**3.1 Biocatalysts prepared by adsorption of recombinant lipase onto mesoporous silica**

The enzymatic active component of the developed heterogeneous lipase-active biocatalysts was recombinant *T. lanuginosus* lipase (designated as r*Pichia*/lip) that was produced extracellular by the methylotrophic yeast *Pichia pastoris* X-33 strain specially constructed by the following genetic engineering manipulations: (1) chemical synthesis of the gene of a mature *T. lanuginosus* lipase taking into account the nucleotide sequence found in the Protein Data Bank (PDB-database); (2) cloning of the synthesized gene into a plasmid vector and production of the constructed recombinant plasmid in *E. coli* cells; (3) transformation of competent *P. pastoris* cells with the obtained plasmid and selection of recombinant yeast clones; and (4) analysis of the selected clones for the ability to produce extracellular and secrete recombinant *T. lanuginosus* lipase into the nutrient culture medium. Finally, conditions for the cultivation and intensive growth of the r*Pichia*/lip strain-producer were optimized in order to increase the concentration of the secreted target enzyme. The lab scale production of recombinant lipase was carried out in a 10-liter BIOK gas-vortex bioreactor (ZAO Sayany, Russia); the lipase concentration in nutrient media reached 2 g/L. Partial purification of recombinant lipase was carried out by precipitation of the secreted r*Pichia*/lip with ammonium sulfate (up to 75% saturation) at 4°C for 16 h. The precipitates were dissolved in distilled water, and further dialysis against a 25 mM acetate buffer pH 4.0 was carried out. The samples of dialyzed and lyophilized r*Pichia*/lip were used for the preparing lipase-active biocatalysts.

Mesoporous silica (SiO2) of KSK™ type was used as a support for immobilization of recombinant *T. lanuginosus* lipase; its textural parameters are as follows: the specific surface area 157 m<sup>2</sup> /g; total pore volumes (VΣ) 0.76 mL/g; average pore diameters 19 nm; and porosity 58%. The lipase-active biocatalysts were prepared by both "spontaneous" and "forcible" adsorption using r*Pichia*/lip solutions at the same concentration. Spontaneous adsorption was carried out while contacting support with lipase solution at a ratio of support weight to solution volume of 1:(3–10) for 24–48 h. For such a long contact, adsorptive immobilization of lipase occurred due to electrostatic and acid-base interactions of enzyme molecules with silica surface. Forcible adsorption of lipase was carried out via moisture capacity impregnation of mesoporous silica by lipase solution, followed by drying granules. When biocatalyst was dried, enzyme molecules lost their hydrated shells and stuck to the surface. In both cases, the amounts of adsorbed enzyme were close in magnitude and equal to 4.2 and 5.4 mg/g for spontaneous and forcible adsorption, respectively. It was found that under exactly the same reaction conditions of esterification and synthesis of *n-*butyl heptanoate, the stationary activity of the spontaneously adsorbed lipase was two-fold less than the activity of forcibly adsorbed enzyme, for example, 2.4 vs. 5.0 U/g, respectively (recall that 1 U = 1 μmol/min). The method of forcible adsorption was characterized not only by comparatively high enzymatic

*Heterogeneous Biocatalysts for the Final Stages of Deep Processing of Renewable Resources… DOI: http://dx.doi.org/10.5772/intechopen.89411*

activity but also by simplicity of its implementation and economical enzyme consumption. For example, a minimal volume of lipase solution used for spontaneous adsorption on 1 g of silica was 3.0 mL, whereas for forcible adsorption, it was 0.8 mL equal to total pore volume (VΣ) of SiO2. All results described here referred to the lipase-active heterogeneous biocatalysts (designated as LipoSil) prepared by forcible adsorption on silica of recombinant lipase, which were used predominantly in esterification processes [15, 16, 18].

Biocatalytic processes of enzymatic esterification were performed at ambient conditions (20 2°C, 1 bar) in unconventional anhydrous media of organic solvents such as hexane and diethyl ether. The saturated fatty acids differing in the number of carbon atoms (C2–C10, C18), as well as aliphatic alcohols differing in the structure of the molecules, namely, the number of carbon atoms (C2–12, C16), the isomerism of the carbon skeleton (*n*- and *iso*-) and OH-group position (*prim-, sec-,* and *tert-*) were studied as substrates in esterification by LipoSil. There were some peculiarities of operation of the preliminary dried lipase-active biocatalysts in nonaqueous organic solvents. A considerable increase of the activity was observed during the 1st–3rd reaction cycles. This phenomena, named preconditioning stage, was due to the ongoing accumulation of formed product—water—in the vicinity of the adsorbed lipase inside the silica-based biocatalyst, and this stage proceeded faster and the higher than the activity of the biocatalyst. For example, if the biocatalytic activities were about 5 and 500 U/g, then the activation of the biocatalysts (preconditioning) proceeded within 24 and 0.5 h, respectively. Calculation showed that under studied conditions upon full conversion of fatty acid, maximal 0.1 mL of water was formed inside for one reaction cycle. Since the total pore volume of silica (0.8 mL/g) was multifold greater than the volume of the water formed, this amount of H2O was firmly held inside KSK™ silica commonly applied as a dehumidifier for industrial gases. Therefore, during esterification, the favorable aqua microenvironment was created for adsorbed lipase, and the activities of the dried biocatalysts increased by 2–4 times. After preconditioning stage, the biocatalytic activity, named stationary, was measured in batch reactor during several tens of reaction cycles. Each reaction cycle was completed to the full conversion of acid, close to 85– 90%. Then, the reaction medium was removed by decantation and the biocatalysts were washed by solvent for 20 h. The next reaction cycle was started by adding fresh reaction medium containing substrates of lipase—acid S1 and double molar excess of alcohol S2. Stationary activity of the biocatalysts was fluctuated in magnitude during the consecutive reaction cycles of the periodic esterification process, perhaps due to the presence of ester (product) residues inside the operating biocatalyst. As it can be seen in **Figure 3**, the operational stability of the prepared biocatalyst was sufficiently high; its stationary activity was retained completely after 38 cycles (900 h) of esterification of various fatty acid. Also, the biocatalysts possessed a high long-term stability; the activity was determined to be 80% of initial one after storage for 9 months in the solvent (hexane and diethyl ether) at ambient temperature and in dried state in refrigerator.

