Smart Core–Shell Microgel Support for Acetyl Coenzyme A

Smart Core–Shell Microgel Support for Acetyl Coenzyme A Synthetase: A Step Toward Efficient Synthesis of ... Publication Date (Web): June 18, 2014. ...
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Smart Core−Shell Microgel Support for Acetyl Coenzyme A Synthetase: A Step Toward Efficient Synthesis of Polyketide-Based Drugs Nidhi C. Dubey,†,‡ Bijay P. Tripathi,† Manfred Stamm,†,‡ and Leonid Ionov*,† †

Department of Nanostructured Materials, Leibniz Institute of Polymer Research Dresden, Hohe Str. 6, D-01069 Dresden, Germany Technische Universitat Dresden, Department of Chemistry, Dresden, 01069, Germany

‡

ABSTRACT: The flexibility in tuning the structure and charge properties of PNIPAm microgels during their synthesis makes them a suitable choice for various biological applications. Two-step free radical polymerization, a common method employed for synthesis of core−shell microgel has been well adopted to obtain cationic poly(N-isopropylacrylamide-aminoethyl methacrylate) (PNIPAm-AEMA) shell and PNIPAm core. Scanning electron microscopy (SEM), dynamic light scattering (DLS), zeta potential, and ninhydrin assay suggests nearly monodispersed particles of cationic nature. Amino groups on the microgel provides suitable attachment point for covalent immobilization of acetyl coenzyme A synthetase (Acs) via 1-ethyl-3-(3-N,N- dimethylaminopropyl) carbodiimide (EDC) chemistry. On immobilization, 61.55% of initial activity of Acs has been retained, while Michaelis−Menten kinetics of the immobilized Acs indicates identical Km (Michaelis constant) but decrease in the Vmax (maximum substrate conversion rate) compared to free enzyme. Immobilized Acs shows an improvement in activity at wide temperature and pH range and also demonstrates good thermal, storage, and operational stability. The Acs−microgel bioconjugate has been successfully reused for four consecutive operation cycles with more than 50% initial activity.

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INTRODUCTION Acetyl CoA is mentioned as the “Hub of metabolism” and is oxidized to produce energy. It plays an essential role as a donor of an acetyl group in the tricitric acid cycle, a neurotransmitter acetylcholine synthesis, and as a precursor molecule in various primary and secondary metabolic reactions such as fatty acid synthesis and polyketide synthesis.1,2 From a practical point of view, acetyl CoA is highly important for biomedical studies such as drug discovery for metabolic disorders and in biotechnological applications for the synthesis of lipids and polyketide-based anticancer drugs.3−5 Polyketides are secondary metabolites synthesized by microorganism and plants as defense molecules. The drugs like tetracycline and erythromycin (antibacterial), doxorubicin and mithramycin (anticancer), EGCG and resverastrol (antioxidants), and Zocor (drug lowering cholesterol levels) comprise this class. The annual market of polyketide-based drugs and products marks a sale of more than $35 billion. Therefore, pharmaceutical companies attach an enormous value to the polyketide compound libraries collected from bacteria, fungi, and plants, and this has led to a rapid increase in commercial biocombinatorial research activity.6−8 The high cost and demand of acetyl CoA for the synthesis of these drugs makes the process highly expensive. To overcome these limitations, different in vitro chemical- and enzyme-based acetyl CoA regeneration systems are employed with the main synthetic process.9,10 One such system is the use of enzyme acetyl CoA synthetase (Acs acetate/CoA ligase, EC © 2014 American Chemical Society

6.2.1.1), which catalyzes the synthesis of acetyl CoA from acetate. The main advantage of this enzyme is its higher substrate specificity, and the two-step reaction is carried out by a single enzyme system, as shown below:11,12 acetate + ATP → acetyl‐AMP + PPi

(1)

acetyl‐AMP + CoA → acetylCoA + AMP

(2)

In view of the low stability and highly expensive nature of enzyme catalysts, suitable approaches are needed to improve the stability as well as reusability to make the process economically viable. Immobilization of enzymes on suitable substrates is a prevailing approach to achieve relatively stable and reusable biocatalyst along with a simplified separation process.13,14 Immobilization of Acs is expected to provide a better alternative to the current methods and make the process more cost efficient and feasible. Mannens et al.15 reported Acs immobilization on glass beads for synthesis of C11 labeled acetyl CoA. The immobilization of enzyme on glass beads was screened for different cross-linking methods and spacer arms. Cyanogen bromide (CNBr) activated glass beads gave maximum enzyme load and were selected for further optimization of enzyme activity and column fabrication to Received: April 30, 2014 Revised: June 17, 2014 Published: June 18, 2014 2776

