The use of isocyanide-based multicomponent reaction for covalent immobilization of Rhizomucor miehei lipase on multiwall carbon nanotubes and graphene nanosheets

Abstract

We describe here a novel and simple method for making bioconjugation and immobilization of Rhizomucor miehei lipase (RML) on carboxylated multiwall carbon nanotubes (MWCNT-COOH) and carboxylated graphene nanosheets (Gr-COOH) by using an isocyanide-based four-component reaction. In this approach, the enzyme supplies amino groups, the support supplies carboxylic acid groups, and the missing components (isocyanide and aldehyde) are added to the reaction medium. This coupling reaction was carried out in water at 25 °C, in which rapid and high enzyme loading were achieved. The maximum loading capacity of 530 mg and 680 mg was obtained for Gr-COOH and MWCNT-COOH, respectively. A variety of techniques including FTIR, Raman spectroscopy, XRD, SEM, and TGA were employed to characterize the immobilized derivatives of RML. The immobilized preparations showed significantly increased thermal stability and co-solvent stability as compared to the soluble enzyme. Kinetic parameters and optimum pH activity of RML and its immobilized preparations were also determined. The K_m values of 0.44, 0.23, and 0.18 mM and the maximum reaction rates (V_max) of 0.09, 0.1, and 0.08 mM min⁻¹ were obtained for MWCNTs-RML, Gr-RML, and free RML, respectively. This approach may provide a general and efficient method to attach biomolecules on a variety of carboxylated solid surfaces at ambient conditions.

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1. Introduction

Enzymes possess efficient properties in terms of high catalytic activity, operations under mild conditions, exclusion of undesirable side reactions, and having different types of selectivity in chemical reactions.¹ Therefore, application of enzymes is increased in several industrial demands.¹ Among them, lipases (with the natural catalytic function of hydrolyzing fatty acid ester bonds) are suitable for organic syntheses because of accepting a wide range of non-natural substrates without requiring cofactors, stability and activity in organic solvents, and availability from several (micro) organisms.^2,3 Application of enzymes in free form are limited by enzyme inactivation under harsh conditions and difficulty with recovering and reusing native enzyme from the reaction medium.^4,5 Therefore, utilizing an economically attractive process that allows improving operational stability of an enzyme and preserving its catalytic properties is a challenge of great interest.^6,7 A great number of materials, both inorganic and organic, have been used as carriers for enzyme immobilization.^8–10 Combined with recent progress in nanotechnology, nanoscale materials with a large surface-to-volume ratio such as metal nanoparticles, nano mesoporous materials, and carbon nanotubes are in the spotlight as immobilization supports.^2,11 In particular, carbon-based nano materials (CNMs) are used widely for immobilization of bio-macromolecules due to their mechanical, thermal, and electrical properties and general biocompatibilities.^12,13 Especially when the support has a porous structure, a highly stable biocatalyst and high immobilization yield can be obtained.¹⁴ In fact, immobilization of biomolecules on these materials is often a key step in their biotechnological and biological applications. Both covalent and non-covalent attachment can be used for immobilization of proteins on the surface of CNMs.^12,15 Although non-covalent immobilization preserves the structure of both the nanomaterial and the protein well, leaching of protein from the CNMs cannot be excluded.^16,17 On the other hand, the best stability and selectivity will be achieved via covalent bonding because of its ability to rigidify the location of the immobilized protein.¹⁸ However, covalent attachment in many cases reduces enzyme activity.¹⁹ Moreover, the surface of CNMs must be modified with proper functional groups to bind with proteins via covalent attachment. Therefore, except for electrochemical methods, there are only small number of protocols for covalent linkage of proteins onto CNMs and these are generally limited to (1) direct coupling of a protein using a diimide-activated amidation procedure, and (2) chemical amination on the surface of the support and subsequent immobilization of a protein by using a proper linker or a cross-linker reagent.²⁰ For example, covalent attachment of bovine serum albumin (BSA) by direct coupling of carboxylic acid groups of carbon nanotubes with surface-exposed amino groups of a protein using N-ethyl-N′-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDAC) or N,N′-dicyclohexylcarbodiimide (DCC) as coupling agents has been reported.²¹ The other universal method for covalent immobilization of proteins on these materials is based on further modification of carboxylic acid groups to produce other electrophilic moieties on the surface of a support.²⁰ Immobilization of enzymes onto CNMs has also been performed using linker molecules such as N-succinimidyl 1-pyrenebutanoate, 1-aminopyrene, polyethylene glycol, and glutaraldehyde.²⁰ Although the amino functionalized CNMs are a well-known support for enzyme immobilization, support modifications are normally time consuming and require more chemical reagents.²²

Shortening the number of steps required for functionalization of a support will decrease total cost of the process. Therefore, there is a big demand for discovery of simple protocols for functionalization of CNMs and subsequent immobilization of biomolecules on them. Recently, we reported the results of using the Ugi four-component reaction for covalent attachment of enzymes on aldehyde functionalized supports.²³ This simple and efficient procedure offers great flexibility and can accommodate different chemistries as well as surfaces of choice. In fact, by utilizing this method a large number of supports and functional groups can be used for immobilization of biomolecules. The Ugi four-component reaction is a condensation reaction between an aldehyde, an amine, a carboxylic acid, and an isocyanide (Scheme 1).