Obviously, a very important property of heterogeneous biocatalysts is the high operational stability, since the productivity calculated by multiplying average activity by 2t½ increases significantly. The esterifying activity of the prepared lipase-active biocatalysts did not practically change during 500–1000 h of operation. Under studded conditions, the productivity was evaluated as 2 tons of product per 1 kg of LipoSil biocatalyst.

The study of the functional properties of enzymes after their immobilization, such as activity, stability, and, importantly, specificity, is of great interest. Of particular scientific and practical interest is the research of the possibility of modulating these properties, by engineering heterogeneous biocatalysts, in particular, by

recombinant *T. lanuginosus* lipase. These lipase-active biocatalysts were prepared both by entrapment of fully disrupted cells (lysates) of recombinant strainproducer r*E*.*coli*/lip inside silica xerogel and its nanocarbon-containing composites [13, 14, 17] and by adsorption of *T. lanuginosus* lipase produced by recombinant lipase on silica [15–18] and carbon aerogel [19]. These biocatalysts were studied in the reactions of tributyrin hydrolysis [13, 19], interesterification of oil-fat blends and vegetable oil triglycerides with ethyl acetate [13, 14], and esterification of fatty acids [15–19]. The two biocatalysts prepared by adsorptive immobilization of

recombinant *T. lanuginosus* lipase are described briefly below.

**mesoporous silica**

*Molecular Biotechnology*

biocatalysts.

**10**

specific surface area 157 m<sup>2</sup>

**3.1 Biocatalysts prepared by adsorption of recombinant lipase onto**

The enzymatic active component of the developed heterogeneous lipase-active biocatalysts was recombinant *T. lanuginosus* lipase (designated as r*Pichia*/lip) that was produced extracellular by the methylotrophic yeast *Pichia pastoris* X-33 strain specially constructed by the following genetic engineering manipulations: (1) chemical synthesis of the gene of a mature *T. lanuginosus* lipase taking into account the nucleotide sequence found in the Protein Data Bank (PDB-database); (2) cloning of the synthesized gene into a plasmid vector and production of the constructed recombinant plasmid in *E. coli* cells; (3) transformation of competent *P. pastoris* cells with the obtained plasmid and selection of recombinant yeast clones; and (4) analysis of the selected clones for the ability to produce extracellular and secrete recombinant *T. lanuginosus* lipase into the nutrient culture medium. Finally, conditions for the cultivation and intensive growth of the r*Pichia*/lip strain-producer were optimized in order to increase the concentration of the secreted target

enzyme. The lab scale production of recombinant lipase was carried out in a 10-liter BIOK gas-vortex bioreactor (ZAO Sayany, Russia); the lipase concentration in nutrient media reached 2 g/L. Partial purification of recombinant lipase was carried out by precipitation of the secreted r*Pichia*/lip with ammonium sulfate (up to 75% saturation) at 4°C for 16 h. The precipitates were dissolved in distilled water, and further dialysis against a 25 mM acetate buffer pH 4.0 was carried out. The samples of dialyzed and lyophilized r*Pichia*/lip were used for the preparing lipase-active

Mesoporous silica (SiO2) of KSK™ type was used as a support for immobilization of recombinant *T. lanuginosus* lipase; its textural parameters are as follows: the

diameters 19 nm; and porosity 58%. The lipase-active biocatalysts were prepared by both "spontaneous" and "forcible" adsorption using r*Pichia*/lip solutions at the same concentration. Spontaneous adsorption was carried out while contacting support with lipase solution at a ratio of support weight to solution volume of 1:(3–10) for 24–48 h. For such a long contact, adsorptive immobilization of lipase occurred due to electrostatic and acid-base interactions of enzyme molecules with silica surface. Forcible adsorption of lipase was carried out via moisture capacity impregnation of mesoporous silica by lipase solution, followed by drying granules. When biocatalyst was dried, enzyme molecules lost their hydrated shells and stuck to the surface. In both cases, the amounts of adsorbed enzyme were close in magnitude and equal to 4.2 and 5.4 mg/g for spontaneous and forcible adsorption, respectively. It was found that under exactly the same reaction conditions of esterification and synthesis of *n-*butyl heptanoate, the stationary activity of the spontaneously adsorbed lipase was two-fold less than the activity of forcibly adsorbed enzyme, for example, 2.4 vs. 5.0 U/g, respectively (recall that 1 U = 1 μmol/min). The method of forcible adsorption was characterized not only by comparatively high enzymatic

/g; total pore volumes (VΣ) 0.76 mL/g; average pore

#### **Figure 3.**

*Operational stability of the lipase-active biocatalyst in periodic esterification of various fatty acids such as butyric (C4:0), enanthic (C7:0), capric (C10:0), and stearic (C18:0) with n-butanol as a function of enzymatic activities depending on a number of reaction cycle. Conditions of esterification: 20 2°C, 0.25 M acid, 0.50 M alcohol, hexane:diethyl ether = 1:1, content of biocatalyst in reaction medium 20.8 wt.%. The amount of adsorbed lipase is 14.0 mg/g.*

that of aliphatic alcohols because the maximal differences in rates were �6 times for