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obtain complete conversion of 1 μmol of acetate.15,16 Reports are also available where Acs was immobilized on different support materials (e.g., Nafion membrane, cellulose fibers, etc.) to study immobilization and biochemical reactions.17,18 In recent decades, a great deal of work has been focused on the preparation of core−shell particles that can adapt their behavior to changes in their environment.19 The core/shell type particles combine the properties of core (inner material) and a shell (outer layer material), which is not possessed by the individual components.20 The particles can be designed in variable shapes and sizes with combinations of different classes of inorganic/organic materials.21 Since the first report of Pelton et al.22 for the synthesis of PNIPAm microgels using precipitation polymerization, the method has been well studied and further modified to obtain microgels of the desired size, shape, and functional groups. Use of microgels as a substrate for the immobilization of the enzyme provides certain advantages. First, microgel particles can easily be synthesized using precipitation or emulsion polymerization techniques.22 Second, the desired chemical groups, suitable for immobilization of enzymes, can be easily introduced on the microgel surface during the synthesis. Third, microgels have a mass density close to water and do not precipitate as fast as SiO2 particles with a similar size, which is very important for an efficient synthetic process in reactors. Forth, due to their hydrophilic nature, microgel particles shall not cause denaturing of the enzyme, which often occurs on hydrophobic surfaces.23,24 PNIPAm microgel particles have been extensively studied due to their pronounced thermal response near physiological temperature, simplicity of synthesis, and high monodispersity. Due to their small size, PNIPAm microgels possess very quick response toward various external stimuli.25 Core−shell microgels with the shell prepared from functional monomers have the additional advantage of carrying a high density of functional groups on the surface and hence achieving a high load of molecules to be conjugated.26 Herein for the first time we report the covalent immobilization of Acs on thermoresponsive PNIPAm-AEMA core−shell microgels with the aim to obtain a reusable and stable bioconjugate catalyst (Figure 1). The cationic core−shell microgels were synthesized using two-stage precipitation polymerization and used to covalently conjugate the Acs enzyme via an amide bond between the amino group of the microgel and the carboxylic group of enzymes. The enzyme

immobilization conditions on microgel were standardized and different reaction and stability parameters were evaluated comparing with the soluble enzyme.