Scheme 1 The Ugi four-component reaction.

We describe here using the Ugi four-component reaction for one-pot immobilization of biomolecules on carboxylic acid-functionalized multiwall carbon nanotubes (MWCNTs-COOH) and graphene nanosheets (Gr-COOH) for the first time. For this, Rhizomucor miehei lipase was selected as a model enzyme in order to assess the potential applicability of CNMs as a carrier for immobilization of biomolecules. The lipase from Rhizomucor miehei is a commercially available enzyme in both soluble and immobilized form. The enzyme is described as a single polypeptide chain of 269 residues with molecular size of 31 600 Da and a pI of 3.8.²⁴ RML is a quite active and stable lipase and presents more advantages in industry compared to other lipases.²⁵ The results of the present protocol are promising to be used not only for immobilization of biomolecules but also for chemical functionalization of CNMs.

2. Experimental

2.1. Materials

The graphite used for preparing graphene nanosheets was expandable graphite, supplied by Shandong Qingdao Graphite Company (China). Carboxylated multiwalled carbon nanotubes (MWNTs-COOH, D: 8–15 nm, length: 50 nm, surface area per unit mass: 233 m² g⁻¹, purity >95%) were obtained from Neutrino Port. Co. (Cheap Tubes Company, USA). The lipase from Rhizomucor miehei, p-nitrophenyl butyrate (p-NPB), cyclohexyl isocyanide, and acetaldehyde were purchased from Sigma. Dioxane, 1-propanol, 2-propanol, THF, DMSO, acetonitrile, sulfuric acid, and nitric acid were purchased from Merck. Other reagents and solvents used were of analytical grade. Fourier transform infrared spectra (FT-IR) were recorded on a Bomen FT-IR-MB-series instrument with a KBr pellet technique. Thermogravimetry (TGA) and differential thermal analysis (DTA) were carried out from 10 °C to 800 °C at a heating rate of 20 °C min⁻¹ in air atmosphere using a STA 503M system from Bähr GmbH, Germany.

2.2. Methods

2.2.1. Preparation of graphene nanosheets. The expandable graphite was heat treated at 950 ± 10 °C for 30 s to obtain expanded graphite. The expanded graphite was immersed in a 75% aqueous alcohol solution in an ultrasonic bath. The mixture was sonicated for 10 h, and then filtered and washed with distilled water and ethyl alcohol. The obtained graphene nanosheets were dried under vacuum at ambient temperature for 12 h and stored at room temperature.

2.2.2. Carboxylation of graphene nanosheets. Carboxylation was carried out by reacting graphene nanosheets with a strong acids oxidizing solution. Graphene nanosheets (1 g) were added to a 3 : 1 mixture of concentrated H₂SO₄ and HNO₃ (120 : 40 mL). The reaction was sonicated in an ultrasonic bath for 1 h and then stirred for 24 h at room temperature. The mixture was centrifuged at 6000 rpm for 1 h and the solid materials were washed with 300 mL of deionized water (repeatedly until no acid was detected in the supernatant) and then with ethanol (2 × 300 mL), followed by vacuum drying. The obtained carboxylated graphene nanosheets were stored at room temperature.

2.2.3. Enzyme immobilization on multi-walled carbon nanotubes and graphene nanosheets. An amount (5 mg) of the MWNTs-COOH (or Gr-COOH) was added to 2 mL of distilled water (pH 7.0) containing a certain amount of RML and acetaldehyde (20 μL). Immobilization of RML was started by addition of 10 μL of cyclohexyl isocyanide to the solution under gently stirring at 25 °C. Periodically, samples of the supernatant were withdrawn and analyzed by activity assay and the Bradford's method²⁶ for determination of protein concentration. Finally, the immobilized RML derivatives were washed by distilled water and stored at 4 °C for further characterization (Section 3.1.).

2.2.4. Enzymatic activity assay. Two methods were used for activity measurement of the derivatives. In the first method the activities of the soluble lipase and its immobilized preparations were analyzed spectrophotometrically by measuring the increment in absorbance at 348 nm (ε = 5150 M⁻¹ cm⁻¹). The increase in absorbance produced by the release of p-nitrophenol in the hydrolysis of p-NPB in 25 mM sodium phosphate buffer at pH 8.0 and 25 °C. Briefly, 0.05–0.2 mL of the lipase suspension or solution (blank or supernatant without further dilution) was added to 2.5 mL of substrate solution (0.8 mM) under magnetic stirring.²⁷ Spontaneous hydrolysis of p-NPB was measured using 2.5 mL of substrate solution (0.8 mM) in the absence of RML as control. Enzyme specific activity is given as 1 μmol of p-nitrophenol released per minute per mg of the enzyme (IU) under the conditions described above. In the second method the activity of the derivatives was assayed in a pH-stat system (Metrohm, Switzerland) by using 20 mL substrate (1% of each substrate, 2% gum arabic, mixed and ultrasonicated for 5 min, pH 8.0 and 25 °C). In this way a certain amount of each derivative was titrated automatically with 0.05 M NaOH. Substrate specificity of the lipase was determined by measuring activity in the presence of C8, C12, and C18 as substrates. Enzyme specific activity is defined as the amount of lipase that releases 1 μmol of fatty acid from the substrate in 1 min at 25 °C.