*Relative activities of the immobilized recombinant T. lanuginosus lipase in reaction of esterification of (a) different saturated fatty acids with n-butanol, and (b) enanthic acid with different alcohols. Conditions of esterification: 20* � *2°C, 0.25 M acid, 0.50 M alcohol, hexane:diethyl ether = 1:1, content of biocatalyst in*

*Heterogeneous Biocatalysts for the Final Stages of Deep Processing of Renewable Resources…*

It was found that immobilized on silica r*Pichia*/lip was not sensitive to isomerism of alcohols' molecule, and the rates of esterification of fatty acids with primary *n-*C4,C5 alcohols were almost 10 � 4% lower than those with *is0*-C4,C5 alcohols [18]. The most pronounced specificity was observed toward the position of OHgroup in alcohol molecules. The rates of esterification of fatty acids with secondary alcohol (*sec-*propanol and *sec-*butanol) was two order of magnitude lower than those

various acids (**Figure 4a**) and only �2 times for alcohols (**Figure 4b**).

*reaction medium 20.8 wt.% the amount of adsorbed lipase 14.0 mg/g.*

*DOI: http://dx.doi.org/10.5772/intechopen.89411*

with primary alcohols, for example, 0.36 U/g vs. 56.1 U/g for *sec-* and *prim*propanol, respectively [18]. The esterification of fatty acid with ternary alcohols (*tert*-butanol) did not occur, and reaction rate was zero. Also, it was found that the substrates (both acid and alcohol) with aromatic and cyclic restudies reacted with

�1

were 0.22 � 0.05 mol/L and 66.7 � 4.0 <sup>μ</sup>mol/L s�<sup>1</sup>

The kinetic parameters such as Michaelis constant (KM) for acid and maximal reaction rate (Vmax) under studied conditions were determined in esterification of enanthic (heptanoic, C7:0) acid with a double excess of *n-*butanol. The kinetic curve as a function of the initial reaction rate from the initial fatty acid concentration lower than 1.0 M was satisfactory approximated by the classic hyperbolic Michaelis equa-

KM is the Michaelis constant, mol/L. At a concentration of enanthic acid above 1.0 M, the reaction rate decreased perhaps due to inactivation of enzyme by high concentration of *n-*butanol as described previously for esterification of capric acid with *iso*pentanol [17]. The values of KM and Vmax determined using Lineweaver-Burk linear approximation and the regression of hyperbolic equation by soft Origin' programs

data published earlier in [16, 17] showed that the Michaelis constant for enanthic (C7) acid was �3 times less than KM for capric (C10) acid during esterification with C4,C5 aliphatic alcohols. This means, that the affinity of adsorbed recombinant lipase toward enanthic acid is higher; thus, the maximum rate Vmax will be achieved

The catalytic properties of the lipase-active heterogeneous biocatalysts such as enzymatic activity, stability, and substrates specificity were investigated in the

at lower initial fatty acid concentrations, and it is important for practice.

, where V and Vmax are the initial observed and maximum reaction

; C0 is the initial substrate concentration, mol/L; and

, respectively. A comparison with

very low rates [15, 19].

KMþC0

rates, respectively, μmol/L�s

**3.2 Conclusion for the part 3.1**

tion, V <sup>¼</sup> Vmax�C0

**13**

**Figure 4.**


#### **Table 1.**

*Matrix of relative activities of the immobilized recombinant T. lanuginosus lipase in reaction of esterification of various pairs of substrates—saturated fatty acids and aliphatic primary n-alcohols.*

selecting the chemical nature of the supports [20, 21]. In our research, the specificity of heterogeneous enzymatic esterification was determined by comparing the reaction rates for various pairs of substrates—saturated fatty acids and aliphatic primary *n-*alcohols—and the matrix of relative units of activities for the LipoSil biocatalyst was composed (**Table 1**) [18].

The following conclusions can be drawn from the data presented in **Table 1**: (1) the rate of esterification of acetic (C2) acid is very low; thus, it is practically impossible to obtain acetate esters; (2) the rate of synthesis of ethyl ester (reaction with ethanol C2) is 1.3–2 times lower than that in reactions with C3–C16 alcohols; (3) the esterification of fatty acids with a number of carbon atoms more than six (C6) occurs 2–5 times faster than that of low molecular weight fatty acids C4–C6; and (4) the maximal observed rate is determined for esterification of enanthic (C7) acid with C3–C8 alcohols. Also, one can see that the immobilized on silica recombinant lipase is more sensitive to the molecular structure of saturated fatty acids than *Heterogeneous Biocatalysts for the Final Stages of Deep Processing of Renewable Resources… DOI: http://dx.doi.org/10.5772/intechopen.89411*

**Figure 4.**

*Relative activities of the immobilized recombinant T. lanuginosus lipase in reaction of esterification of (a) different saturated fatty acids with n-butanol, and (b) enanthic acid with different alcohols. Conditions of esterification: 20* � *2°C, 0.25 M acid, 0.50 M alcohol, hexane:diethyl ether = 1:1, content of biocatalyst in reaction medium 20.8 wt.% the amount of adsorbed lipase 14.0 mg/g.*

that of aliphatic alcohols because the maximal differences in rates were �6 times for various acids (**Figure 4a**) and only �2 times for alcohols (**Figure 4b**).