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EXPERIMENTAL SECTION

Materials and Methods. N-Isopropylacrylamide (97%) (NIPAm) was recrystallized from n-hexane and dried in vacuum before use. N,N′-Methylenebisacrylamide (≥99.5%; MBA), sodium dodecyl sulfate (SDS), ammonium persulfate (≥98.0%; APS), 2-aminoethyl methacrylate hydrochloride (90%; AEMA), S-acetyl-coenzyme A synthetase (EC 6.2.1.1, >3 units/mg, from baker’s yeast; Acs), 1ethyl-3-(3-N,N-dimethylaminopropyl) carbodiimide (EDC), and all other chemicals were obtained from Sigma-Aldrich (Germany). All the chemicals were used as received unless otherwise stated. The 30 and 10 kDa molecular weight cut off (MWCO) centrifugal filter tubes were purchased from Millipore. Water used throughout the investigation was purified to a resistance of 18 MΩ (Millipore) and filtered through a 0.45 μm nylon filter to remove particulate matter. Synthesis of PNIPAm-AEMA Core−Shell Microgels. The core−shell microgel particles were synthesized by a two-step free radical precipitation polymerization method, as shown in Figure 2a.27,28 A 64 mM total monomer concentration of NIPAm (700 mg) and MBA (21 mg) was taken for the synthesis of the core. While, for shell synthesis, a 37 mM solution of NIPAm (400 mg), MBA (12 mg), and AEMA (10 mg) was used. Except the initiator, all ingredients were first dissolved in water and filtered through Whatman filter paper and heated under stirring to 70 °C under a gentle stream of argon for 1 h. First the core was synthesized by rapid addition of 1 mL of APS solution (2 mM) into a monomer solution containing SDS (2 mM). The reaction was allowed to proceed for 4 h at 70 °C under continuous stirring. After polymerization, the solution was filtered through a 0.45 μm pore size filter. For synthesis of shell, SDS (0.7 mM) was added to a 20 mL solution of core microgel and the volume was made up to 75 mL using Millipore water. The solution was further heated under stirring in a nitrogen gas atmosphere to 70 °C. A total of 25 mL of monomer solution for shell synthesis was added to the heated core microgel solution, and the reaction was initiated with the addition 1 mL of APS solution (1.5 mM). The reaction mixture was continuously stirred for 4 h at 70 °C. Thus, obtained core−shell microgel solution was allowed to cool at room temperature and filtered with a suitable membrane. The unreacted monomers and small molecular weight polymers were removed from the microgel solution by continuous dialysis for a week using a 10 kDa MWCO dialysis tube against a daily change of water. Dynamic Light Scattering, Zeta Potential, and Morphology Analysis. The dynamic light scattering (DLS) measurements under different temperatures were performed for both core and core−shell microgels to analyze the particle size and its thermoresponsive behavior. Zeta potential measurements of both particles were also carried out to identify the charge behavior. Both DLS and zeta potential experiments were done using Zeta sizer Nano 3000HS (Malvern Instruments/U.K.), equipped with a 633 nm He/Ne laser and a noninvasive back scatter (NIBS) technology. Before the size measurements, samples were thermally equilibrated for 10 min and data were acquired by averaging 30 measurements, with a 10 s integrating time for each measurement. Volume phase transition temperature was determined with respect to temperature (24−40 °C). Zeta potential was obtained at pH 7, and the values were the average of three successive readings. Furthermore, the morphology of the synthesized microgel was also characterized by scanning electron microscopy. SEM was performed on vacuum-dried microgels on a silicon wafer using a NEON 40 FIB-SEM workstation (Carl Zeiss AG, Germany) operated at 3 kV, after 3 nm thick sputter coating of platinum. Ninhydrin Assay. Free amino group content on microgel particles was determined using a ninhydrin test in a 96-well plate format. The ninhydrin reagent was prepared in ethanol at a concentration of 50 mg/mL. Equal volumes of microgel solution and ninhydrin reagent were mixed in a 96-well plate and incubated at 50 °C for 20 min. The

Figure 1. Schematic representation of the synthesis of acetyl-CoA by acetyl CoA synthetase immobilized on PNIPAm-AEMA core−shell microgel. 2777

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Figure 2. Synthesis and properties of PNIPAm-AEMA core−shell microgel. (a) Scheme of synthesis; (b) Hydrodynamic radius of PNIPAm core (red circle) and PNIPAm-AEMA core−shell microgel (black triangle) as a function of temperature determined by DLS; (c) Scanning electron microscopy image of core−shell microgel after vacuum drying at room temperature. absorbance was checked at 590 nm using a plate reader (TICAN, Austria). The concentration of amino group was determined from standard plot of glycine as standard.29 Determination of the Enzymatic Activity and Protein Concentration. The activity of free acetyl CoA synthetase and their bioconjugates with PNIPAm-AEMA microgel was studied in borate buffer (0.1 M, pH 8) in the presence of potassium acetate (20 mM), MgCl2 (4 mM), glutathione (4 mM), ATP (1 mM), and coenzyme A (500 μM). The reaction was terminated by the addition of acidic molybdate reagent. The pyrophosphate−molybdate complex formed was further reduced by the addition of mercaptoethanol and Eikonogen reagent. The obtained colored product was quantified by measuring the absorption at 590 nm and pyrophosphate concentration was determined (ε = 2.7 × 104 M−1 cm−1).30 One unit of enzyme activity was defined as the amount of enzyme required to form 1 μmol pyrophosphate per min at pH 8 and at 37 °C.31 Bradford micro assay with a sensitivity of 1−10 μg/mL was used to determine protein concentration using bovine serum albumin as the standard protein.32 Immobilization of Enzyme on Microgel. Acs was covalently immobilized to the PNIPAm-AEMA core−shell microgel using carbodiimide chemistry in borate buffer (0.1 M) at pH 8.33,34 Inorganic phosphate present in commercial enzyme preparation (interferes with molybdate assay) was removed by filtration using 30 kDa centrifuge filter tubes (Millipore). The immobilization condition