2.2.5. Determination of the amount of RML bound to the carriers. The concentration of RML in the solutions was determined by the Bradford's method. The reported yields of immobilization were calculated as the ratio of protein bound to the support to the initial amount of protein. Yields are expressed as percentage.

2.2.6. Thermal inactivation of RML immobilized preparations. Free enzyme and immobilized preparations of RML were incubated in 25 mM sodium phosphate at pH 7.0 at different temperatures. The suspension of each sample was withdrawn periodically and their activities were measured using the p-NPB assay.

2.2.7. Stability of RML immobilized preparations in presence of organic solvents. Free enzyme and immobilized preparations were incubated in a total volume of 1 mL solution containing 25 mM sodium phosphate buffer pH 7.0 and 10% or 20% of dioxane, 1-propanol, 2-propanol, methanol, acetaldehyde, DMSO, and THF at 25 °C. The activity of each sample was measured using the p-NPB assay.

2.2.8. Leaching experiment. Different preparations of MWNTs-RML and Gr-RML (5 mg) were incubated in a solution containing 1 M of NaCl with vigorous magnetic stirring for 24 h. Then the concentration of RML in the supernatant was measured by using enzyme activity assay and the Bradford's method.

2.2.9. Determination of kinetic parameters. Kinetic parameters of the free and immobilized preparations of RML were determined using different p-NPB concentrations in the range of 0.063–1.25 mM in 25 mM sodium phosphate buffer (pH 8.0). K_m and V_max values of the free and immobilized lipase were calculated via non-linear regression fitting of the of the Michaelis–Menten equation using Prism 6 (Graphpad Software).

3. Results and discussion

3.1. Covalent immobilization of RML on graphene nanosheets

The choice of the reactive functional group on carbon-based materials was simply made, keeping in mind that one of the most common functional groups which can be easily introduced on the surface of CNMs is carboxylic acid groups. To demonstrate the utility of the proposed method, RML was immobilized on carboxylated graphene nanosheets. This novel approach is based on the use of the Ugi multi-component reaction for covalent attachment of enzymes. In this four-component reaction the support supplies carboxylic acid groups, the enzyme supplies amino groups, and the two missing components (isocyanide and aldehyde) are added to the reaction medium. Extremely mild condition (water and 25 °C) was used to perform this coupling reaction. Similar to our previous report,²³ negligible immobilization of RML on both supports was observed in sodium phosphate buffer solution even at low ionic strength (5 mM) after a long time incubation. Further investigation is needed to find the reasons for a negative effect of impurities on enzyme immobilization in this procedure. Carboxylated graphene nanosheets have a unique planar structure and intriguing mechanical properties that have attracted intensive interests. To investigate the effect of the amount of offered enzyme on immobilization yield and final activity of the immobilized derivatives, different amounts of RML (400, 500, 800, and 1000 mg) were offered to immobilize on 1 g of Gr-COOH under gentle stirring for 5 h. Further incubation of the enzyme for a long time (24 h) had a negligible effect on the immobilization yield in all cases. However, complete immobilization of up to 100 mg of the enzyme was performed shortly after 30 minutes of incubation. Table 1 shows the parameters of immobilization of RML on Gr-COOH using various enzyme-to-support weight ratios. Immobilization of 400 mg of RML on Gr-COOH produced 86% of immobilization yield after 5 h of incubation. It means that 340 mg of the offered enzyme is immobilized on the support at this condition. Increasing the enzyme to 500 mg caused a decrease in the immobilization yield to 73%. However, the amount of the immobilized enzyme improved to 365 mg on 1 g of the support. With further increasing the amount of offered enzyme to 800 and 1000 mg, lower immobilization yields were obtained. This investigation also showed that the maximum loading capacity of 1 g of Gr-COOH is about 530 mg of the enzyme. To date different loading capacities for carboxylated graphene nanosheets have been reported in the literature. To the best of our knowledge this is the highest reported capacity of immobilization of an enzyme on this support via covalent linkage. However, physical adsorption of 800 mg g⁻¹ of acid pectinase onto graphene nanosheets has been achieved previously.²⁸

Table 1 Immobilization of Rhizomucor miehei lipase on multiwall carbon nanotubes and graphene nanosheets