It was found that immobilized on silica r*Pichia*/lip was not sensitive to isomerism of alcohols' molecule, and the rates of esterification of fatty acids with primary *n-*C4,C5 alcohols were almost 10 � 4% lower than those with *is0*-C4,C5 alcohols [18]. The most pronounced specificity was observed toward the position of OHgroup in alcohol molecules. The rates of esterification of fatty acids with secondary alcohol (*sec-*propanol and *sec-*butanol) was two order of magnitude lower than those with primary alcohols, for example, 0.36 U/g vs. 56.1 U/g for *sec-* and *prim*propanol, respectively [18]. The esterification of fatty acid with ternary alcohols (*tert*-butanol) did not occur, and reaction rate was zero. Also, it was found that the substrates (both acid and alcohol) with aromatic and cyclic restudies reacted with very low rates [15, 19].

The kinetic parameters such as Michaelis constant (KM) for acid and maximal reaction rate (Vmax) under studied conditions were determined in esterification of enanthic (heptanoic, C7:0) acid with a double excess of *n-*butanol. The kinetic curve as a function of the initial reaction rate from the initial fatty acid concentration lower than 1.0 M was satisfactory approximated by the classic hyperbolic Michaelis equation, V <sup>¼</sup> Vmax�C0 KMþC0 , where V and Vmax are the initial observed and maximum reaction rates, respectively, μmol/L�s �1 ; C0 is the initial substrate concentration, mol/L; and KM is the Michaelis constant, mol/L. At a concentration of enanthic acid above 1.0 M, the reaction rate decreased perhaps due to inactivation of enzyme by high concentration of *n-*butanol as described previously for esterification of capric acid with *iso*pentanol [17]. The values of KM and Vmax determined using Lineweaver-Burk linear approximation and the regression of hyperbolic equation by soft Origin' programs were 0.22 � 0.05 mol/L and 66.7 � 4.0 <sup>μ</sup>mol/L s�<sup>1</sup> , respectively. A comparison with data published earlier in [16, 17] showed that the Michaelis constant for enanthic (C7) acid was �3 times less than KM for capric (C10) acid during esterification with C4,C5 aliphatic alcohols. This means, that the affinity of adsorbed recombinant lipase toward enanthic acid is higher; thus, the maximum rate Vmax will be achieved at lower initial fatty acid concentrations, and it is important for practice.

#### **3.2 Conclusion for the part 3.1**

The catalytic properties of the lipase-active heterogeneous biocatalysts such as enzymatic activity, stability, and substrates specificity were investigated in the

selecting the chemical nature of the supports [20, 21]. In our research, the specificity of heterogeneous enzymatic esterification was determined by comparing the reaction rates for various pairs of substrates—saturated fatty acids and aliphatic primary *n-*alcohols—and the matrix of relative units of activities for the LipoSil

*Matrix of relative activities of the immobilized recombinant T. lanuginosus lipase in reaction of esterification of*

*various pairs of substrates—saturated fatty acids and aliphatic primary n-alcohols.*

*Operational stability of the lipase-active biocatalyst in periodic esterification of various fatty acids such as butyric (C4:0), enanthic (C7:0), capric (C10:0), and stearic (C18:0) with n-butanol as a function of enzymatic activities depending on a number of reaction cycle. Conditions of esterification: 20 2°C, 0.25 M acid, 0.50 M alcohol, hexane:diethyl ether = 1:1, content of biocatalyst in reaction medium 20.8 wt.%. The*

The following conclusions can be drawn from the data presented in **Table 1**: (1) the rate of esterification of acetic (C2) acid is very low; thus, it is practically impossible to obtain acetate esters; (2) the rate of synthesis of ethyl ester (reaction with ethanol C2) is 1.3–2 times lower than that in reactions with C3–C16 alcohols; (3) the esterification of fatty acids with a number of carbon atoms more than six (C6) occurs 2–5 times faster than that of low molecular weight fatty acids C4–C6; and (4) the maximal observed rate is determined for esterification of enanthic (C7) acid with C3–C8 alcohols. Also, one can see that the immobilized on silica recombinant lipase is more sensitive to the molecular structure of saturated fatty acids than

biocatalyst was composed (**Table 1**) [18].

**Figure 3.**

*Molecular Biotechnology*

**Table 1.**

**12**

*amount of adsorbed lipase is 14.0 mg/g.*

esterification of various saturated fatty acids with aliphatic alcohols. These biocatalysts were prepared by immobilizing recombinant *T. lanuginosus* lipase onto mesoporous silica by impregnation method. Particular attention was paid to the study of the substrate specificity of immobilized lipase. It was found that forcibly adsorbed on silica *T. lanuginosus* lipase demonstrated broad substrate specificity. Saturated fatty acids with a number of carbon atoms 6 or more (till 18) and alcohols with a number of carbon atoms 3 or more (till 16) reacted with comparatively high reaction rates. The reaction rates depended slightly on isomerism (*n*- and *iso*-) of carbon skeleton of C4-C5 alcohols, whereas the rates depended strongly on the position of OH-group, and secondary and ternary alcohols did not react with fatty acids. Comparing the rates of esterification of various pairs of substrates using primary alcohols, a matrix of relative biocatalytic activities was composed. According to this matrix, the rates of synthesis of C4-C18 esters were sufficiently high, while the rate of synthesis of acetate ester was very low. Under the same reaction conditions, the maximal rate was observed in esterification of enanthic (C7:0) acid with butanol. The classical Michaelis-Menten kinetics was inherent for biocatalytic esterification of this acid with double molar excess of alcohol, and the main kinetic parameter, Michaelis constant for acid, was determined to be 0.22.mol/L.