was standardized before conjugation with respect to EDC, microgel, and salt concentration. A total of 6 mL of mixture solution containing microgel solution (10 mg/mL), enzyme solution (1 mg/mL), and EDC (1 mg/mL) were incubated at 10 °C for 4 h. After the completion of the incubation period, an equal volume of tris-Cl buffer (0.1 M, pH 8) was added to the reaction mixture to quench extra EDC. Thus, bioconjugate formed was precipitated by adding salt solution (1 M NaCl) and obtained by centrifugation at 12000 rpm for 5 min at 25 °C. Finally, the bioconjugate was desalinated by repeated centrifugation with borate buffer using 10 kDa MWCO centrifuge filter tubes. The amount of enzyme loaded to microgel was determined by subtracting the residual protein content in the supernatant from the initial protein content. The enzyme and bioconjugate were stored in borate buffer (0.1 M) containing glutathione (1 mM) and magnesium chloride (1 mM; pH 8). The number of molecules per unit volume (Nm and Ne) for microgel and enzyme were calculated using the following formula:29,35

N=

V vi

(1)

where V is the total volume of particles per unit volume of dispersion (mL) and νi is the mean volume of a particle. The value of V was found as 2778

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Figure 3. Standardization of immobilization parameters by checking the (a) effect of EDC, (a) microgel, and (c) salt on activity of enzyme. Relative activities are normalized activities of enzymes with respect to highest activity of respective enzymes and experiment. Each experiment was done in duplicate error bars in figures show standard deviations.

V=

W ρ

under a broad spectrum of parameters such as temperature, storage time, and reusability. The thermal stability of the prepared conjugate and free enzymes was assessed by determining the activity under standard conditions after subjecting to various temperature ranges from 32 to 60 °C for 10 min, followed by cooling on ice.38 Similarly, the storage stability was monitored by measuring the activity of enzyme stored at 4 °C in 0.1 M borate buffer. For control experiments, assay for free enzyme was run in parallel. Reusability of immobilized enzyme was also evaluated by consecutive operation cycles of enzymatic reaction for 15 min, conjugate were obtained as mentioned in immobilized enzyme preparation and further washing with 0.1 M borate buffer. The procedure was repeated using fresh aliquot of substrate.

(2)

W is the mass of wet microgel or enzyme and ρ is the density since the microgel particles were highly swollen in water ρ for microgel was assumed to be 1.00 g mL−1 and average protein density for enzyme 1.35 g mL−1. The value of νi was determined as vi =

⎛4⎞ 3 ⎜ ⎟π R ⎝3⎠ h

(3)

where Rh is the hydrodynamic radius. Using these data, the number of enzyme molecules on single microgel particles was calculated from Ne/ Nm. Effect of Temperature, pH, and Substrate Concentration. The effect of various reaction parameters such as temperature, pH, and substrate concentration on immobilized and free enzymes reactivity was studied. Optimum pH condition for both immobilized and free enzymes was evaluated in the range of 5−9. The enzymes were assayed in 0.1 M 2-(N-morpholino)ethanesulfonic acid (MES) buffer (pH 5− 6.5) and 0.1 M borate buffer (pH 7−9) at 37 °C for 15 min for respective pH. The effect of temperature on enzymes reactivity was studied at different temperatures ranging from 25 to 55 °C in 0.1 M borate buffer (pH 8) for 15 min. The enzyme activity was normalized with respect to maximum activity for given experimental condition and the final data were represented as relative activity. The kinetic constants for acetate as a substrate were obtained by varying its concentration from 0.5 to 10 mM at a constant cofactor concentration (500 μM ATP and 100 μM Co enzyme A).36,37 Assuming Michaelis− Menten kinetics for Acs, which is represented by following equation:

V=

Vmax[S] K m + [S]

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RESULTS AND DISCUSSION Synthesis of PNIPAm-AEMA Core−Shell Microgels. The core−shell microgel particles were synthesized by two-step free radical precipitation polymerization method which is described in Figure 2a. The basis for using core−shell morphology was to provide high amine group density on the surface of microgel particles, which enable better enzyme load.27 In the first step, the core was synthesized by polymerization and cross-linking of NIPAm with MBA in the presence of SDS as surfactant. The anionic surfactant has major role in controlling the shape and size of the core particle. In second step, the presynthesized core was used as seed for the synthesis of shell using an additional cationic comonomer AEMA to obtain amino functionalized core−shell microgel.27,28 During the synthesis of core, homogeneous nucleation mechanism and enhanced anionic surface charge density (a combined effect of sulfate ions and SDS) leads to the formation of small sized with low polydispersity core.22 While during shell synthesis, the cationic monomer plays an important role in chain transfer mechanism and controlling the size and shape of final microgel particles. The size and morphology of the core and core−shell microgels was studied by DLS and SEM characterizations. The size variations of microgels with respect to temperature are depicted in Figure 2b. The size of the core and core−shell microgels in water at 24 °C was found to be 150 and 320 nm, respectively. While at 40 °C, the size changed to 60 and 180 nm for core and core−shell microgel, respectively. PNIPAm exhibits reversible transition from a solvated random coil to a desolvated globular state at 32 °C, due to the disruption of hydrogen bonds and the dominance of the hydrophobic part of PNIPAm at high temperature, causing water to expel out of its structure.22,39 The microgels possess a

