation volume depends on the pressure. The ES complex is compressed in a stronger way than the transition state (TS). The volume levels at low and high pressures are arranged assuming a volume reduction of both ES and TS by pressure.
ES complex to the transition state. In the literature, sources of activation volumes are discussed.6−8,34,35 Exposure of hydrophobic and charged amino acid residues to water during the formation of the transition state will cause a higher hydration density and a negative activation volume. Indeed, Mozhaev et al. have attributed the observed negative activation volume of αCT to the increased hydration of the transition state dipole.13 Moreover, small conformational changes of the enzyme forming the transition state can be linked to the generation or the filling of void volumes inside the enzyme. In both cases of free and adsorbed α-CT, we observe a more positive activation volume in the higher pressure range. This may be explained by a stronger compression of the ES complex relative to the transition state, as illustrated in the volume diagram (Figure 5). If VES is compressed in a stronger way than V‡, the activation volume, which is the difference V‡ − VES, becomes larger (more positive). From conformational studies it is clear that α-CT does not undergo significant changes in the secondary and tertiary structure in the pressure range of 1− 2000 bar (Figure 3 and Figure S3). However, we could speculate that the substrate binding in the ES complex leaves some void volume at ambient pressure. At about 500 bar, the substrate molecule might be pushed more deeply into the pocket of the enzyme in a way that substrate-enzyme fitting is optimized and VES becomes smaller. Upon adsorption of α-CT on the silica particles, the activation volumes are increasing (Figure 5). It is likely that an increase of the transition state volume, V‡, is the reason for this finding. When an enzyme is catalyzing a chemical reaction, conformational flexibility of the enzyme is mandatory. However, in the adsorbed state, this conformational flexibility will be reduced. As a consequence, the transition state might have a nonperfect, distorted geometry with an increased void volume, which increases V‡ and ΔV‡ of α-CT in the adsorbed state. Moreover, the location of water molecules in the active site of the enzyme appears to be a crucial factor. There might be gaps in the transition state that are just too small to accommodate a water molecule. There is also a heterogeneous orientation of the enzyme molecules on the silica particles. Thus, there are different enzyme−silica interactions that could lead to different volume profiles. In particular, when the
This mechanism is much more complex than the peptide cleavage by α-CT. The two intermediates in the above scheme are denoted as compound I and compound II. In this study, HRP catalyzes the oxidation of the Amplex Red substrate to the resorufin product by hydrogen peroxide. Resorufin is detected by its fluorescence (see Materials and Methods section). In the enzymatic assay, we have used a large excess of hydrogen peroxide (1 mM) and Amplex Red (50 μM) that bind to HRP (0.026 μM). This excess disfavors the back reaction and ensures the maximum enzymatic rate. The rate-limiting steps will then be the successive transfers of two electrons from an Amplex Red molecule to the heme group. Recent single molecule fluorescence experiments are consistent with a two-electron mechanism, where the two electrons originate from a single Amplex Red molecule and fluorescent resorufin is formed while it is still confined in the enzyme.39 Alternatively, it has been proposed that HRP oxidizes an Amplex Red molecule by a oneelectron transfer only. Afterward, the formed nonfluorescent radical intermediate is released from the enzyme and reacts to resorufin by dismutation.40 The measured enzymatic activity of HRP using Amplex Red as the substrate is shown in Figure 6. The observed slope of the product fluorescence, S, is plotted as a function of pressure, p, on a logarithmic scale (see eq 3). In both cases, free HRP and HRP that is adsorbed on silica particles, the slope of the product fluorescence, and thus the associated rate constant of product formation (see eq 2) decreases as the pressure increases. This indicates positive activation volumes, which increase the activation free energies by the amount pΔV‡. As 15500
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Overall, the volume profiles of free and adsorbed HRP appear to be similar (Fig. 7). Presumably, the mechanism of the enzymatic reaction does not change upon adsorption, and no large distortion of the active site occurs when HRP interacts with the silica particles. However, in contrast to α-CT, activation volumes of HRP tend to be smaller at higher pressures. Above 500 bar, an activation volume of only +2 mL mol−1 indicates that the volumes of the transition state and the ES complex are very similar, and pressure has little effect on the enzymatic rate. Apparently, pressure can compress the transition state stronger than the ES complex leading to a reduction of the activation volume, ΔV‡ = V‡ − VES. So far, a molecular interpretation of the volume diagrams given in Figure 7 is difficult. Assuming the transfers of the two electrons from Amplex Red to HRP are rate-limiting, it can be concluded that the ES complex grows in volume to form the transition state, where the electron transfer can happen, because V‡ > VES. Probably, electron transfer requires some shift of Amplex Red away from the optimum binding position thereby creating some void volume. However, at pressures above about 500 bar, the increased volume of the transition state is suppressed resulting in V‡ ≈ VES. This finding is supported by earlier compressibility measurements of HRP in the low and high pressure range.41 Below 100 bar, HRP has a large compressibility, whereas HRP is less compressible at higher pressures. It has been concluded that the enzyme has soft domains at ambient conditions, which are compressed and become rigid at higher pressures.41 Thus, a larger compressibility of HRP at ambient conditions is linked to a larger density fluctuation, which could be the reason for the larger activation volumes of free and adsorbed HRP that are observed below 500 bar (Figure 7). At higher pressures, a rigid compressed structure of HRP is consistent with little volume change during the oxidation of Amplex Red by HRP. The effect of pressure on the activity of free and adsorbed HRP was found to be partially reversible only. After pressure release from 1000 to 1 bar, the fluorescence intensity was not linear with time, but was curved upward. This suggests a slow recovery of the original conformation. Recovered activities range around 90%.