It is well known that, from a practical point of view, the stability of heterogeneous biocatalysts is a very important characteristic that determines the biocatalyst productivity. The prepared lipase-active biocatalysts possessed considerably high operational stability in a periodic batch process of low-temperature esters'synthesis carried out in unconventional anhydrous media of organic solvents (hexane and diethyl ether). The enzymatic activity of the biocatalysts was completely retained for several tens of reaction cycles.

It was concluded that due to the remarkable catalytic properties combined with simplicity of immobilization method, the prepared lipase-active biocatalysts are promising for practical use in organic synthesis including the production of valuable esters.

#### **3.3 Biocatalysts prepared by adsorption of lipase on carbon aerogel**

Immobilization of recombinant *T. lanuginosus* lipase (SIGMA Co.) was carried out by spontaneous adsorption onto macroporous carbon aerogel [19]. It should be noted that carbon aerogels are a unique class of porous materials with a very low density (less than 0.1 g/mL) and a porosity of up to 90–99%. These materials are novel promising adsorbents for enzyme immobilization with great potential for practical implementation. Macroporous carbon aerogel (MCA) for our research has been produced by *in situ* synthesis of multi-walled carbon nanotubes (CNTs) via catalytic high-temperature decomposition of ethylene over the supported Fe:Co catalyst. The carbon aerogel was obtained in the form of ball-shaped granules of 1–10 mm in diameter (**Figure 5a**).

The three-dimensional framework and rigid macrostructure of MCA were formed by chaotic interlacing carbon nanotubes (**Figure 6a**). The number of walls and the diameter of CNTs determined by high resolution transmission electron microscopy (HRTEM) were equal to 12–14 and 15–25 nm respectively (**Figure 6b**). The density of MCA was 0.06 g/mL. The texture parameters of carbon aerogel were as follows: the specific surface area (SBET) was 80–110 m<sup>2</sup> /g, total pore volume (VΣ) was 10-14 mL/g; macropores of 0.5–1 μm in diameter were predominant in texture; and the volumes of meso- and micropores did not exceed 2% of VΣ.

the 2nd plateau (**Figure 7**). Considering the molecular weight of the lipase, 30,000 Da, we calculated the surface concentration of the adsorbed lipase,

*lipase. Conditions of adsorption: 20 2°C, 0.02 M Na-phosphate buffer pH 7.0, 24 h.*

estimated to be 20 nm as confirmed by HRTEM images [19]. It is known from literature [22] and PDB-database, that crystallographic size of the lipase molecule is 3.5 4.5 5.0 nm; in aqueous media, the diameters of hydrated molecules increase twice. The *T. lanuginosus* lipase has the ability to be associated into dimers due to hydrophobic interaction between hydrophobic lids and pockets of active site [23]. So, the first plateau corresponded to the formation of the 1st adsorptive layer that was dense and uniform in distribution of the adsorbed lipase on the surface of MCA. The second plateau can be attributed to the 2nd also dense adsorptive layer formed

*Adsorption of recombinant lipase on macroporous carbon aerogel depending on initial concentration of soluble*

*(a) Photo of granules of macroporous carbon aerogel and (b) photo of agglomeration of fine dispersed carbon*

*Heterogeneous Biocatalysts for the Final Stages of Deep Processing of Renewable Resources…*

*(a) SEM image of the inside of the granules of macroporous carbon aerogel; (b) HRTEM image of carbon*

*nanotubes (CNTs) under the influence of increasing concentration of tributyrin.*

*DOI: http://dx.doi.org/10.5772/intechopen.89411*

, and then the area occupied by one molecule of the

. The diameter of one adsorbed molecule of lipase was

<sup>2</sup> <sup>10</sup><sup>15</sup> molecules per 1 m<sup>2</sup>

**Figure 5.**

**Figure 6.**

**Figure 7.**

**15**

*nanotubes forming MCA.*

adsorbed lipase, 500 nm<sup>2</sup>

When studying adsorption of recombinant lipase on MCA, it was found that adsorption graph contained two "plateaus" corresponding to adsorption of 100 mg/g, or 0.8 mg/m<sup>2</sup> of carbon aerogel for the 1st plateau, and 200 mg/g for *Heterogeneous Biocatalysts for the Final Stages of Deep Processing of Renewable Resources… DOI: http://dx.doi.org/10.5772/intechopen.89411*

#### **Figure 5.**

esterification of various saturated fatty acids with aliphatic alcohols. These

0.22.mol/L.

*Molecular Biotechnology*

valuable esters.

**14**

for several tens of reaction cycles.

1–10 mm in diameter (**Figure 5a**).

biocatalysts were prepared by immobilizing recombinant *T. lanuginosus* lipase onto mesoporous silica by impregnation method. Particular attention was paid to the study of the substrate specificity of immobilized lipase. It was found that forcibly adsorbed on silica *T. lanuginosus* lipase demonstrated broad substrate specificity. Saturated fatty acids with a number of carbon atoms 6 or more (till 18) and alcohols with a number of carbon atoms 3 or more (till 16) reacted with comparatively high reaction rates. The reaction rates depended slightly on isomerism (*n*- and *iso*-) of carbon skeleton of C4-C5 alcohols, whereas the rates depended strongly on the position of OH-group, and secondary and ternary alcohols did not react with fatty acids. Comparing the rates of esterification of various pairs of substrates using primary alcohols, a matrix of relative biocatalytic activities was composed. According to this matrix, the rates of synthesis of C4-C18 esters were sufficiently high, while the rate of synthesis of acetate ester was very low. Under the same reaction conditions, the maximal rate was observed in esterification of enanthic (C7:0) acid with butanol. The classical Michaelis-Menten kinetics was inherent for biocatalytic esterification of this acid with double molar excess of alcohol, and the main kinetic parameter, Michaelis constant for acid, was determined to be