Offered enzyme (mg g^−1)	Yield^a (%)		mg enzyme per g support		Desorption^b (mg)		Specific activity^c (U mg^−1 RML)
	Gr	MWCNTs	Gr	MWCNTs	Gr	MWCNTs	Gr	MWCNTs
a Yield is defined as the percentage of the soluble enzyme that becomes attached to the supports after 5 h.b Desorption was performed in a solution containing 1 M NaCl.c Specific activity (U mg^−1 lipase) is expressed as micromole of substrate hydrolyzed per minute per mg of RML. The specific activity of the soluble RML was 5.9.
400	86	92	340	368	12	10	5.7	5.8
500	73	90	365	450	17	11	5.8	5.6
800	56	80	448	640	13	14	5.5	5.5
1000	53	68	530	680	21	19	5.3	5.1

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A desorption experiment was performed by incubating the immobilized derivatives in a solution containing 1 M NaCl under vigorous stirring in order to ensure that the immobilization is covalent. After 24 h, the solid was separated from the supernatant by centrifugation. The Bradford assay showed a release of only 12–21 (3.5–4%) of the immobilized enzyme in solution, clearly proving that by using the multicomponent approach immobilization is performed via covalent binding. The results also showed no detectable immobilization in directed attachment of RML on Gr-COOH without using isocyanide and aldehyde.

The specific activity of the immobilized preparations of RML on graphene nanosheets (Gr-RML) was also investigated as a function of the amount of loaded RML. The enzymatic activity was characterized by measuring the hydrolysis rate of p-NPB as a model substrate. The catalytic hydrolysis of p-NPB produces equal moles of p-nitrophenol which is determined spectrophotometrically. The results showed a little decrease (90–98%) in specific activity of the immobilized derivatives compared to specific activity of free RML, presenting remarkable efficiency of our method for enzyme immobilization. The results also showed that with increasing the amount of enzyme, the specific activity of the final derivative decreases. Decrease in enzyme specific activity is most likely because of diffusion limitation of substrate/product after immobilization of the enzyme.¹⁸

3.2. Covalent immobilization of RML on carboxylated multiwall carbon nanotubes (MWCNTs-COOH)

Covalent immobilization of RML on carboxylated multiwall carbon nanotubes was examined using the Ugi four-component approach. Various amounts of the enzyme were offered to the support. Immobilization of up to 100 mg of the enzyme was completed quickly after 15 minutes of incubation while for the higher amounts of RML longer times were needed. As Table 1 shows, both immobilization yields and loading capacities were greatly improved compared to Gr-COOH, suggesting that MWCNTs-COOH is a more efficient immobilization matrix for proteins. For example, with 400 mg of RML in the initial solution, 92% of immobilization yield was obtained. That means that 368 mg of the enzyme was immobilized on the support after 5 h of incubation. By increasing the amount of offered enzyme to 500, 800, and 1000 mg, immobilization yields decreased to 90, 80, and 68% respectively. Despite of these low immobilization yields, the loading amount of RML on the support dramatically improved. The higher capacity of carboxylated multiwall carbon nanotubes compared to Gr-COOH can be explained by its three dimensional structure which enables higher enzyme loading.²⁹ The maximum loading capacity of MWCNTs was determined as 680 mg which was much higher than those reported previously.^30,31 The results also showed that in higher immobilization yields, the specific activity of the obtained derivatives slightly decreased. For example, with immobilization of 680 mg of the enzyme, specific activity of the derivative decreased to 5.1, which is about 86% of the specific activity of the free enzyme. In fact, in a high enzyme loading diffusion of the substrate to the enzyme and of the product away from the enzyme becomes limited. To confirm the covalent nature of the enzyme attachment, a desorption experiment was performed. The results showed that only 3% of the immobilized enzyme has physical interactions and leached out from the support, indicating that the enzyme was irreversibly bound to the support. A desorption experiment for the derivative obtained from direct immobilization of RML on MWCNTs-COOH without using coupling agents (isocyanide and aldehyde) was also performed using the same conditions. More than 98% of the adsorbed enzyme was released to the solution, confirming that physical adsorption is the main mechanism of immobilization with the condition that was used.

3.3. Characterization of the immobilized derivative of MWCNTs-COOH

Characterization of the MWCNTs-COOH before and after immobilization of RML was carried out by using scanning electron microscopy (SEM), infrared spectroscopy (IR), thermal gravimetric analysis (TGA), X-ray diffraction analysis (XRD), and Raman spectroscopy.

Fig. 1 shows the high-resolution SEM images of the carboxyl-functionalized MWCNTs/graphene nanosheets and their RML immobilized derivatives. The surface of the MWCNTs-COOH has uniform and smooth layers and a clear rode-like shape, whereas its aggregates structure in the image of MWCNTs-RML is clear evidence for successful immobilization of the enzyme. It also shows that the immobilized preparation become thicker due to attachment of RML on the surface of MWCNTs. In other words, the diameter of MWCNTs-RML is slightly increased as compared to that of MWCNTs-COOH. These morphological characteristics are similar to the images reported in the previous study of enzyme organophosphate hydrolase on functionalized carbon nanotubes.³²

Fig. 1 Scanning electron micrographs of (a) MWCNTs-COOH, (b) MWCNTs-RML, (c) Gr-COOH, and (d) Gr-RML.