Figure 6. Pressure dependence of the enzymatic activity of free and adsorbed HRP. The slopes of the plots are determined by linear fits. They are proportional to the negative activation volume according to eq 3. Each error bar covers the results of four independent measurements. S and S0 are the slopes of the fluorescence intensity as a function of time under high and ambient pressure, respectively.
found for α-CT, the plots of ln(S/S0) versus p (Figure 6) are nonlinear, suggesting nonconstant activation volumes of free and adsorbed HRP in the pressure range 1−2000 bar. To estimate activation volumes, we have divided the studied pressure range into a lower and higher pressure range in a similar way as described above for α-CT. From the linear fits (Figure 6) and eq 3, we can derive activation volumes of +16 mL mol−1 (1−500 bar) and +2 mL mol−1 (500−2000 bar) in the case of free HRP. For adsorbed HRP, corresponding values of +27 mL mol−1 (1−250 bar) and +2 mL mol−1 (250−1500 bar) are obtained (note that the error bars in Figure 6 are covering the results of four independent measurements). All activation volumes obtained in this way are summarized in a volume diagram (Figure 7).
4. CONCLUSIONS The effect of pressure on the enzymatic activity of free and adsorbed α-CT and HRP has been studied. α-CT cleaves a peptide bond, whereas HRP oxidizes a substrate. Generally, the immobilization of enzymes on carrier particles provides an easy method to remove expensive enzymes from reaction media for the reuse in additional processes. Because enzymes can lose some biological activity at aqueous−solid interfaces, it is interesting to study pressure as a new tool to activate enzymes in the adsorbed state. In the case of free α-CT, it is known that the enzymatic rate is increasing with increasing pressure. We could confirm this effect, but in addition, we have also observed that the pressure activation of free α-CT is more pronounced in the lower pressure range up to about 500 bar. Upon adsorption on silica particles, a positive contribution is added to the activation volume of α-CT. However, up to 500 bar, the activation volume of adsorbed α-CT is still negative, and the enzymatic activity can still be enhanced by the application of pressure. This finding demonstrates a new potential strategy to optimize enzymatic reactions on carrier particles. On the other hand, the observed activation volumes of HRP are generally positive and similar in the free and adsorbed states. However,
Figure 7. Activation volumes of free and adsorbed HRP (not to scale, ΔV‡ given in mL mol−1). In both cases, the activation volume depends on the pressure. The TS is compressed in a stronger way than the ES complex. The volume levels at low and high pressures are arranged assuming a volume reduction of both ES and TS by pressure. 15501
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(11) García, A. F.; Butz, P.; Tauscher, B. Mechanism-based irreversible inactivation of horseradish peroxidase at 500 MPa. Biotechnol. Prog. 2002, 18, 1076−1081. (12) Schmidt, T. F.; Caseli, L.; Viitala, T.; Oliveira, O. N. Enhanced activity of horseradish peroxidase in Langmuir−Blodgett films of phospholipids. Biochim. Biophys. Acta 2008, 1778, 2291−2297. (13) Mozhaev, V. V.; Lange, R.; Kudryashova, E. V.; Balny, C. Application of high hydrostatic pressure for increased activity and stability of enzymes. Biotechnol. Bioeng. 1996, 52, 320−331. (14) Taniguchi, Y.; Suzuki, K. Pressure inactivation of αchymotrypsin. J. Phys. Chem. 1983, 87, 5185−5193. (15) Kurinomaru, T.; Tomita, S.; Hagihara, Y.; Shiraki, K. Enzyme hyperactivation system based on a complementary charged pair of polyelectrolytes and substrates. Langmuir 2014, 30, 3826−3831. (16) You, C.-C.; Agasti, S. S.; De, M.; Knapp, M. J.; Rotello, V. M. Modulation of the catalytic behavior of α-chymotrypsin at monolayerprotected nanoparticle surfaces. J. Am. Chem. Soc. 2006, 128, 14612− 14618. (17) Czeslik, C.; Winter, R. Effect of temperature on the conformation of lysozyme adsorbed to silica particles. Phys. Chem. Chem. Phys. 2001, 3, 235−239. (18) The Molecular Probes Handbook, http://www.lifetechnologies. com. (19) Ö jteg, G.; Lundahl, P.; Wolgast, M. The net electric charge of proteins. A comparison of determinations by Donnan potential measurements and by gel electrophoresis. Biochim. Biophys. Acta 1989, 991, 317−323. (20) Phelps, C.; Forlani, L.; Antonini, E. The hydrogen ion equilibria of horseradish peroxidase and apoperoxidase. Biochem. J. 1971, 124, 605−614. (21) Ui, N. Isoelectric points and conformation of proteins. Biochim. Biophys. Acta 1971, 229, 582−589. (22) Marini, M. A.; Wunsch, C. The hydrogen ion equilibria of chymotrypsinogen and α-chymotrypsin. Biochemistry 1963, 2, 1454− 1460. (23) Wong, P. T. T.; Moffat, D. J. A new internal pressure calibrant for high-pressure infrared spectroscopy of aqueous systems. Appl. Spectrosc. 1989, 43, 1279−1281. (24) Smeller, L.; Meersman, F.; Fidy, J.; Heremans, K. High-pressure FTIR study of the stability of horseradish peroxidase. Effect of heme substitution, ligand binding, Ca++ removal, and reduction of the disulfide bonds. Biochemistry 2003, 42, 553−561. (25) Akasaka, K.; Nagahata, H.; Maeno, A.; Sasaki, K. Pressure acceleration of proteolysis: A general mechanism. Biophysics 2008, 4, 29−32. (26) Sun, Z.; Winter, R. Temperature- and pressure-induced unfolding of α-chymotrypsin. In Advances in High Pressure Bioscience and Biotechnology II; Winter, R., Ed.; Springer: Berlin, 2003; pp 117− 120. (27) Koo, J.; Czeslik, C. Volume changes of proteins adsorbed on silica particles. Soft Matter 2012, 8, 11670−11676. (28) Devineau, S.; Zanotti, J.-M.; Loupiac, C.; Zargarian, L.; Neiers, F.; Pin, S.; Renault, J. P. Myoglobin on silica: A case study of the impact of adsorption on protein structure and dynamics. Langmuir 2013, 29, 13456−13472. (29) Byler, D. M.; Susi, H. Examination of the secondary structure of proteins by deconvolved FTIR spectra. Biopolymers 1986, 25, 469− 487. (30) Pappa, H. S.; Cass, A. E. G. A step towards understanding the folding mechanism of horseradish peroxidase. Eur. J. Biochem. 1993, 212, 227−235. (31) Malinin, A. S.; Rakhnyanskaya, A. A.; Bacheva, A. V.; Yaroslavov, A. A. Activity of an enzyme immobilized on polyelectrolyte multilayers. Polymer Sci. 2011, 53, 52−56. (32) Blow, D. M. Structure and mechanism of chymotrypsin. Acc. Chem. Res. 1976, 9, 145−152. (33) Northrop, D. B. Effects of high pressure on enzymatic activity. Biochim. Biophys. Acta 2002, 1595, 71−79.