(4)

where V is the rate of the reaction, [S] is the concentration of the substrate, Km is the apparent constant, and Vmax is the maximum of reaction velocity. The values for Km and Vmax were estimated with the help of Lineweaver−Burk plot for the Michaelis−Menten equation expressed as follows:

K 1 1 1 = m − + V Vmax [S] Vmax

(5)

Fluorescence Study. The tertiary structure of free and immobilized Acs was characterized by tryptophan fluorescence measurements using a microtiter plate reader (Infinite M200 Pro Tecan, Austria). The excitation wavelength was set at 280 nm, and the emission spectrum was recorded from 310 to 390 nm. Further temperature-dependent enzyme conformation change was recorded in the temperature range 25−42 °C. Thermal, Storage, and Operational Stability Study. The longterm use of enzyme catalyst is based on their stability and reactivity 2779

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microgel was determined to be 68% of the initial Acs, suggesting the immobilized enzyme of 0.02 mg per mg of dry microgel particles. From this protein data, ∼24 Acs molecules were calculated to be bound on the surface of each microgel particle using eqs 1−3.29 The activity of immobilized Acs was found to be 0.23 mU/mg of carrier, which was about 61.55% of the initial activity of Acs. The attachment of enzyme to microgel using carbodiimide chemistry resulted in more active enzyme with respect to total bound enzyme. In the case of immobilization on CNBr-activated glass beads, 100% enzyme immobilization was achieved, but only 23% of enzyme was reported to be active.15 The decrease in enzyme activity after immobilization is the effect of several factors such as change in enzyme confirmation or modification of important residues due to covalent immobilization, and addition to this immobilization matrix also imparts additional factors like steric hindrance, partition effects, and mass transfer constraints.38 Effect of Temperature, pH, and Substrate on Acs Activity. Temperature and pH are two important parameters that influence activity of enzymes. It is important to study these factors to determine any change in conformation of enzyme on binding to the support.42 pH-dependent activity of enzyme was studied from pH range 5−9 and the properties of immobilized enzyme were compared with those of free enzyme. It can be seen from Figure 4a that free enzyme has optimal pH at 8.0,

volume phase transition from a swollen state to a collapsed state at or near this temperature. Furthermore, this transition temperature is also affected by the presence of functional comonomer in the structure.22,27 Figure 3a confirmed that the synthesized microgel particles exhibit the phase transition at around 32 °C, which is near to the LCST of PNIPAm. The morphology of the core−shell microgel was characterized with the help of SEM, and the image is depicted in Figure 2c. It is evident from the SEM image that the microgels were nearly uniform in size and shape. The cationic nature of prepared core−shell microgel was confirmed by measuring the zeta potential. The zeta potential value of diluted microgel sample in deionized water at pH 7 was found to be 8.60 mV, which clearly indicates the cationic behavior of microgel particles. This behavior of microgels was originated due to the presence of free −NH2 groups in shell. The presence of the free −NH2 group of AEMA comonomer in the shell was also confirmed by Ninhydrin assay. The free amino group concentration was obtained as 71 μM/mg of dry microgel with respect to glycine as a standard. Thus, the positive value of zeta potential and free amine content in microgel confirms that the microgels were sufficiently positively charged.29,40 Immobilization of Acetyl CoA Synthetase on PNIPAmAEMA Core−Shell Microgels. It is known from the structure of the active site of acetyl CoA synthetase that lysine 675 residues is very critical in the first step of the reaction, where acetate is activated to acetyl phosphate.12 Therefore, for covalent conjugation of enzyme to microgel, the carboxylic acid of enzyme was targeted using zero length cross-linker EDC.33 The effect of EDC concentrations on Acs immobilization to microgel was studied from 0.25 to 5 mg/mL concentration. As shown in Figure 3a, the enzyme activity increased with an increase in EDC concentration up to 1 mg/ mL; above this concentration, there was a sharp decline in enzyme activity. Since EDC is a cross-linker, at higher concentration, it cross-links enzyme molecules to one another at a faster rate, leading to a decrease in enzyme activity. To determine the working concentration of microgel for enzyme immobilization, different concentrations of microgel from 1 to 10 mg/mL were incubated with 1 mg/mL of enzyme in the presence of EDC. The highest concentration of microgel for this study was maintained to be 10 mg/mL to avoid a very viscous solution above this concentration. From Figure 3b it can be seen that the enzyme activity increases with an increase in microgel concentration. Therefore, for further use, a working concentration of 10 mg/mL microgel was found suitable, which is a 1:10 proportion of enzyme to microgel for conjugation protocol. To obtain the bioconjugate after the immobilization process, the microgels were precipitated from a reaction mixture at room temperature with the help of salt.41 Hence, it was important to study the effect of salt concentration on enzyme activity. The conjugate was subjected to different concentrations of NaCl solution and activity was determined. At a given centrifugal condition, 1 M NaCl solution was optimum to obtain complete bioconjugate. Also, Figure 3c shows that no significant reduction in enzyme activity at 1 M NaCl concentration was observed as compared to 10−1000-fold less concentration of NaCl. After standardization of immobilization conditions, the conjugation was carried out at optimized conditions. From the Bradford method, the amount of Acs immobilized onto

