On the activity loss of hydrolases in organic solvents: II. a mechanistic study of subtilisin Carlsberg
© Castillo et al; licensee BioMed Central Ltd. 2006
Received: 10 August 2006
Accepted: 22 December 2006
Published: 22 December 2006
Enzymes have been extensively used in organic solvents to catalyze a variety of transformations of biological and industrial significance. It has been generally accepted that in dry aprotic organic solvents, enzymes are kinetically trapped in their conformation due to the high-energy barrier needed for them to unfold, suggesting that in such media they should remain catalytically active for long periods. However, recent studies on a variety of enzymes demonstrate that their initial high activity is severely reduced after exposure to organic solvents for several hours. It was speculated that this could be due to structural perturbations, changes of the enzyme's pH memory, enzyme aggregation, or dehydration due to water removal by the solvents. Herein, we systematically study the possible causes for this undesirable activity loss in 1,4-dioxane.
As model enzyme, we employed the protease subtilisin Carlsberg, prepared by lyophilization and colyophilization with the additive methyl-β-cyclodextrin (MβCD). Our results exclude a mechanism involving a change in ionization state of the enzyme, since the enzyme activity shows a similar pH dependence before and after incubation for 5 days in 1,4-dioxane. No apparent secondary or tertiary structural perturbations resulting from prolonged exposure in this solvent were detected. Furthermore, active site titration revealed that the number of active sites remained constant during incubation. Additionally, the hydration level of the enzyme does not seem to affect its stability. Electron paramagnetic resonance spectroscopy studies revealed no substantial increase in the rotational freedom of a paramagnetic nitroxide inhibitor bound to the active site (a spin-label) during incubation in neat 1,4-dioxane, when the water activity was kept constant using BaBr2 hydrated salts. Incubation was also accompanied by a substantial decrease in Vmax/KM.
These results exclude some of the most obvious causes for the observed low enzyme storage stability in 1,4-dioxane, mainly structural, dynamics and ionization state changes. The most likely explanation is possible rearrangement of water molecules within the enzyme that could affect its dielectric environment. However, other mechanisms, such as small distortions around the active site or rearrangement of counter ions, cannot be excluded at this time.
Enzymes have been successfully employed to catalyze a number of transformations and chiral resolutions of biological and industrial importance in organic solvents [1–10]] It is well documented that enzyme catalysis in these non-natural media offers some advantages over that in natural-aqueous environment, such as: prevention of autolysis (in case of proteases) and increased thermostability [11, 12]. In fact, it has been shown that enzymes can perform catalysis in organic solvents at temperatures far above those temperatures that denature enzymes in aqueous systems . Despite this, enzyme inactivation in organic solvents has been reported, and studies have addressed this issue. For example, Fagain et al. (2003) have recently reviewed bioreactor stability, shelf life and operational stability of a variety of enzymes suspended in neat organic solvents and aqueous-organic solvent mixtures .
We have previously reported that the activity of the serine protease subtilisin Carlsberg is significantly reduced upon storage in several organic solvents, irrespective of its preparation, hydration, hydrophobicity of the solvent, the reaction temperature and of the substrates used . A subsequent study using various lipases, an esterase, and α-chymotrypsin showed that for these enzymes most of their initial high activity also plunges upon exposure to organic solvents . It was found that some additives and modes of enzyme preparation (such as chemical modification with polyethylene glycol) were able to reduce activity loss of the enzymes in various organic solvents. The other variables studied (enzyme hydration, the reaction temperature, the organic solvent and the enzymes themselves) were not found to affect the rapid loss of activity observed. Furthermore, experiments using structurally defined cross-linked enzyme crystals (CLECs) [11, 16] suggested that the mechanism of inactivation might not involve large structural changes of the enzyme catalyst. The activity loss was reversible upon re-lyophilizing the enzyme from an aqueous buffer, indicating that inactivation did not involve autolysis or irreversible denaturation of the enzyme .
Those findings outline a clear drawback in the use of biocatalysts for applications that require prolonged exposure to organic solvents, and showed the need for understanding the mechanism of enzyme inactivation during storage in these media. Herein we study this detrimental effect in detail using two different preparations of the model enzyme subtilisin Carlsberg.