In the FTIR spectrum of MWCNTs-COOH (Fig. 2(a)), the peaks at 1764 and 3291 cm⁻¹ are due to the C=O and O–H stretching vibrations of the carboxylic groups. For the MWCNTs-RML (Fig. 2(b)) the C=O stretching frequencies of the amide bond formed by the functionalization reaction appeared at 1666 and 1538 cm⁻¹ which clearly indicate the successful attachment of RML on MWCNTs. Furthermore, the bands at 2928 and 2863 cm⁻¹ can be clearly assigned to the C–H stretching vibrations of the attached RML on MWCNTs.

Fig. 2 FT-IR spectra of (a) MWCNTs-COOH, (b) MWCNTs-RML, (c) Gr-COOH, and (d) Gr-RML.

The successful immobilization of RML on MWCNTs-COOH was further confirmed by thermogravimetric analysis (Fig. 3(a and b)). While the MWCNTs-COOH only showed a minor mass loss at 492 °C, the mass loss of 29% in sample MWCNTs-RML at the same temperature corresponds to removal of the enzyme molecules bond to the support surface. A similar observation was reported by Shi et al.³³ for MWCNTs-lipase preparations.

Fig. 3 TG analysis of (a) MWCNTs-COOH, (b) MWCNTs-RML, (c) Gr-COOH, and (d) Gr-RML.

Fig. 4(a and b) shows typical XRD patterns of MWCNTs-COOH and MWCNTs-RML. The first diffraction peak for the samples at the angle (2θ) of 25.70° can be indexed as the C (0 0 2) reflection of the hexagonal structure.^34,35 The other characteristic diffraction peaks at 2θ of about 44°, 64.48°, and 78.01° are associated with C (1 0 0), C (0 0 4), and C (1 1 0) diffractions, respectively.³⁴ There is no significant difference between the two patterns of MWCNTs-COOH and MWCNTs-RML, indicating that the carbon crystallite structure of MWCNTs was not affected by enzyme loading in the immobilization process.

Fig. 4 XRD analysis of (a) MWCNTs-COOH, (b) MWCNTs-RML, (c) Gr-COOH, and (d) Gr-RML.

Raman scattering was used for investigating disorder in the carbon skeleton of MWCNTs-COOH before and after immobilization of RML (Fig. 5(a and b)). The most characteristic peaks at 1338, 1565, and 2700 cm⁻¹ in the Raman spectra of both samples correspond to the graphite D-band, G-band, and 2D band, respectively. The G-band at 1338 cm⁻¹ can be attributed to sp²-hybridized carbons of the Raman-active E_2g mode of graphite while the D-band at 1338 cm⁻¹ is attributed to either sp³-hybridized carbons or to structural defect sites of the sp²-hydridized carbon network.³⁶ The relative intensity of D to G band (I_D/I_G) can be used to determine the degree of disorder in the carbon network. The profiles of Raman spectra of both MWCNTs-COOH (1.06) and MWCNTs-RML (0.98) are almost the same, suggesting that MWCNTs retains its basic structural properties during the immobilization of RML.

Fig. 5 Raman spectra of (a) MWCNTs-COOH, (b) MWCNTs-RML, (c) Gr-COOH, and (d) Gr-RML.

3.4. Characterization of the immobilized derivative of Gr-COOH

Functionalized graphene nanosheets before and after immobilization were also systematically characterized using SEM, IR, TGA, XRD, and Raman spectroscopy. Fig. 1(c and d) shows the SEM image of carboxylated-graphene nanosheets which confirms that there is no damage effect on the graphene nanosheets even under the strongest oxidation treatment of using mixtures of nitric acid sulfuric acid. The SEM micrograph of the obtained Gr-COOH also shows irregular particles with smooth surfaces. The changes in the surface morphology of the Gr-COOH after covalent immobilization of the enzyme are also observed.

In the IR spectrum of Gr-COOH (Fig. 2(c)), the peak at 3312 cm⁻¹ can be attributed to the –OH vibration stretching. Besides, it also shows other bands at 1662 cm⁻¹ for carboxyl groups and 1419 cm⁻¹ for C–O. In the immobilized derivative, the strong bands at 1667 and 1540 cm⁻¹ are according to N–H in-plane stretching of the amide I and amide II presented.³⁷ In addition, appearance of C–N stretching frequency at 1247 cm⁻¹ and aliphatic C–H stretch band at 2855–2940 cm⁻¹ clearly demonstrated successful functionalization of the support by RML (Fig. 2(d)).

The successful immobilization of RML on graphene nanosheets can also be concluded from TGA curves. As shown in Fig. 3(c and d), compared with Gr-COOH, Gr-RML shows much lower thermal stability. Its main mass loss takes place around 231–478 °C, which can be attributed to the removal of the enzyme from the surface of the support. The observed small mass loss over the temperature range between 478 and 705 °C can also be assigned to the removal of more stable functional groups.

The XRD pattern of Gr-COOH gives a distinguishable (002) graphite peak at 26° together with three other peaks at 44.22°, 54.41°, and 64.43° with low intensities. As Fig. 4(d) shows, after immobilization of the enzyme the XRD profile is not changed, suggesting that the crystalline structure of the support remained almost unchanged after enzyme attachment.