above about 500 bar, very small activation volumes are only observed. Although molecular interpretations of the activation volumes determined in this study are difficult, the obtained volume profiles provide some framework for mechanistic discussions. The observed pressure-dependence of the activation volumes of α-CT and HRP suggest that the transition state of HRP is more compressed under pressure than the ES complex, whereas the ES complex of α-CT is more compressed under pressure than the transition state. Apparently, the transition state of α-CT is rather rigid and cannot easily be compressed, whereas the transition state of HRP exhibits some void volume as compared to the initial ES complex.
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ASSOCIATED CONTENT
S Supporting Information *
Supporting Information is given on the calculation of the surface coverage of the silica particles by enzyme molecules, the activity measurements of the filtrates after enzyme adsorption on silica particles (Figure S1), the pressure dependence of the fluorescence intensity of the assay products (Figure S2), and the Trp fluorescence of α-CT as a function of pressure (Figure S3) and temperature (Figure S4). This material is available free of charge via the Internet at http://pubs.acs.org.
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AUTHOR INFORMATION
Corresponding Author
*E-mail [email protected]. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This work has been supported by the Deutsche Forschungsgemeinschaft (DFG). REFERENCES
(1) Talbert, J. N.; Goddard, J. M. Enzymes on material surfaces. Colloids Surf., B 2012, 93, 8−19. (2) Mateo, C.; Palomo, J. M.; Fernandez-Lorente, G.; Guisan, J. M.; Fernandez-Lafuente, R. Improvement of enzyme activity, stability and selectivity via immobilization techniques. Enzyme Microb. Technol. 2007, 40, 1451−1463. (3) Rabe, M.; Verdes, D.; Seeger, S. Understanding protein adsorption phenomena at solid surfaces. Adv. Colloid Interface Sci. 2011, 162, 87−106. (4) Czeslik, C.; Royer, C.; Hazlett, T.; Mantulin, W. Reorientational dynamics of enzymes adsorbed on quartz: A temperature-dependent time-resolved TIRF anisotropy study. Biophys. J. 2003, 84, 2533−2541. (5) Royer, C. A. Revisiting volume changes in pressure-induced protein unfolding. Biochim. Biophys. Acta 2002, 1595, 201−209. (6) Boonyaratanakornkit, B. B.; Park, C. B.; Clark, D. S. Pressure effects on intra- and intermolecular interactions within proteins. Biochim. Biophys. Acta 2002, 1595, 235−249. (7) Eisenmenger, M. J.; Reyes-De-Corcuera, J. I. High-pressure enhancement of enzymes: A review. Enzyme Microb. Technol. 2009, 45, 331−347. (8) Mozhaev, V. V.; Heremans, K.; Frank, J.; Masson, P.; Balny, C. Exploiting the effects of high hydrostatic pressure in biotechnological applications. Trends Biotechnol. 1994, 12, 493−501. (9) Butz, P.; Greulich, K. O.; Ludwig, H. Volume changes during enzyme reactions: Indications of enzyme pulsation during fumarase catalysis. Biochemistry 1988, 27, 1556−1563. (10) Anese, M.; Nicoli, M. C.; Dall’Aglio, G.; Lerici, C. R. Effect of high pressure treatments on peroxidase and polyphenoloxidase activities. J. Food Biochemistry 1995, 18, 285−293. 15502
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(34) Low, P. S.; Somero, G. N. Activation volumes in enzymatic catalysis: Their sources and modification by low-molecular-weight solutes. Proc. Natl. Acad. Sci. U. S. A. 1975, 72, 3014−3018. (35) Northrop, D. B. Unusual origins of isotope effects in enzymecatalysed reactions. Philos. Trans. R. Soc. B 2006, 361, 1341−1349. (36) Welinder, K. G. Amino acid sequence studies of horseradish peroxidase. Eur. J. Biochem. 1979, 96, 483−502. (37) Veitch, N. C. Horseradish peroxidase: A modern view of a classic enzyme. Phytochemistry 2004, 65, 249−259. (38) Rodríguez-López, J. N.; Lowe, D. J.; Hernández-Ruiz, J.; Hiner, A. N. P.; García-Cánovas, F.; Thorneley, R. N. F. Mechanism of reaction of hydrogen peroxide with horseradish peroxidase: Identification of intermediates in the catalytic cycle. J. Am. Chem. Soc. 2001, 123, 11838−11847. (39) Zheng, D.; Lu, H. P. Single-molecule enzymatic conformational dynamics: spilling out the product molecules. J. Phys. Chem. B 2014, 118, 9128−9140. (40) Gorris, H. H.; Walt, D. R. Mechanistic aspects of horseradish peroxidase elucidated through single-molecule studies. J. Am. Chem. Soc. 2009, 131, 6277−6282. (41) Smeller, L.; Fidy, J. The enzyme horseradish peroxidase is less compressible at higher pressures. Biophys. J. 2002, 82, 426−436.
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Activation Volumes of Enzymes Adsorbed on Silica Particles Vitor Schuabb and Claus Czeslik* Department of Chemistry and Chemical Biology, TU Dortmund University, D-44221 Dortmund, Germany S Supporting Information *
ABSTRACT: The immobilization of enzymes on carrier particles is useful in many biotechnological processes. In this way, enzymes can be separated from the reaction solution by filtering and can be reused in several cycles. On the other hand, there is a series of examples of free enzymes in solution that can be activated by the application of pressure. Thus, a potential loss of enzymatic activity upon immobilization on carrier particles might be compensated by pressure. In this study, we have determined the activation volumes of two enzymes, α-chymotrypsin (α-CT) and horseradish peroxidase (HRP), when they are adsorbed on silica particles and free in solution. The experiments have been carried out using fluorescence assays under pressures up to 2000 bar. In all cases, activation volumes were found to depend on the applied pressure, suggesting different compressions of the enzyme-substrate complex and the transition state. The volume profiles of free and adsorbed HRP are similar. For α-CT, larger activation volumes are found in the adsorbed state. However, up to about 500 bar, the enzymatic reaction of α-CT, which is adsorbed on silica particles, is characterized by a negative activation volume. This observation suggests that application of pressure might indeed be useful to enhance the activity of enzymes on carrier particles.