The results previously reported from this and other laboratories demonstrated that most enzymes lose their initial high activity exponentially during the first hours of incubation in a variety of organic solvents until a constant activity value is reached. So far, all of the enzymes studied show some residual activity after a long incubation period (usually from 4 to 7 days), which appears to persist indefinitely [14, 15, 17]. However, no mechanistic explanation for this phenomenon has been provided so far. Herein we examine the structure, the flexibility, the number of active sites remaining, the powder morphology, and the pH profile of subtilisin Carlsberg as some of the possible causes that could contribute to the observed decrease in enzyme activity after prolonged exposure to organic media. We had previously demonstrated that subtilisin Carlsberg instability was independent of: (a) the mode of enzyme preparation (lyophilized, co-lyophilized and CLECs – all showed a similar stability profile); (b) the physicochemical properties of the solvents used; and (c) the substrates . Subsequent studies involving different preparations of α-chymotrypsin, pig-liver esterase and several lipases showed that this was not unique to subtilisin, but rather a common effect shared by several enzymes .
First, we decided to study the enzyme-powder morphology after suspension in 1,4-dioxane since it has been suggested that incubation could result in larger and more compact aggregates, which could reduce the enzyme activity.
Active-site titration and Michaelis constant
Castillo et al., Table 1 Kinetic parameters of subtilisin C. co-lyophilized with methyl-β-cyclodextrin
102 ± 3
16 ± 2
130 ± 10
319 ± 48
Enzyme pH profile
Active site spin labeling
Equation 1: τ(s) = 6.5 × 10-10 ΔH0 [(h0/h-1)1/2-1]
EPR studies of subtilisin C. in 1,4-dioxane with 0.5 % water and controlling the water activity
Spin-labeled lyophilized subtilisin C. was suspended in 1,4-dioxane, water (0.5 % v/v) was added, and the EPR spectra were recorded at day 0 (no incubation) and day 4 of storage in this solvent. The results show that in the presence of water, at day zero (no incubation), the spin label has a higher degree of mobility than that of the enzyme suspended in the neat solvent (ΔH 0 is smaller while h i /(h i +h a ) is larger – consistent with increased mobility of the spin label, (Figure 7)). Incubation of the hydrated enzyme for 5 days in this solvent shows only a moderate increase in the spin label mobility, as determined from the values of Tau, ΔH 0 and h i /(h i +h a ). However, when these experiments were repeated controlling the water activity, no increase in enzyme flexibility was observed (Figure 7). Since hydration has a clear effect on the enzyme flexibility, and it has been well documented that added water can alter the enzyme activity, we decided to look at the effect of hydration on enzyme storage stability further.
Enzyme storage stability in the presence of 0.5% water and controlling the water activity
It is clear that loss of activity is not due to any major change in structure. Other possible mechanisms include changes in the ionization state of critical residues in the enzyme, changes in the enzyme flexibility, or in the enzyme's hydration level which could also have an effect on its flexibility. Our stability data obtained with 0.5% water and with controlled water activity rules out hydration of the enzyme as the cause. Change in hydration is one factor that may affect both activity and flexibility without significant change in protein structure. Our experiments varying the water content of the organic solvent in which the enzyme is incubated are informative here. The quite different changes in flexibility seen with different water levels suggest that hydration is a major contributor to these effects.
In contrast, the much smaller effects of water level on inactivation over time suggest that this process is not so closely related to hydration (or indeed to flexibility changes). However, we certainly cannot exclude processes involving protein-bound water molecules. Activity could be affected by the loss or movement of just one or two water molecules close to the active site. Such changes might even be opposite to those in the total amount of enzyme-bound water.
The pH memory experiments tend to exclude changes in the ionization state as the cause, but the increase in KM observed could be the result of small but critical changes in the structure around the active-site. It is also interesting to compare these findings with the inactivation of subtilisin Carlsberg, in silica immobilised or CLEC form, during continuous operation in acetonitrile based media . In this case no activity can be recovered on returning the catalyst to aqueous media , and CD, infrared and fluorescence measurements reveal extensive structural changes (Ganesan et al (2006), and further unpublished work). Clearly the mechanisms involved here are quite different than the storage stability studied in the present paper, perhaps the immobilized enzyme used in that study is more susceptible to structural or flexibility changes than the "native" enzyme?