Raman spectroscopy was also used as an effective tool to investigate the structural characteristics and properties of Gr-RML. Fig. 5(c and d) shows a moderate D-band (∼1348 cm⁻¹), a moderate G-band (∼1570) cm⁻¹ and a strong 2D band (∼2714 cm⁻¹) in the Raman spectrum of Gr-COOH. On the other hand, three Raman bands are observed for Gr-RML: a band with relative intensity at 1452 cm⁻¹ (D-band), a band at 1509 cm⁻¹ (G-band), and a band with high intensity in 2428 cm⁻¹ region. Compared with Gr-COOH (1.19), the D/G intensity of Gr-RML (1.02) is almost similar; implying that immobilization of RML by the Ugi four-component reaction has no significant effect on the graphite structure of Gr-COOH.

3.5. Thermal and co-solvent stability of the free and immobilized derivatives

Organic solvents can cause conformational changes in enzyme structure, thus changing its catalytic properties. This is due to the fact that small amounts of water molecules are required for enzymatic function. Organic solvents, particularly those having log P values below 2, strongly distort this essential water–enzyme interaction, thereby inactivating or denaturing the enzyme structure. In order to investigate the effect of immobilization on the stability of RML, seven water-miscible solvents (10% and 20% of 1-propanol, 2-propanol, dioxane, THF, acetonitrile, DMSO, and methanol) with log P < 2 were used. Both immobilized derivatives showed improved stability compared to the soluble enzyme, retaining 100% of their initial activities after 24 h of incubation in the presence of 10% of the seven organic solvents, while the soluble enzyme retained 68–80% of its initial activity at this condition (results not shown).

Further investigation was performed using 20% of all solvents in order to determine the effect of increasing organic solvents percentage on the stability of both free enzyme and the immobilized derivatives. Fig. 6(c) shows the results of enzyme inactivation in the presence of 20% of seven chosen organic solvents. Both immobilized derivatives (MWCNTs-RML and Gr-RML) retain 100% of their initial activities after 24 h of incubation in the presence of 20% of methanol, while the free enzyme loses 80% of its activity under the same conditions. However, increasing the percentage of methanol to 50% leads to almost 75% and 90% decrease in activity of MWCNTs-RML and Gr-RML, respectively.

Fig. 6 Thermal (a, b) and co-solvent (c) stability of free RML and immobilized preparations.

Incubation of the soluble RML in the presence of 20% dioxane leads to 72% decrease in initial activity of RML, while immobilized derivatives are stable (more than 95%) at the same condition. On the other hand, while Gr-RML is quite stable in presence of 20% acetonitrile, MWCNTs-RML and free RML keep 75% and 20% of their activities after 24 h of incubation, respectively. The most promising result was obtained from an investigation on co-solvent stability of MWCNTs-RML. This derivative retains 71–100% of its initial activity after 24 h incubation in the presence of 20% of the seven organic solvents.

Thermal stability of free RML and its immobilized derivatives (MWNTs-RML and Gr-RML) was also investigated with various temperatures (Fig. 6(a and b)). A complete profile of thermal stability of the free RML was reported in our previous study.²³ The soluble RML is relatively unstable and loses 80% of initial activity at 50 °C after 6 h of incubation. Rapid inactivation of RML was also observed by increasing the temperature to 55 °C after 2.5 h of incubation. As results show, immobilization of RML on MWNTs-COOH leads to significant improvement in thermal stability of the immobilized derivative (Fig. 6(a)). MWNTs-RML is quite stable and retains 100% of its initial activity after 24 h of incubation at 45 °C. Furthermore, the results reveal retaining about 93% and 80% of initial activity MWNTs-RML during 24 h of incubation at 50 °C and 55 °C, respectively. By increasing the temperature to 60 °C, only 24% of the activity of MWNTs-RML disappears during 8 h of incubation which clearly confirms the positive effect of the immobilization procedure on stability of RML. The improved thermal stability is also observed for RML immobilized on functionalized Gr-COOH. As can be seen from Fig. 6(b), Gr-RML has great thermal resistance by retaining 100% of its activity after 24 h of incubation at 45–50 °C. This derivative also shows about 94% of its activity after 24 h of incubation at 55 °C. Furthermore, covalent immobilization of RML increases the stability of the obtained derivative with retaining 65% activity after 8 h incubation at 60 °C. Complete inactivation of the derivative is observed after 24 h incubation of the derivative at 60 °C. In comparision, the thermal stability of MWNTs-RML is more improved compared to immobilized preparation obtained from immobilization of RML on Gr, most likely because of the three dimensional structure of MWNTs which keeps away the enzyme structure from distorting agents. Another experiment was performed in order to study thermal stability of the derivatives at 45 °C for a period of 5 days. The results showed that free RML, Gr-RML, and MWNTs-RML retain their initial activities in the first four days at this condition. In the fifth day of incubation 69, 94, and 100% of activities were observed for free RML, Gr-RML, and MWNTs-RML, respectively. These results further confirmed the positive effect of both immobilization and the structure of the carrier on stability of the immobilized enzyme.