It is well known that, from a practical point of view, the stability of heterogeneous biocatalysts is a very important characteristic that determines the biocatalyst productivity. The prepared lipase-active biocatalysts possessed considerably high operational stability in a periodic batch process of low-temperature esters'synthesis carried out in unconventional anhydrous media of organic solvents (hexane and diethyl ether). The enzymatic activity of the biocatalysts was completely retained

It was concluded that due to the remarkable catalytic properties combined with simplicity of immobilization method, the prepared lipase-active biocatalysts are promising for practical use in organic synthesis including the production of

Immobilization of recombinant *T. lanuginosus* lipase (SIGMA Co.) was carried out by spontaneous adsorption onto macroporous carbon aerogel [19]. It should be noted that carbon aerogels are a unique class of porous materials with a very low density (less than 0.1 g/mL) and a porosity of up to 90–99%. These materials are novel promising adsorbents for enzyme immobilization with great potential for practical implementation. Macroporous carbon aerogel (MCA) for our research has been produced by *in situ* synthesis of multi-walled carbon nanotubes (CNTs) via catalytic high-temperature decomposition of ethylene over the supported Fe:Co catalyst. The carbon aerogel was obtained in the form of ball-shaped granules of

The three-dimensional framework and rigid macrostructure of MCA were formed by chaotic interlacing carbon nanotubes (**Figure 6a**). The number of walls and the diameter of CNTs determined by high resolution transmission electron microscopy (HRTEM) were equal to 12–14 and 15–25 nm respectively (**Figure 6b**). The density of MCA was 0.06 g/mL. The texture parameters of carbon aerogel were

was 10-14 mL/g; macropores of 0.5–1 μm in diameter were predominant in texture;

When studying adsorption of recombinant lipase on MCA, it was found that

100 mg/g, or 0.8 mg/m<sup>2</sup> of carbon aerogel for the 1st plateau, and 200 mg/g for

/g, total pore volume (VΣ)

as follows: the specific surface area (SBET) was 80–110 m<sup>2</sup>

and the volumes of meso- and micropores did not exceed 2% of VΣ.

adsorption graph contained two "plateaus" corresponding to adsorption of

**3.3 Biocatalysts prepared by adsorption of lipase on carbon aerogel**

*(a) Photo of granules of macroporous carbon aerogel and (b) photo of agglomeration of fine dispersed carbon nanotubes (CNTs) under the influence of increasing concentration of tributyrin.*

#### **Figure 6.**

*(a) SEM image of the inside of the granules of macroporous carbon aerogel; (b) HRTEM image of carbon nanotubes forming MCA.*

#### **Figure 7.**

*Adsorption of recombinant lipase on macroporous carbon aerogel depending on initial concentration of soluble lipase. Conditions of adsorption: 20 2°C, 0.02 M Na-phosphate buffer pH 7.0, 24 h.*

the 2nd plateau (**Figure 7**). Considering the molecular weight of the lipase, 30,000 Da, we calculated the surface concentration of the adsorbed lipase, <sup>2</sup> <sup>10</sup><sup>15</sup> molecules per 1 m<sup>2</sup> , and then the area occupied by one molecule of the adsorbed lipase, 500 nm<sup>2</sup> . The diameter of one adsorbed molecule of lipase was estimated to be 20 nm as confirmed by HRTEM images [19]. It is known from literature [22] and PDB-database, that crystallographic size of the lipase molecule is 3.5 4.5 5.0 nm; in aqueous media, the diameters of hydrated molecules increase twice. The *T. lanuginosus* lipase has the ability to be associated into dimers due to hydrophobic interaction between hydrophobic lids and pockets of active site [23]. So, the first plateau corresponded to the formation of the 1st adsorptive layer that was dense and uniform in distribution of the adsorbed lipase on the surface of MCA. The second plateau can be attributed to the 2nd also dense adsorptive layer formed

probably by further dimerization of lipase molecules. As it turned out, under studied conditions, the lipase adsorbed molecules formed two dense adsorptive layers; the amount of lipase in each adsorptive layer was equal to 100 mg per 1 g of MCA (**Figure 7**).

It was found that 85–90% of the amount of adsorbed lipase was very firmly attached to the surface of carbon aerogel and did not desorb [19]. Desorption was investigated using various selective reagents in order to clarify mechanism of lipase binding on carbon surface. For example, the amount of lipase desorbed using 1 M sodium chloride did not exceed 2%; hence, electrostatic interactions were negligible. The most effective desorption, 10 2%, was observed using distilled water and C2-C4 alcohols'solutions; hence, weak Van der Waals interactions were predominant. The lipase was also efficiently desorbed by emulsifier (gum arabic) perhaps due to breaking hydrophobic interactions. Thus, study on selective desorption showed that the strong adsorption of *T. lanuginosus* lipase on carbon nanotubes forming aerogel occurred exclusively due to hydrophobic-hydrophobic interactions.