Our studies of secondary and tertiary structure show that the enzyme remains structurally defined during prolonged incubation in organic solvents. Changes in the enzyme hydration or aggregation do not seem to be responsible for the observed poor storage stability, although it is possible that in the presence of water, we could be observing two competing effects (activation and fast deactivation) which prevent us from clearly separating the two effects (low storage stability from a deactivation/activation effect of added water). The flexibility of the active-site spin label increases with increased water concentration, but it remains constant during the storage period when the water activity is controlled by added hydrated salts. The observed decrease in activity is also accompanied by an increase in KM and a decrease in Vmax, showing loss of the catalytic efficiency of the enzyme.
These results exclude structural changes, flexibility, hydration, and changes of the enzyme ionization state as possible mechanism for the observed low storage stability, but do not exclude possible depletion or rearrangement of water molecules around the active site, or small structural perturbations around the active site or movements of counter ions. Further comprehensive studies will be required to ascertain this.
Enzymes and reagents
Subtilisin Carlsberg (alkaline protease from Bacillus licheniformis, EC 220.127.116.11) and sec-phenethyl alcohol were purchased from Sigma/Aldrich (St. Louis, MO). The solvents were purchased from Aldrich in the anhydrous form (Sure/Seal bottles, water content below 0.005%). Vinyl butyrate was purchased from TCI America (Portland, OR). The spin label 4-ethoxyfluorophosphinyloxy-TEMPO (4-EFPO-TEMPO) was purchased from Toronto Research Chemicals (North York, Canada).
Enzyme preparation and kinetic measurements
Subtilisin C. powder was prepared by 24 h lyophilization from a solution of 5 mg/mL in a 20 mM potassium phosphate buffer at pH 7.8. Colyophilization with MβCD was done by lyophilizing the enzyme from a buffer solution containing the additive at a 1:6 weight ratio of enzyme to additive. The model transesterification reaction between sec-phenethyl alcohol and vinyl butyrate was followed by gas chromatography. The GC instruments (HP 6850 and HP 6890, fitted with Chirasil CB columns, FID detectors and He as carrier gas) were calibrated with the chiral ester products of the reactions. The product peak areas and retention times were the same in the presence or absence of the substrates. The substrates (70 mM alcohol and 200 mM vinyl butyrate) and the solvent (1.0 mL) were stored over activated 3 Å molecular sieves prior to their use. All kinetic experiments were terminated before 10% of the product was formed. The enzyme enantioselectivity was determined by measuring the initial rates of enantiomeric product formation . The retention times of the "R" and "S" products were obtained using samples of the pure enantiomers synthesized from the corresponding alcohol enantiomers. The enzyme enantioselectivity is equal to the ratio: [kcat/KM]R/[kcat/KM]S = VR[S]/VS[R] .
Enzyme hydration and controlled activity experiments
Water activity was controlled using hydrated salt pairs as described . The mixture of BaBr2•1H2O/BaBr2 was prepared by placing the anhydrous salt (5 g/16.8 mmol) over pure water (75.8 mg/4.2 mmol) in a sealed vessel saturated with water vapor . The enzyme activity measurements performed at the resulting water activity of 0.008 were carried out as described above with the following modifications: The mixture of BaBr2•1H2O/BaBr2 (100 mg) was added to the solvent (1 mL) containing the enzyme and vinyl butyrate. The suspension was equilibrated by shaking at 45°C and 300 rpm for 60 min. Then, the alcohol was added and the reaction monitored as described above.
Enzyme stability in organic solvents
Enzyme stability was determined by measuring the enzyme activity (initial velocity) as a function of the incubation time (for up to 4 days) as follows: vials containing the enzyme (lyophilized powder: 10 mg, and co-lyophilized: 1.0 mg) suspended in 1.0 mL 1,4-dioxane under N2, were incubated at 45°C under constant shaking (300 rpm). To measure the enzyme activity at different incubation times, the substrates were added to initiate the reaction and the kinetics followed as previously described.
Michaelis constants (KM), Vmaxand enzyme concentration
The apparent Michaelis constants and Vmax were determined by plotting Vs vs Vs/[S] measured for the model transesterification reaction between phenyl alcohol and vinyl butyrate. The alcohol concentration was increased from 1 to 100 mM, while the activated ester concentration was kept constant at 200 mM. The active enzyme concentration was determined following a published procedure .
pH profile experiments
The enzyme was lyophilized and co-lyophilized from 20 mM potassium phosphate buffers of varying pH, adjusted as needed with dilute solutions of NaOH or HCl. The initial activities (for the transesterification reaction between sec-phenethyl alcohol and vinyl butyrate) at day 0 and after a 7-day incubation period were measured for each of the enzymes (prepared from buffers of different pH).