1. INTRODUCTION Enzymes are nature’s catalysts. Their function has been optimized in a perfect way in order to catalyze chemical reactions under mild conditions, such as neutral pH-values, ambient pressure, and almost ambient temperatures. Usually, they are highly specific for a particular reaction pathway, enabling yields with high enantiomeric excess. Therefore, enzymes are very attractive for use in biotechnological, pharmaceutical, and biomedical processes. The major drawbacks of using enzymes compared to chemical catalysts are their relatively low stability and their relatively high cost. Expensive enzymes are worth being recovered from the reaction media to be reused in many cycles. Removing an enzyme from a reaction mixture can be carried out easily by filtration or centrifugation, when the enzyme is immobilized on carrier particles large enough to be retained or sedimented.1,2 However, any deviation from the protein’s natural environment, such as an aqueous−solid interface, can cause a structural change and denaturation of the protein.3 Furthermore, interactions with a solid substrate can also slow down the dynamics of a protein.4 Additionally, overall, enzymatic activity at interfaces may also be decreased by an unfavorable orientation of the enzyme molecules because the active site can be blocked either by the material surface or by neighboring enzyme molecules. To preserve enzymatic activity at aqueous−solid interfaces, different strategies have been proposed and are applied.1,2 For example, the use of hydrophilic surfaces, the incorporation of an enzyme into porous materials, and genetic engineering can be helpful to overcome limited enzymatic activity at aqueous− solid interfaces. In this study, we will explore pressure as a new tool to control and potentially enhance enzymatic activity at © 2014 American Chemical Society
aqueous−solid interfaces. Generally, the effect of pressure on enzymes is two-fold, because enzymes can be regarded as a normal protein with a folded conformation, which can be destroyed under pressure, and as a biological catalyst, whose rate can be modified under pressure. Although a folded conformation of a protein represents an almost perfect dense packing, some void volumes and cavities remain, which are filled by water upon unfolding. Thus, there is a negative volume change, ΔV, associated with the unfolding of proteins (at least at ambient temperatures).5,6 Other contributions to ΔV are discussed in the literature, such as electrostriction upon exposure of charged and polar residues to the solvent.6 Due to this volume decrease, protein unfolding and subsequent loss of enzymatic activity is often observed at pressures of several 1000 bar. In addition, the catalytic properties of enzymes have been shown to sensitively depend on pressure below the pressure of unfolding in the folded state.7−9 A transition state is favored under pressure, when it has a smaller volume than the reactants. This is expressed as negative activation volume. The activation volume, ΔV‡, is related to the change of the rate constant, k, with pressure, p, according to8,9 ⎛ ∂ ln k ⎞ ΔV ‡ ⎜ ⎟ =− RT ⎝ ∂p ⎠T
(1)
Received: September 9, 2014 Revised: November 19, 2014 Published: December 5, 2014 15496
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somewhat larger diameters have been found from dynamic light scattering experiments.17 Whereas Ludox AM silica particles have a negative surface charge, Ludox CL silica particles are coated with alumina, converting their charge from negative to positive. Enzymatic Assays. α-CT was adsorbed on silica particles by mixing stock solutions of α-CT (250 μL of a 2 mg mL−1 solution) and Ludox AM (68 μL of a 1.21 g mL−1 solution) in 682 μL buffer solution (20 mM Tris-HCl, pH = 7.8) and incubating the mixed solution at 4 °C over 3 h. The calculated surface coverage of the silica particles by α-CT is on the order of 10−2 = 1% (calculation given in the Supporting Information). To probe the enzymatic activity of free or adsorbed α-CT (in the absence or presence of silica particles), samples contained 1.4 μg mL−1 of α-CT, 20 mM Tris-HCl buffer (pH = 7.8), and 1.57 mg mL−1 EnzCheck substrate. This substrate consists of a fluorophore and a quencher moiety that are covalently connected via an amino acid chain. Upon chain cleavage by α-CT, the quencher separates from the fluorophore that is then free to emit fluorescence.18 Experiments were carried out with a K2 fluorescence spectrometer from ISS (Champaign, Illinois, USA) with a high pressure sample cell from ISS. Fluorescence was excited at 502 nm with a Xe arc lamp and detected at 528 nm in photon counting mode. All measurements were performed at 25 °C. HRP was adsorbed on silica particles by mixing stock solutions of HRP (1.1 μL of a 23 μg mL−1 solution) and Ludox CL (100 μL of a 1.23 g mL−1 solution) in 1 mL buffer solution (50 mM sodium phosphate, pH = 7.4) and incubating the mixed solution at 4 °C over 3 h. The calculated surface coverage of the silica particles by HRP is on the order of 10−7 = 10−5 % (calculation given in Supporting Information). To probe the enzymatic activity of free or adsorbed HRP (in the absence or presence of silica particles), samples contained 1.16 μg mL−1 of HRP (0.026 μM), 50 mM sodium phosphate buffer (pH = 7.4), 50 μM Amplex Red substrate, and 1 mM H2O2. HRP catalyzes the oxidation of the Amplex Red substrate to the resorufin product by H2O2. Because resorufin is fluorescent, it can be quantified easily.18 Fluorescence was also measured using the K2 fluorescence spectrometer. Fluorescence of resorufin was excited at 530 nm and detected at 585 nm. All measurements were performed at 25 °C. Degree of Adsorption. Ludox CL silica particles have a positive surface charge due to alumina coating and interact with negatively charged amino acid residues of HRP. The net charge of HRP at neutral pH-values is very close to 0.19,20 α-CT, which has a small positive net charge at neutral pH-values (the isoelectric point is at pH = 9),21,22 can bind to the negatively charged Ludox AM silica particles. To determine the degree of enzyme adsorption on the silica particles, we have used Millipore Amicon Ultra centrifugal filters (50 kDa), which can only be passed by free, nonadsorbed enzyme molecules. Silica particles with adsorbed enzyme molecules cannot pass. The filters were loaded with the mixed solutions of enzymes and silica particles. After centrifugation at 14000g for 10 min at 4 °C, the filtrates were analyzed for free enzymes. No free HRP or α-CT could be detected by UV-spectroscopy. Moreover, no enzymatic activity of HRP or α-CT could be detected in the filtrate when carrying out the enzymatic assays described above. The fluorescence intensity did not change with time upon addition of the corresponding substrate (Figure S1, Supporting Information). Thus, within the sensitivity of the applied UV and fluorescence measurements, all enzyme molecules are adsorbed in the presence of the silica particles. Data Collection and Analysis. Fluorescence intensity is proportional to the fluorophore concentration, c. Thus, the rate of product formation, dc/dt, is proportional (not equal) to the slope of the fluorescence intensity that is plotted as a function of time (Figure 2). To remove all instrumental factors affecting the fluorescence intensity, the slope observed under pressure, S, has been normalized to the slope observed under ambient pressure, S0. Each fluorescence assay started with a few minutes of data collection at 1 bar followed by a 1−2 min period, during which the pressure has been increased, and a further time period under high pressure. The ratio of the slopes measured directly before and after the pressure change can be related to the ratio of the corresponding rate constants, because the concentrations are very similar before and after the pressure change:
Positive and negative activation volumes are illustrated in Figure 1. Depending on the sign of the activation volume, enzymatic reactions can proceed faster (negative sign) or slower (positive sign) under pressure.