Lipase-active biocatalysts prepared by adsorptive immobilization of recombinant *T. lanuginosus* lipase on macroporous carbon aerogel were studied in the periodic processes of bioconversion of triglycerides and fatty acids such as tributyrin hydrolysis, interesterification of vegetable oil with ethyl acetate, and esterification of saturated fatty acids (butyric C4:0, capric C10:0, and stearic C18:0) with *iso*-pentanol.

surface of carbon nanotubes, namely correctly and incorrectly (**Figure 8b**). The incorrect orientation (No. 1 in **Figure 8b**) with blocking the active site by the surface was realized more likely in the incompact 1st adsorptive layer; in this case, the observed lipase activity had a minimum value. The correct orientation (No. 3 in **Figure 8b**) may be realized in the dense 1st adsorptive layer, so the formation of active enzyme-substrate complex occurred; and in this case, the observed lipase activity had a maximum value. A further decrease in specific activities after the formation of the 1st adsorptive layer was due to covering the active lipase by the molecules in subsequent 2nd adsorptive layer (scheme in **Figure 7**) and possible partially correct orientation (No. 2 in **Figure 8b**). The obtained data suggest that the main reasons for a loss of enzymatic activity upon adsorption of the lipase on hydrophobic carbon aerogel are as follows: (1) incorrect orientation of enzyme molecule on the carbon surface and blockage of the active site, as well as possible deformation of enzyme molecule; and (2) dehydration of the biocatalysts due to removal of essential water from the vicinity of adsorbed lipase via efficient adsorp-

*Specific hydrolytic activity of lipase adsorbed on carbon aerogel depending on adsorption. Conditions of hydrolysis: 20 2°C, 0.02 M Na-phosphate buffer pH 7.0, 0.02 M tributyrin, 1.0 M glycerol, 0.6% gum*

*Heterogeneous Biocatalysts for the Final Stages of Deep Processing of Renewable Resources…*

It has been found that activity and stability of the prepared lipase-active hetero-

As mentioned above, esterification of acid with alcohols was accompanied by the formation of water and the corresponding ester, and, as a result, accumulation of essential water molecules inside biocatalysts in the vicinity of the adsorbed lipase occurred. So, the stability of the prepared biocatalysts was much higher in esterification than in hydrolysis and interesterification. And the biocatalysts operated during esterification without a loss of activity for more than several hundred hours in the nonaqueous media of anhydrous organic solvents. As seen in **Figure 9**, the

performed either in aqueous reaction media (hydrolysis) or in nonaqueous media (interesterification and esterification). The stability of the biocatalysts was quite low when triglycerides participated in reactions as lipase substrates. For example, the biocatalysts lost 90% of initial activity during six reaction cycles of tributyrin hydrolysis [19]. Another example, the stability of adsorbed on MCA r*Pichia*/lip was determined during interesterification of linseed oil with ethyl acetate in order to produce the valuable product, vitamin F—ethyl esters of ω-3 fatty acids. The conversion of triglycerides was 87 and 66% in the 1st and 3–5th reaction cycles, respec-

geneous biocatalysts depended strongly on the type of enzymatic reaction

tively, and then biocatalysts inactivated due to rapid dehydration.

tion of hydrophobic triglycerides.

*arabic, content of biocatalyst in reaction medium 3.3 wt.%.*

*DOI: http://dx.doi.org/10.5772/intechopen.89411*

**Figure 8.**

**17**

The activity of the lipase-active biocatalysts and specific activity of adsorbed on MCA lipase were measured in hydrolysis of emulsified tributyrin under conditions when there were no diffusion limitations for mass transfer of the substrate from reaction media toward the adsorbed lipase [19]. Note that specific activity of adsorbed enzyme (in U/mg) is calculated by dividing the experimentally observed activity of the biocatalyst (in U/g) by the amount of adsorbed enzyme (in mg/g). The specific activities of the lipase adsorbed both on MCA granules and nonporous fine powders of carbon nanotubes were compared with each other; the values were found to be 160 and 12 U/mg, respectively. This significant difference was due to the relative rigidity of ball-shaped aerogel granules that prevented agglomeration of carbon nanotubes in "oil-in-water" triglyceride emulsion as presented in **Figure 5b**. Since water-immiscible hydrophobic triglyceride molecules were adsorbed efficiently on hydrophobic carbon nanotubes, with increasing tributyrin concentration, agglomerates of round shape containing this substrate were formed from CNTs (**Figure 5b**). As a result, the required hydrolysis water is displaced from the vicinity of adsorbed lipase and its specific activity fell down as mentioned above.

The maximal hydrolytic specific activity of the adsorbed *T. lanuginosus* lipase was measured to be 700 U/mg (vs. 14,000 U/mg for soluble lipase), i.e., activity significantly decreased upon adsorption of lipase on MCA. Another reason discussed below may be probably incorrect orientation of adsorbed lipase on a highly hydrophobic carbon surface of CNTs.

The results of study on dependence of the specific activity of adsorbed lipase on the adsorption value are presented in **Figure 8a**. Initially, when adsorption of the lipase was small, the observed activities were extremely low. The specific activity increased dramatically and reached maximal values, >700 U/mg, in a region close to the formation of the 1st protein dense layer at adsorption of 100–110 mg/g (**Figure 8a**). Then, the specific activity fell down as adsorption increased (**Figure 8a**). As a result, maximal hydrolytic activity of the biocatalysts, 75,000 U/ g, was observed at the adsorption corresponding to the formation of the 1st protein layer. The activity of the biocatalyst with the double adsorptive layer was found to be 1.6-fold lower, 47,000 U/g. In order to explain maximum on the curve in **Figure 8a**, we proposed that the adsorbed lipase can be oriented differently on the

*Heterogeneous Biocatalysts for the Final Stages of Deep Processing of Renewable Resources… DOI: http://dx.doi.org/10.5772/intechopen.89411*