Figure 4. Effect of pH and temperature on activity of free (▲) and immobilized Acs (red circle): (a) pH-dependent studies on the activities of free and immobilized Acs was carried out at 37 °C in buffers with different pH values; (b) Effect of different temperatures on the activities of free and immobilized Acs was studied in borate buffer pH 8. Relative activities are normalized activities of enzymes with respect to highest activity of respective enzymes and experiment. Each experiment was done in triplicate and error bars in figures show standard deviations calculated with three data points.

which is close to previous report on same enzyme,43 while for immobilized Acs, pH was shifted toward more alkalinity. The electrostatic interactions of enzyme with the matrix leads to unequal partitioning of H+ and OH− between the microenvironment of the immobilized enzyme and the bulk phase. This is attributed to the Donnan partition effect where charged support often leads to displacements in the pH activity profile of immobilized enzymes.44 The temperature dependence on the activity of free and immobilized Acs was studied in the temperature range of 25 to 55 °C. The results presented in Figure 4b show that the immobilized enzyme was active at a broad temperature range and exhibited optimum temperature of reaction at 37 °C similar to free enzyme. These results can be attributed to rigidification 2780

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free enzyme was recorded with the change of temperature from 25 to 42 °C. Whereas in the case of immobilized enzyme (Figure 5b), no prominent change in intensity was observed below VPTT of microgel. Moreover, with the increase in temperature beyond 32 °C, only 12% reduction in intensity was seen at 42 °C. The comparatively low reduction in intensity of immobilized enzyme shows improved thermal stability of enzyme after immobilization on microgel because of structural rigidity.44,49 Stability of Immobilized Acs. The attachment of enzyme to a suitable polymer matrix provides longer shelf life and thermal stability to the biocatalyst. The enhancement in thermal stability of immobilized enzyme was also checked by determining the activity after incubation of enzyme at specific temperatures (32−60 °C). The thermal stability of conjugate enzyme was found comparatively higher than the free enzyme (Figure 6a). The enhancement of rigidity of enzyme structure upon attachment to support resists change in native conformation of enzyme. Protection of enzyme conformation from environmental changes due to restricted mobility of the covalently immobilized enzyme on support imparts enhanced thermal stability.44 The similar behavior of PNIPAm−enzyme bioconjugate with other enzymes has also been reported, which shows higher thermal stability upon immobilization.50−52 In solution the native enzyme tends to acquire thermodynamically stable conformation which leads to distortion of structure and inactivation of enzyme. At a temperature below the LCST of polymer, it probably prevents the unfolding of enzyme by forming a hydrated surface layer, like a protective colloid.34,44 Figure 6b shows the data of storage stability for free and immobilized enzymes. The immobilized Acs maintains more than 90% of its initial activity after stored for 9 days at 4 °C, while free Acs demonstrates only 63% of its initial activity under the similar conditions. This indicates that the storage stability of enzyme was improved remarkably after the immobilization on PNIPAm-AEMA core−shell microgel. This result is comparable to report on Acs immobilized on CNBractivated glass beads where they obtain relative activity between 80 and 90% on the ninth day of storage.15 Reusability of bioconjugate depends on the extent of stabilization, which is directly related to the extent of covalent bond formation and electrostatic interactions. Immobilization of enzyme allows switching of conjugate from the soluble to the insoluble form in solution making easy recovery from the reaction mixture.53,54 The operation stability of this immobilized Acs was studied to estimate the number of times immobilized enzyme can be reused. For the first cycle the free enzyme activity was 12 mU/mL, this value is 2.4 times higher than immobilized enzyme activity. After the first cycle, the recovery of free enzyme from solution was not achieved. Where as from Figure 6c, it can be seen that the immobilized Acs preserved 50% of its initial activity after four consecutive operations.