Figure 1. Schematic simplified volume diagrams of enzymatic reactions. The enzyme-substrate (ES) complex transforms into the transition state (TS), which finally dissociates into the product (P) and the enzyme (E). The transition state can have a higher or a lower volume than the ES complex. As a consequence, the enzymatic reaction is inhibited or accelerated under pressure, respectively. The volume change from the ES complex to the transition state is called the activation volume, ΔV‡.
Here, we have studied two enzymes, horseradish peroxidase (HRP) and α-chymotrypsin (α-CT). A peroxidase catalyzes the oxidation of a substrate by H2O2. In the literature, it is reported that carrot peroxidase loses activity up to 2000 bar, but shows increased activity at 5000 bar.10 However, for HRP, an inactivation has been observed under pressures up to 5000 bar, but this pressure effect seems to depend on the kind of the used substrate.11 Enhanced activity has been reported for HRP that is immobilized in a Langmuir−Blodgett film of a phospholipid.12 α-CT is hydrolyzing peptide bonds. The catalytic activity of α-CT is known to be enhanced at high pressures.13,14 For example, an increase in pressure to 4700 bar at 20 °C results in an acceleration of the hydrolysis by a factor of 6.5.13 Furthermore, α-CT activation can also be achieved by simply adding a polyelectrolyte that is oppositely charged as compared with the substrate.15 When α-CT adsorbs on oppositely charged particles, positively charged substrates are preferred demonstrating the importance of electrostatic interactions for the substrate binding and the release of product.16 Thus, HRP and α-CT appear to be interesting candidates to study the combined effects of an aqueous-solid interface and pressure. As we will show here, pressure has a pronounced effect on enzymatic activities at interfaces and might even be used to enhance activities of immobilized enzymes. Furthermore, we present activation volumes of adsorbed enzymes for the first time, which provide valuable information about the enzymatic mechanisms.
2. MATERIALS AND METHODS Enzymes and Substrates. HRP and Amplex Red substrate (10acetyl-3,7-dihydroxyphenoxazine) were purchased from Thermo Fisher Scientific (Assay Kit A22188). α-CT from bovine pancreas was purchased from Sigma-Aldrich (C7762). The EnzCheck Peptidase/Protease Assay Kit from Thermo Fisher Scientific (Assay Kit E33758) was used to probe the α-CT activity. Ludox AM and Ludox CL colloidal silica particles were purchased from Sigma-Aldrich and used for immobilization of the enzymes. They have an average diameter of about 12 nm according to the manufacturer, although 15497
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added to the sample as an internal pressure sensor. The shift of the 983 cm−1 band to higher wavenumbers is proportional to the applied pressure.23 All spectra are corrected for background measured with buffer solution. D2O was used as the solvent.
3. RESULTS AND DISCUSSION 3.1. Pressure Stability of HRP and α-CT. In this study, we have performed enzymatic assays under pressures up to 2000 bar at 25 °C. It is helpful to separate pressure effects on the enzyme kinetics from pressure effects on the protein conformation. Free HRP and α-CT do not unfold in solution in this pressure range.24−26 However, in the adsorbed state, the conformational stability of proteins is usually lowered. This means that the temperature and the pressure of protein unfolding are significantly lowered after adsorption at an aqueous−solid interface.17,27,28 Thus, we have analyzed the conformation of HRP and α-CT when they are adsorbed on silica particles and, for comparison, free in solution as a function of pressure. Using FTIR spectroscopy in combination with a diamond anvil cell, the amide I′ band (in D2O) has been recorded. This band is known to be sensitive for the secondary structure of a protein. Any change in the secondary structure leads to a change of the band shape, because the amide I′ band is a superposition of the infrared absorption bands of the various secondary structure elements, which have all individual wavenumbers.29 In Figure 3, the amide I′ bands of HRP and αFigure 2. Typical fluorescence intensity data as recorded to probe the enzymatic activities of α-CT and HRP. The fluorescence intensity has been measured at 1 bar and after pressure increase. The slope of the data is proportional to the rate of the enzymatic reaction. The slopes before and after the pressure increase have been determined by linear fits.