FTIR studies were conducted with a Nicolet Magna-IR System 560 optical bench as described [20, 28, 31, 32]. A total of 256 scans at 2 cm-1 resolution using Happ-Ganzel apodization were averaged to obtain each spectrum. Aqueous spectra were measured using spacers of <15 μm thickness. Lyophilized protein powders (lyophilized and colyophilized) were measured as KBr pellets (1 mg of protein per 200 mg of KBr). Pellets were produced using a Spectra Tech Macro-Micro KBr Die Kit and Carver 12-ton hydraulic press. Enzyme powders in organic solvents were prepared by sonication for 2 min in a sonication bath and measured in a FTIR cell equipped with CaF2 windows and using 100 μm thick spacer. When necessary, spectra were corrected for the solvent background and water vapor contributions in an interactive manner using the Nicolet OMNIC 3.1 software to obtain the protein's vibration spectra. Each protein sample was measured at least five times [20, 32].
All spectra were analyzed by second derivatization in the amide I region (1700–1600 cm-1). The second derivative spectra were obtained with the derivative function of Omnic 3.1 software (Nicolet). The final protein spectrum were smoothed with an 11-point smoothing function (10.6 cm-1). Amide I second derivative spectra were also used to calculate the spectral correlation coefficient (see below).
Calculation of the correlation coefficient
Spectral correlation coefficient values (SCC) to quantify procedure-induced protein structural perturbations were calculated as described using the amide I second derivative spectra [19, 28]. After correcting the spectra for the background and all water vapor contributions, the second-derivative spectra in the amide I spectral region were calculated and smoothed twice with a 10.6 cm-1 function. Next, these second-derivative spectra were saved for the spectral range from 1700–1600 cm-1 on identical wavenumber scales and with identical data spacing. The data for the reference spectrum (e.g. the native in aqueous medium) and the spectrum with the varied condition (e.g. after suspension in organic solvent) were imported into the program Sigma Plot and the correlation coefficient was calculated as described in detail by Griebenow et al . An SCC of 1.0 indicates that two spectra being compared are the same.
Circular Dichroism experiments
Far and near UV Circular Dichroism spectra (CD) of lyophilized and co-lyophilized subtilisin C. were studied in 1,4-dioxane, before and after incubation for 4 days as described. CD spectra were measured at room temperature in a Jasco 810 spectropolarimeter fitted with a rotating sample cell holder. Full details are described in Ganesan et al (2006). Far UV CD and near UV CD spectra were measured using cells of path length 0.01 cm and 0.5 cm, respectively. The spectra were an average of 5 scans collected at a scan speed of 50 nm/min and bandwidth 1 nm for far UV and 1.5 nm for near UV. After the measurement was completed the content of the sample cell was collected (with rinsing), air dried and dissolved in deionized water and protein concentration was estimated from UV absorption at 280 nm.
Preparation of spin labeled subtilisin
Subtilisin Carlsberg was spin labeled at the active site Ser-221 with 4-ethoxyfluorophosphinyloxy-TEMPO (Figure 8) by the method of Morrisett and Broomfield with minor modifications in the published protocol . After the reaction, the remaining free radical was removed using Sephadex G25 desalting columns (Amersham Biosciences). The spin labeled enzyme free of any unbound spin label was thereafter lyophilized and used for the EPR analysis. Active site titration using a published protocol  revealed a 100% reduction on the number of enzyme-active-sites (accompanied also by loss of enzyme activity), demonstrating that indeed, spin-labeling took place at the active-site. Organic solvents dried overnight over molecular sieves were used for the EPR analysis. The incubation of labeled enzyme in organic solvents was carried out at room temperature. All EPR spectra were recorded at room temperature on a Bruker EMX EPR spectrometer with a microwave power of 2.0 mW, a microwave frequency of 9.7 GHz and a sweep width of 100G (resolution 1024 points). Every spectrum was obtained from multiple scans.
We thank EPSRC and BBSRC (UK) for support of the CD studies. Contract grant sponsor: National Institute of Health, grant numbers: S06 GM-08216, S06 GM08102, P20 RR16439, and by NIH Grant Number P20 RR-016470 from the INBRE Program of the National Center For Research Resources.
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