of protein conformation due to covalent immobilization of enzyme making it less susceptible to the temperature-induced conformational changes.44 Michaelis−Menten kinetics gives a direct estimation of enzyme activity affected due to interruption of steps involved from enzyme substrate binding to final product release. Immobilization has influence on the activity of enzyme by making structural changes, which may lead to denaturation blocking the active site inhibiting diffusion of the substrate to the enzyme or general microenvironment effects.45The apparent Km and Vmax values calculated from Lineweaver− Burk plot is presented in Table 1. Apparent Km values of acetate Table 1. Kinetic Parameters of Immobilized and Free Enzymes for Acetate Substrate at 37 °C enzyme

Km (μM)

Vmax (U/min)

free enzyme immobilized enzyme

170 ± 30 188 ± 20

8.5 × 10−3 ± 0.4 × 10−3 5.6 × 10−3 ± 0.1 × 10−3

for immobilized and free enzymes are close to each other. The Michaelis constant (Km) is a direct measure of the enzyme substrate affinity. The marginal increase of Km of immobilized enzyme indicates that the Acs−substrate binding continues to be efficient. While the maximum velocity (Vmax), which is the highest rate of substrate conversion when the enzyme is fully saturated with the substrate. In the present study, a 34% reduction in Vmax was observed for immobilized enzyme as compared to free enzymes. As Vmax is dependent on active enzyme concentration, the decrease in maximum velocity may be due to inactivation of enzyme particles or due to limited diffusion of substrate to immobilized enzyme.45,46 Fluorescence Study. Tryptophan (Trp) fluorescence study provides information about the structures of protein adsorbed to optically transparent substrates.47 The changes observed in fluorescence intensity and λmax can be attributed to the change in Trp environment, hinting to altered enzyme conformation.48 From Figure 5 the fluorescence spectra of both free and

Figure 5. Fluoroscence emission spectra (Ex280/Em 310−390)) at different temperatures of free (a) and immobilized (b) Acs in 0.01 M phosphate buffer pH 7.4 (protein concentration = 0.05 mg/mL).

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CONCLUSION The novelty of work lies in investigation of core−shell thermal responsive PNIPAm-AEMA microgel as a support for acetyl CoA synthetase to obtain reusable biocatalyst. The cationic microgel synthesized was nearly uniform in size and contained a sufficient amino group for enzyme attachment. On immobilization of Acs at standardized conditions, the reaction parameters like temperature and pH were optimum for a wide range. The developed and optimized procedure of immobilization allows

immobilized Acs shows λmax at 330 nm, which indicates that there is no significant change in immobilized enzyme native conformation. Further, the temperature-dependent intensity profile was studied from the temperature range 25−42 °C to determine conformational stability in enzyme after immobilization. The temperature-dependent fluorescence spectra of free enzyme (Figure 5a) indicate a linear decrease in the intensity with increasing temperature. About 30% decrease in intensity of 2781