S k = S0 k0
(2)
The specific meaning of the rate constants is discussed below. At 1 bar, the slope, S0, was found to be absolutely constant over time in both enzyme assays, indicating that the enzymes were saturated with substrates and were working with maximum rates. Thus, S/S0 = 1 at 1 bar for both enzymes. It follows from eq 1 that a plot of ln(S/S0) as a function of pressure, p, yields a curve whose slope contains the activation volume:
d ln(S /S0) ΔV ‡ =− dp RT
(3)
This equation can be applied to all pressures, p, and does not require a constant activation volume. When the activation volume is pressure dependent, a change in the slope of ln(S/S0) versus p is observed. In addition, the raw slope, as taken from the fit under high pressure (Figure 2), has been corrected for the pressure dependence of the fluorescence quantum yield of the product. This dependence has been measured using an enzymatic assay with free HRP or α-CT (without silica particles) that was allowed to run for about 5 h until the fluorescence intensity of the product reached a constant value, and no additional product was formed anymore. Then, the steady-state fluorescence intensity of this assay was measured as a function of pressure. After normalization to 1 bar (Figure S2), the steady-state fluorescence intensity, I(p), was used to correct the raw slope, Sraw(p), by calculating S(p) = Sraw(p)/I(p). Fourier Transform Infrared (FTIR) Spectroscopy. Infrared absorption spectra of HRP and α-CT were recorded using a Nicolet 6700 Fourier transform infrared spectrometer from Thermo Fisher Scientific at a spectral resolution of 2 cm−1. Sample solutions were analyzed in a thermostated diamond anvil cell. Barium sulfate was
Figure 3. Amide I′ band in the FTIR spectra of free and adsorbed αCT and HRP (selected data). Each diagram contains three overlapping amide I′ bands in the pressure range 1−2300 bar indicating the pressure stability of the secondary structures. Data curves refer to 1 bar (solid line), 1000 bar (dashed line), and 2300 bar (dotted line).
CT both in the adsorbed state on silica particles and free in solution are plotted. Apparently, no significant change of the band shape of the amide I′ bands can be observed, when the pressure is increased up to 2 kbar, indicating stable secondary structures of the enzymes. Indeed, the pressure of unfolding has been reported to be 12 kbar in the case of HRP24 and about 4 kbar in the case of α-CT.25,26 In addition, we have also probed 15498
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the tertiary structure of α-CT by recording the Trp fluorescence band (unfortunately, the Trp fluorescence of HRP was too weak to observe due to intramolecular Trp heme energy transfer30). A red shift of the Trp fluorescence band indicates an exposure of buried Trp residues to water in the course of protein unfolding. As can be seen in Figure S3, the Trp bands of free and adsorbed α-CT do not shift with pressure, clearly indicating an intact tertiary structure up to 2000 bar. For comparison, the Trp band of α-CT has also been measured as a function of temperature (Figure S4). Here, a pronounced red-shift of this band is observed upon thermal unfolding. Thus, pressure-induced changes of the enzyme kinetics of HRP and α-CT (see below) are not related to any pressure-induced partial unfolding of these enzymes. 3.2. Pressure Effects on the Enzymatic Activity of Free and Adsorbed α-CT. α-CT from bovine pancreas has a molar mass of about 25 000 g mol−1 and an isoelectric point of about 9 giving a positive net charge to the enzyme at neutral pHvalues.21,22 However, α-CT binds to and is active on both negatively and positively charged surfaces of polyelectrolyte layers.,31 and it binds to silica particles in a quantitative way (see Materials and Methods section). α-CT cleaves peptide bonds, preferentially those formed by the aromatic amino acids Trp, Tyr, and Phe.32 It hydrolyzes a peptide bond via an initial nucleophilic attack of the residue Ser195, which is located in the active site. The mechanism is of Michaelis−Menten type:32 k1
Figure 4. Pressure dependence of the enzymatic activity of free and adsorbed α-CT. The slopes of the plots are determined by linear fits. They are proportional to the negative activation volume according to eq 3. Each error bar covers the results of two independent measurements. S and S0 are the slopes of the fluorescence intensity as a function of time under high and ambient pressure, respectively.
α ‐CT + S ⇀ α ‐CT·S ↽⎯⎯⎯ k−1
k2
→ P2−α‐CT + P1 k3
⎯⎯⎯→ α ‐CT + P1 + P2 H 2O
Here, S is the substrate, which is hydrolyzed into the products P1 and P2. As described in the Materials and Methods section, we used a large excess of substrate (a few mg mL−1) that saturates the enzyme (a few μg mL−1). In this case, the rate of the enzymatic reaction is determined by step 2 and step 3 with a rate constant of kcat = k2k3/(k2 + k3).9 It has been proposed that k2 is small and rate-limiting for a peptide hydrolysis (this study), whereas k3 is small and rate-limiting for an ester hydrolysis.13,14,32 A back reaction is also avoided by a high substrate concentration. Therefore, the ΔV‡-values of α-CT, as observed in this study, probably refer to step 2 in the reaction scheme, where the enzyme is acylated (P2−α-CT) and an amine (P1) is released. In Figure 4a, logarithmic plots of the normalized fluorescence slopes, S/S0, are shown for α-CT free in solution. According to eq 3, the change of ln(S/S0) with pressure, p, is proportional to the negative activation volume, −ΔV‡. Apparently, the logarithmic plot is not consistent with a constant activation volume over the whole pressure range studied. Indeed, pressure-dependent activation volumes are described as the more usual case.33 The activation volume is the difference of the volumes of the transition state and the enzyme−substrate (ES) complex, ΔV‡ = V‡ − VES. If these volumes, V‡ and VES, are compressed to different extents, a change of ΔV‡ with pressure is the result. In our case, the curved data in Figure 4a can be approximated with two linear ranges from 1 to 500 bar and from 500 to 2000 bar. These linear ranges yield activation volumes of −67 mL mol−1 (1−500 bar) and −15 mL mol−1 (500−2000 bar) using eq 3.