3.6. Substrate specificity of free and immobilized RML

Three different substrates with acyl chain lengths of C8, C12, and C18 were also used to investigate the activity and substrate specificity of free and immobilized preparations of RML by using a pH-stat system at 25 °C and pH 8.0 (Fig. 7). The results show that by increasing the chain length, activity of RML and its immobilized derivatives decreases. The maximum specific activity of 24 890, 17 662, and 14 970 U mg⁻¹ was observed for free RML while using C8, C12, and C18 triacylglycerides as substrate, respectively. Decrease in enzyme activity by increasing the chain length was also observed for the immobilized preparations. Furthermore, the results show that with the same triacylglyceride, the specific activity of RML decreases after immobilization on both supports. The maximum decrease in enzyme activity (43%) of the immobilized preparations was observed for MWCNs-RML in hydrolyzing a C18 triacylglyceride as compared to the free RML.

Fig. 7 The activity and substrate specificity of free and immobilized preparations of RML toward substrates with three different acyl chain lengths.

3.7. Kinetic study and evaluation of optimal pH activity of free and immobilized preparations of RML

The optimum pH for free RML is at pH 7.5, whereas the optimum pH of the Gr-RML and MWCNTs-RML are shifted by 1 and 1.5 units towards the alkaline side, respectively (Fig. 8).

Fig. 8 Optimum pH activity of RML and immobilized preparations.

This increase in optimum pH can be explained by the effect of immobilization on alteration of the microenvironment of the enzyme. A similar increase in the optimum pH value after enzyme immobilization has been previously documented.^23,38

Kinetic parameters of RML and its immobilized derivatives were determined by spectrophotometrically measuring the initial reaction rates for each form with varying amounts of substrate (p-NPB) (Fig. 9).

Fig. 9 Michaelis–Menten plot for the free and immobilized preparations of RML.

K_m values for MWCNTs-RML and Gr-RML were 0.44 and 0.23 mM, respectively which were slightly more than that of the free RML (0.18 mM). The higher values of K_m represent lower affinity between enzymes and substrates. The effect of immobilization in increasing the K_m values can be explained by slightly decreased access of the substrate to the active site of enzyme in the immobilized form.³⁹ The maximum reaction rates (V_max) of the free RML, MWCNTs-RML, and Gr-RML were 0.08, 0.09, and 0.1 mM min⁻¹, respectively. The observed results are in accordance with the values reported by Zhang et al.⁴⁰

4. Conclusion

Covalent immobilization of RML on carboxylated multiwall carbon nanotubes and carboxylated graphene nanosheets was performed via a four-component reaction using an extremely mild condition. This immobilization procedure was performed at room temperature in water and small amounts of offered protein (up to 100 mg); it was accomplished in a very short time (15–30 min) without significant decrease in specific activity of the enzyme. This short immobilization time will maximize the survival rate of particularly unstable proteins. The results showed that the applied approach can be used as a simple, rapid, useful and versatile method for covalent linkage of biomolecules onto carbon-based solid materials. This protocol can also easily be used to conjugate any other useful molecules with amine groups, such as nucleic acids, antibodies, and even chemical compounds on carboxylated solid supports. The leaching experiment confirmed the immobilization of RML on the supports via irreversible covalent linkage. The results showed strongly increased capacity of MWCNTs-COOH, and Gr-COOH for covalent immobilization of enzymes. The immobilized derivatives also showed greatly improved enzyme stability compared to the free enzyme.

Acknowledgements

This work was financially supported by the National Institute of Genetic Engineering and Biotechnology for which the authors are thankful.