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Figure 6. Stability parameters of free (▲) and immobilized Acs (red circle): (a) Thermal stability of immobilized and free Acs. The activity of free and immobilized Acs were determined after incubating at respective temperatures for a 10 min and cooled on ice. (b) Storage stability at 4 °C was monitored by checking the activity of free and immobilized Acs in borate buffer (0.1 M, pH 8). (c) Operation stability of immobilized Acs was checked for 5 cycles at standard reaction conditions. Relative activities are normalized activities of enzymes with respect to highest activity of respective enzymes and experiment. Each experiment was done in triplicate and error bars in figures show standard deviations calculated with three data points. (9) Woodyer, R. D., Johannes, T. W.; Zhao, H. In Enzyme Technology; Pandey, A., Webb, C., Soccol, C. R., Larroche, C., Eds.; Springer Science: Delhi, India, 2006; pp 85−104. (10) Mishra, P. K.; Drueckhammer, D. G. Chem. Rev. 2000, 100, 3283−3309. (11) Patel, S. S.; Walt, D. R. J. Biol. Chem. 1987, 262, 7132−1134. (12) Jogl, G.; Tong, L. Biochemistry 2004, 43, 1425−1431. (13) Sheldon, R. A. Adv. Synth. Catal. 2007, 349, 1289−1307. (14) Kudina, O.; Zakharchenko, A.; Trotsenko, O.; Tokarev, A.; Ionov, L.; Stoychev, G.; Puretskiy, N.; Pryor, S. W.; Voronov, A.; Minko, S. Angew. Chem., Int. Ed. 2014, 53, 483−487. (15) Mannens, G.; Siegers, G.; Lambrecht, R.; Claeys, A. Biohim. Biophys. Acta 1988, 959, 214−219. (16) Mannens, G.; Slegers, G.; Lambrecht, R.; Goethals, P. J. Labelled Compd. Rad. 1988, 25, 695−705. (17) Sokic-Lazic, D.; Minteer, S. D. Biosens. Bioelectron. 2008, 24, 939−944. (18) Takagi, K.; Mochizuki, M.; Sakamoto, I.; Teranishi H. Carriers for Immobilization of Physiologically Active Substances. U.S. Patent, 4,610,962, Sept. 9, 1986. (19) Cayre, O. J.; Chagneux, N.; Biggs, S. Soft Matter 2011, 7, 2211− 2234. (20) Ramli, R. A.; Laftah, W. A.; Hashim, S. RSC Adv. 2013, 3, 15543−15565. (21) Ghosh Chaudhuri, R.; Paria, S. Chem. Rev. 2012, 112, 2373− 2433. (22) Pelton, R.; Hoare, T. In Microgel Suspensions: Fundamentals and Applications; Fernandez-Nieves, A., Wyss, H. M., Mattsson, J., Weitz, D. A., Eds.; Wiley-VCH: Weinheim, 2011; pp 3−32. (23) Hamerska-Dudra, A.; Bryjak, J.; Trochimczuk, A. W. Enzyme Microb. Technol. 2006, 38, 921−925. (24) Popat, A.; Hartono, S. B.; Stahr, F.; Liu, J.; Qiao, S. Z.; Lu, G. Q. Nanoscale 2011, 3, 2801−2818. (25) Lyon, L. A.; Meng, Z.; Singh, N.; Sorrell, C. D.; John, A. S. Chem. Soc. Rev. 2009, 38, 865−874. (26) Nayak, S.; Lyon, L. A. Angew. Chem., Int. Ed. 2005, 44, 7686− 7708. (27) Nayak, S.; Lee, H.; Chmielewski, J.; Lyon, L. A. J. Am. Chem. Soc. 2004, 126, 10258−10259. (28) Silva, C. S. O.; Baptista, R. P.; Santos, A. M.; Martinho, J. M. G.; Cabral, J. M. S.; Taipa, M. A. Biotechnol. Lett. 2006, 28, 2019−2025. (29) Zhang, H.; Mardyani, S.; Chan, W. C. W; Kumacheva, E. Biomacromolecules 2006, 7, 1568−1572. (30) Putnins, R. F.; Yamada, E. W. Anal. Biochem. 1975, 68, 185− 195.

retaining activity of enzyme on microgel particles. Its thermal, storage, and operational stability are significantly improved after immobilization, indicating that PNIPAm-AEMA core−shell microgel is the favorable carrier for enzyme immobilization. Moreover, the immobilized enzyme could be reused for four consecutive operation cycles with more than 50% initial activity that opens new perspective for efficient synthesis of many drugs.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Fax: +49-3514658281. Tel.: +493514658271. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank Ms. A. Caspari and Dr. C. Bellman for assisting in DLS and zeta potential measurements. The authors are grateful to Dr. D. Appelhans and N. Polikarpov for their help in microgel synthesis, Dr. M. Müller, Dr. M. Maitz, and Dr. M. Tsurkan for allowing instrument facilities. B.P.T. acknowledges Alexander von Humboldt Foundation for AvH Postdoctoral Fellowship.

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