A negative activation volume indicates that the volume of the transition state is smaller than the volume of the ES complex. In this way, high pressure favors the formation of the transition state, because the activation free energy is lowered by pΔV‡. The outcome is a faster enzymatic reaction. Negative activation volumes of α-CT have also been found in the literature.13,14 Taniguchi et al. report activation volumes in the range from −4.4 to −35 mL mol−1 for the enzyme deacylation (step 3 in the reaction scheme).14 Mozhaev et al. studied the acylation of the enzyme (step 2 in the reaction scheme) and observed an activation volume of about −10 mL mol−1 at 20 °C and −25 mL mol−1 at 50 °C.13 These values are in good agreement with our value of −15 mL mol−1 at 25 °C above 500 bar. However, the strongly negative activation volume observed in this study below 500 bar has not been resolved so far. The corresponding data of the α-CT catalysis in the adsorbed state on silica particles are shown in Figure 4b. Remarkably, as found for free α-CT, α-CT that is adsorbed on silica particles has a nonconstant activation volume in the pressure range of 1−2000 bar. There is a slight increase of the enzymatic rate up to 500 bar reaching a maximum at about 750 bar. At higher pressures, the enzymatic rate is decreasing with increasing pressure. From linear fits, activation volumes of −4 mL mol−1 (1−500 bar) and +5 mL mol−1 (1000−2000 bar) can be determined (Figure 4b). When comparing the data of free α-CT (Figure 4a) with those of adsorbed α-CT (Figure 4b), similar pressure profiles can be observed except for an overall reduction of the slopes and a corresponding increase of the activation volumes upon adsorption of α-CT on silica 15499
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enzyme surface enclosing the active site is binding to the silica surface, volume changes occurring during the enzymatic reaction are likely to be distorted. This aspect must also be considered in the interpretation of ΔV‡, and a single pathway might not be enough in the adsorbed state of the enzyme. On the other hand, conformational changes and a lowered thermodynamic stability of the folded state of the adsorbed enzyme do not play a significant role, as can be concluded from the measured FTIR (Figure 3) and fluorescence spectra (Figure S3), which are discussed in section 3.1. However, a clear molecular picture of the immobilization and pressure effects on the α-CT catalysis is not possible at this point. The effect of pressure on the activity of free and adsorbed αCT was found to be partially reversible only. After pressure release from 1000 to 1 bar, the fluorescence intensity was not linear with time, but was curved downward. This suggests a reduced activity due to substrate depletion. Recovered activities are ranging around 73% (free α-CT) and 92% (adsorbed αCT). 3.3. Pressure Effects on the Enzymatic Activity of Free and Adsorbed HRP. For comparison, we have also studied the effect of pressure on the catalytic activity of free and adsorbed HRP. HRP has a molar mass of about 44 000 g mol−1, is mainly α-helical, and binds heme as prosthetic group.36,37 HRP catalyzes the oxidation of a substrate by H2O2. The mechanism is based on the initial binding of an oxygen atom from H2O2 to the iron atom of heme. Then, two electrons are transferred successively from the substrate to the heme, and a water molecule is finally released after uptake of two protons:37,38
particles. Apparently, adsorption adds a positive contribution (in a mathematical sense) to the activation volume. A volume diagram of α-CT catalysis is given in Figure 5. Activation volumes are illustrated as arrows pointing from the
Figure 5. Activation volumes of free and adsorbed α-CT (not to scale, ΔV‡ given in mL mol−1). In both cases, the activation volume depends on the pressure. The ES complex is compressed in a stronger way than the transition state (TS). The volume levels at low and high pressures are arranged assuming a volume reduction of both ES and TS by pressure.
ES complex to the transition state. In the literature, sources of activation volumes are discussed.6−8,34,35 Exposure of hydrophobic and charged amino acid residues to water during the formation of the transition state will cause a higher hydration density and a negative activation volume. Indeed, Mozhaev et al. have attributed the observed negative activation volume of αCT to the increased hydration of the transition state dipole.13 Moreover, small conformational changes of the enzyme forming the transition state can be linked to the generation or the filling of void volumes inside the enzyme. In both cases of free and adsorbed α-CT, we observe a more positive activation volume in the higher pressure range. This may be explained by a stronger compression of the ES complex relative to the transition state, as illustrated in the volume diagram (Figure 5). If VES is compressed in a stronger way than V‡, the activation volume, which is the difference V‡ − VES, becomes larger (more positive). From conformational studies it is clear that α-CT does not undergo significant changes in the secondary and tertiary structure in the pressure range of 1− 2000 bar (Figure 3 and Figure S3). However, we could speculate that the substrate binding in the ES complex leaves some void volume at ambient pressure. At about 500 bar, the substrate molecule might be pushed more deeply into the pocket of the enzyme in a way that substrate-enzyme fitting is optimized and VES becomes smaller. Upon adsorption of α-CT on the silica particles, the activation volumes are increasing (Figure 5). It is likely that an increase of the transition state volume, V‡, is the reason for this finding. When an enzyme is catalyzing a chemical reaction, conformational flexibility of the enzyme is mandatory. However, in the adsorbed state, this conformational flexibility will be reduced. As a consequence, the transition state might have a nonperfect, distorted geometry with an increased void volume, which increases V‡ and ΔV‡ of α-CT in the adsorbed state. Moreover, the location of water molecules in the active site of the enzyme appears to be a crucial factor. There might be gaps in the transition state that are just too small to accommodate a water molecule. There is also a heterogeneous orientation of the enzyme molecules on the silica particles. Thus, there are different enzyme−silica interactions that could lead to different volume profiles. In particular, when the
This mechanism is much more complex than the peptide cleavage by α-CT. The two intermediates in the above scheme are denoted as compound I and compound II. In this study, HRP catalyzes the oxidation of the Amplex Red substrate to the resorufin product by hydrogen peroxide. Resorufin is detected by its fluorescence (see Materials and Methods section). In the enzymatic assay, we have used a large excess of hydrogen peroxide (1 mM) and Amplex Red (50 μM) that bind to HRP (0.026 μM). This excess disfavors the back reaction and ensures the maximum enzymatic rate. The rate-limiting steps will then be the successive transfers of two electrons from an Amplex Red molecule to the heme group. Recent single molecule fluorescence experiments are consistent with a two-electron mechanism, where the two electrons originate from a single Amplex Red molecule and fluorescent resorufin is formed while it is still confined in the enzyme.39 Alternatively, it has been proposed that HRP oxidizes an Amplex Red molecule by a oneelectron transfer only. Afterward, the formed nonfluorescent radical intermediate is released from the enzyme and reacts to resorufin by dismutation.40 The measured enzymatic activity of HRP using Amplex Red as the substrate is shown in Figure 6. The observed slope of the product fluorescence, S, is plotted as a function of pressure, p, on a logarithmic scale (see eq 3). In both cases, free HRP and HRP that is adsorbed on silica particles, the slope of the product fluorescence, and thus the associated rate constant of product formation (see eq 2) decreases as the pressure increases. This indicates positive activation volumes, which increase the activation free energies by the amount pΔV‡. As 15500