References

1. J. L. Porter, R. A. Rusli and D. L. Ollis, ChemBioChem, 2015, 17, 197–203.
2. Z. Habibi, M. Mohammadi and M. Yousefi, Process Biochem., 2013, 48, 669–676.
3. P. Adlercreutz, Chem. Soc. Rev., 2013, 42, 6406–6436.
4. A. A. Homaei, R. Sariri, F. Vianello and R. Stevanato, J. Chem. Biol., 2013, 6, 185–205.
5. M. Babaki, M. Yousefi, Z. Habibi, J. Brask and M. Mohammadi, Biochem. Eng. J., 2015, 101, 23–31.
6. O. Barbosa, C. Ortiz, Á. Berenguer-Murcia, R. Torres, R. C. Rodrigues and R. Fernandez-Lafuente, Biotechnol. Adv., 2015, 33, 435–456.
7. C. Garcia-Galan, Á. Berenguer-Murcia, R. Fernandez-Lafuente and R. C. Rodrigues, Adv. Synth. Catal., 2011, 353, 2885–2904.
8. F. Jia, B. Narasimhan and S. Mallapragada, Biotechnol. Bioeng., 2014, 111, 209–222.
9. Q. Zhang, Z. Zheng, C. Liu, C. Liu and T. Tan, Colloids Surf., B, 2016, 140, 446–451.
10. J. C. S. D. Santos, O. Barbosa, C. Ortiz, A. Berenguer-Murcia, R. C. Rodrigues and R. Fernandez-Lafuente, ChemCatChem, 2015, 7, 2413–2432.
11. K. Min and Y. J. Yoo, Biotechnol. Bioprocess Eng., 2014, 19, 553–567.
12. J. Shen, M. Shi, B. Yan, H. Ma, N. Li, Y. Hu and M. Ye, Colloids Surf., B, 2010, 81, 434–438.
13. J. Zhang, F. Zhang, H. Yang, X. Huang, H. Liu, J. Zhang and S. Guo, Langmuir, 2010, 26, 6083–6085.
14. C. Mateo, J. M. Palomo, G. Fernandez-Lorente, J. M. Guisan and R. Fernandez-Lafuente, Enzyme Microb. Technol., 2007, 40, 1451–1463.
15. S. A. Ansari, R. Satar, S. Chibber and M. J. Khan, J. Mol. Catal. B: Enzym., 2013, 97, 258–263.
16. M. Garmroodi, M. Mohammadi, A. Ramazani, M. Ashjari, J. Mohammadi, B. Sabour and M. Yousefi, Int. J. Biol. Macromol., 2016, 86, 208–215.
17. M. Ashjari, M. Mohammadi and R. Badri, J. Mol. Catal. B: Enzym., 2015, 115, 128–134.
18. M. Mohammadi, Z. Habibi, S. Dezvarei, M. Yousefi, S. Samadi and M. Ashjari, Process Biochem., 2014, 49, 1314–1323.
19. M. Mohammadi, Z. Habibi, S. Dezvarei, M. Yousefi and M. Ashjari, Food Bioprod. Process., 2015, 94, 414–421.
20. J. H. Bartha-Vári, M. I. Toşa, F. D. Irimie, D. Weiser, Z. Boros, B. G. Vértessy, C. Paizs and L. Poppe, ChemCatChem, 2015, 7, 1122–1128.
21. W. Huang, S. Taylor, K. Fu, Y. Lin, D. Zhang, T. W. Hanks, A. M. Rao and Y.-P. Sun, Nano Lett., 2002, 2, 311–314.
22. A. Star, Y. Liu, K. Grant, L. Ridvan, J. F. Stoddart, D. W. Steuerman, M. R. Diehl, A. Boukai and J. R. Heath, Macromolecules, 2003, 36, 553–560.
23. M. Mohammadi, M. Ashjari, S. Dezvarei, M. Yousefi, M. Babaki and J. Mohammadi, RSC Adv., 2015, 5, 32698–32705.
24. R. C. Rodrigues and R. Fernandez-Lafuente, J. Mol. Catal. B: Enzym., 2010, 64, 1–22.
25. M. Noël, P. Lozano and D. Combes, Bioprocess Biosyst. Eng., 2005, 27, 375–380.
26. M. M. Bradford, Anal. Biochem., 1976, 72, 248–254.
27. Z. Habibi, M. Mohammadi and M. Yousefi, Process Biochem., 2013, 48, 669–676.
28. Y. Liu, Q. Li, Y.-Y. Feng, G.-S. Ji, T.-C. Li, J. Tu and X.-D. Gu, Chem. Pap., 2014, 68, 732–738.
29. S. Iijima, Nature, 1991, 354, 56–58.
30. H. Zhou, Y. Qu, C. Kong, D. Li, E. Shen, Q. Ma, X. Zhang, J. Wang and J. Zhou, Colloids Surf., B, 2014, 116, 365–371.
31. P. Asuri, S. S. Karajanagi, E. Sellitto, D. Y. Kim, R. S. Kane and J. S. Dordick, Biotechnol. Bioeng., 2006, 95, 804–811.
32. G. Mechrez, M. A. Krepker, Y. Harel, J.-P. Lellouche and E. Segal, J. Mater. Chem. B, 2014, 2, 915–922.
33. Q. Shi, D. Yang, Y. Su, J. Li, Z. Jiang, Y. Jiang and W. Yuan, J. Nanopart. Res., 2007, 9, 1205–1210.
34. T. A. Saleh, M. Gondal, Q. Drmosh, Z. Yamani and A. Al-Yamani, Chem. Eng. J., 2011, 166, 407–412.
35. C. Lu, F. Su and S. Hu, Appl. Surf. Sci., 2008, 254, 7035–7041.
36. I. Pavlidis, T. Tsoufis, A. Enotiadis, D. Gournis and H. Stamatis, Adv. Eng. Mater., 2010, 12, B179–B183.
37. J. Li, Y.-B. Wang, J.-D. Qiu, D.-C. Sun and X.-H. Xia, Anal. Bioanal. Chem., 2005, 383, 918–922.
38. R. Su, P. Shi, M. Zhu, F. Hong and D. Li, Bioresour. Technol., 2012, 115, 136–140.
39. M. Tu, X. Zhang, A. Kurabi, N. Gilkes, W. Mabee and J. Saddler, Biotechnol. Lett., 2006, 28, 151–156.
40. C. Zhang, S. Luo and W. Chen, Talanta, 2013, 113, 142–147.

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