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Overall, the volume profiles of free and adsorbed HRP appear to be similar (Fig. 7). Presumably, the mechanism of the enzymatic reaction does not change upon adsorption, and no large distortion of the active site occurs when HRP interacts with the silica particles. However, in contrast to α-CT, activation volumes of HRP tend to be smaller at higher pressures. Above 500 bar, an activation volume of only +2 mL mol−1 indicates that the volumes of the transition state and the ES complex are very similar, and pressure has little effect on the enzymatic rate. Apparently, pressure can compress the transition state stronger than the ES complex leading to a reduction of the activation volume, ΔV‡ = V‡ − VES. So far, a molecular interpretation of the volume diagrams given in Figure 7 is difficult. Assuming the transfers of the two electrons from Amplex Red to HRP are rate-limiting, it can be concluded that the ES complex grows in volume to form the transition state, where the electron transfer can happen, because V‡ > VES. Probably, electron transfer requires some shift of Amplex Red away from the optimum binding position thereby creating some void volume. However, at pressures above about 500 bar, the increased volume of the transition state is suppressed resulting in V‡ ≈ VES. This finding is supported by earlier compressibility measurements of HRP in the low and high pressure range.41 Below 100 bar, HRP has a large compressibility, whereas HRP is less compressible at higher pressures. It has been concluded that the enzyme has soft domains at ambient conditions, which are compressed and become rigid at higher pressures.41 Thus, a larger compressibility of HRP at ambient conditions is linked to a larger density fluctuation, which could be the reason for the larger activation volumes of free and adsorbed HRP that are observed below 500 bar (Figure 7). At higher pressures, a rigid compressed structure of HRP is consistent with little volume change during the oxidation of Amplex Red by HRP. The effect of pressure on the activity of free and adsorbed HRP was found to be partially reversible only. After pressure release from 1000 to 1 bar, the fluorescence intensity was not linear with time, but was curved upward. This suggests a slow recovery of the original conformation. Recovered activities range around 90%.
Figure 6. Pressure dependence of the enzymatic activity of free and adsorbed HRP. The slopes of the plots are determined by linear fits. They are proportional to the negative activation volume according to eq 3. Each error bar covers the results of four independent measurements. S and S0 are the slopes of the fluorescence intensity as a function of time under high and ambient pressure, respectively.
found for α-CT, the plots of ln(S/S0) versus p (Figure 6) are nonlinear, suggesting nonconstant activation volumes of free and adsorbed HRP in the pressure range 1−2000 bar. To estimate activation volumes, we have divided the studied pressure range into a lower and higher pressure range in a similar way as described above for α-CT. From the linear fits (Figure 6) and eq 3, we can derive activation volumes of +16 mL mol−1 (1−500 bar) and +2 mL mol−1 (500−2000 bar) in the case of free HRP. For adsorbed HRP, corresponding values of +27 mL mol−1 (1−250 bar) and +2 mL mol−1 (250−1500 bar) are obtained (note that the error bars in Figure 6 are covering the results of four independent measurements). All activation volumes obtained in this way are summarized in a volume diagram (Figure 7).
4. CONCLUSIONS The effect of pressure on the enzymatic activity of free and adsorbed α-CT and HRP has been studied. α-CT cleaves a peptide bond, whereas HRP oxidizes a substrate. Generally, the immobilization of enzymes on carrier particles provides an easy method to remove expensive enzymes from reaction media for the reuse in additional processes. Because enzymes can lose some biological activity at aqueous−solid interfaces, it is interesting to study pressure as a new tool to activate enzymes in the adsorbed state. In the case of free α-CT, it is known that the enzymatic rate is increasing with increasing pressure. We could confirm this effect, but in addition, we have also observed that the pressure activation of free α-CT is more pronounced in the lower pressure range up to about 500 bar. Upon adsorption on silica particles, a positive contribution is added to the activation volume of α-CT. However, up to 500 bar, the activation volume of adsorbed α-CT is still negative, and the enzymatic activity can still be enhanced by the application of pressure. This finding demonstrates a new potential strategy to optimize enzymatic reactions on carrier particles. On the other hand, the observed activation volumes of HRP are generally positive and similar in the free and adsorbed states. However,
Figure 7. Activation volumes of free and adsorbed HRP (not to scale, ΔV‡ given in mL mol−1). In both cases, the activation volume depends on the pressure. The TS is compressed in a stronger way than the ES complex. The volume levels at low and high pressures are arranged assuming a volume reduction of both ES and TS by pressure. 15501
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above about 500 bar, very small activation volumes are only observed. Although molecular interpretations of the activation volumes determined in this study are difficult, the obtained volume profiles provide some framework for mechanistic discussions. The observed pressure-dependence of the activation volumes of α-CT and HRP suggest that the transition state of HRP is more compressed under pressure than the ES complex, whereas the ES complex of α-CT is more compressed under pressure than the transition state. Apparently, the transition state of α-CT is rather rigid and cannot easily be compressed, whereas the transition state of HRP exhibits some void volume as compared to the initial ES complex.
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ASSOCIATED CONTENT
S Supporting Information *
Supporting Information is given on the calculation of the surface coverage of the silica particles by enzyme molecules, the activity measurements of the filtrates after enzyme adsorption on silica particles (Figure S1), the pressure dependence of the fluorescence intensity of the assay products (Figure S2), and the Trp fluorescence of α-CT as a function of pressure (Figure S3) and temperature (Figure S4). This material is available free of charge via the Internet at http://pubs.acs.org.
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AUTHOR INFORMATION
Corresponding Author
*E-mail [email protected]. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This work has been supported by the Deutsche Forschungsgemeinschaft (DFG). REFERENCES
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