**Figure 8.**

probably by further dimerization of lipase molecules. As it turned out, under studied conditions, the lipase adsorbed molecules formed two dense adsorptive layers; the amount of lipase in each adsorptive layer was equal to 100 mg per 1 g of MCA

It was found that 85–90% of the amount of adsorbed lipase was very firmly attached to the surface of carbon aerogel and did not desorb [19]. Desorption was investigated using various selective reagents in order to clarify mechanism of lipase binding on carbon surface. For example, the amount of lipase desorbed using 1 M sodium chloride did not exceed 2%; hence, electrostatic interactions were negligible. The most effective desorption, 10 2%, was observed using distilled water and C2-C4 alcohols'solutions; hence, weak Van der Waals interactions were predominant. The lipase was also efficiently desorbed by emulsifier (gum arabic) perhaps due to breaking hydrophobic interactions. Thus, study on selective desorption showed that the strong adsorption of *T. lanuginosus* lipase on carbon nanotubes forming aerogel occurred exclusively due to hydrophobic-hydrophobic interactions. Lipase-active biocatalysts prepared by adsorptive immobilization of recombinant *T. lanuginosus* lipase on macroporous carbon aerogel were studied in the periodic processes of bioconversion of triglycerides and fatty acids such as tributyrin hydrolysis, interesterification of vegetable oil with ethyl acetate, and esterification of saturated fatty acids (butyric C4:0, capric C10:0, and stearic C18:0)

The activity of the lipase-active biocatalysts and specific activity of adsorbed on MCA lipase were measured in hydrolysis of emulsified tributyrin under conditions when there were no diffusion limitations for mass transfer of the substrate from reaction media toward the adsorbed lipase [19]. Note that specific activity of adsorbed enzyme (in U/mg) is calculated by dividing the experimentally observed activity of the biocatalyst (in U/g) by the amount of adsorbed enzyme (in mg/g). The specific activities of the lipase adsorbed both on MCA granules and nonporous fine powders of carbon nanotubes were compared with each other; the values were found to be 160 and 12 U/mg, respectively. This significant difference was due to the relative rigidity of ball-shaped aerogel granules that prevented agglomeration of carbon nanotubes in "oil-in-water" triglyceride emulsion as presented in **Figure 5b**. Since water-immiscible hydrophobic triglyceride molecules were adsorbed efficiently on hydrophobic carbon nanotubes, with increasing tributyrin concentration, agglomerates of round shape containing this substrate were formed from CNTs (**Figure 5b**). As a result, the required hydrolysis water is displaced from the vicinity

of adsorbed lipase and its specific activity fell down as mentioned above.

the formation of the 1st protein dense layer at adsorption of 100–110 mg/g (**Figure 8a**). Then, the specific activity fell down as adsorption increased

(**Figure 8a**). As a result, maximal hydrolytic activity of the biocatalysts, 75,000 U/ g, was observed at the adsorption corresponding to the formation of the 1st protein layer. The activity of the biocatalyst with the double adsorptive layer was found to be 1.6-fold lower, 47,000 U/g. In order to explain maximum on the curve in **Figure 8a**, we proposed that the adsorbed lipase can be oriented differently on the

highly hydrophobic carbon surface of CNTs.

The maximal hydrolytic specific activity of the adsorbed *T. lanuginosus* lipase was measured to be 700 U/mg (vs. 14,000 U/mg for soluble lipase), i.e., activity significantly decreased upon adsorption of lipase on MCA. Another reason discussed below may be probably incorrect orientation of adsorbed lipase on a

The results of study on dependence of the specific activity of adsorbed lipase on the adsorption value are presented in **Figure 8a**. Initially, when adsorption of the lipase was small, the observed activities were extremely low. The specific activity increased dramatically and reached maximal values, >700 U/mg, in a region close to

(**Figure 7**).

*Molecular Biotechnology*

with *iso*-pentanol.

**16**

*Specific hydrolytic activity of lipase adsorbed on carbon aerogel depending on adsorption. Conditions of hydrolysis: 20 2°C, 0.02 M Na-phosphate buffer pH 7.0, 0.02 M tributyrin, 1.0 M glycerol, 0.6% gum arabic, content of biocatalyst in reaction medium 3.3 wt.%.*

surface of carbon nanotubes, namely correctly and incorrectly (**Figure 8b**). The incorrect orientation (No. 1 in **Figure 8b**) with blocking the active site by the surface was realized more likely in the incompact 1st adsorptive layer; in this case, the observed lipase activity had a minimum value. The correct orientation (No. 3 in **Figure 8b**) may be realized in the dense 1st adsorptive layer, so the formation of active enzyme-substrate complex occurred; and in this case, the observed lipase activity had a maximum value. A further decrease in specific activities after the formation of the 1st adsorptive layer was due to covering the active lipase by the molecules in subsequent 2nd adsorptive layer (scheme in **Figure 7**) and possible partially correct orientation (No. 2 in **Figure 8b**). The obtained data suggest that the main reasons for a loss of enzymatic activity upon adsorption of the lipase on hydrophobic carbon aerogel are as follows: (1) incorrect orientation of enzyme molecule on the carbon surface and blockage of the active site, as well as possible deformation of enzyme molecule; and (2) dehydration of the biocatalysts due to removal of essential water from the vicinity of adsorbed lipase via efficient adsorption of hydrophobic triglycerides.

It has been found that activity and stability of the prepared lipase-active heterogeneous biocatalysts depended strongly on the type of enzymatic reaction performed either in aqueous reaction media (hydrolysis) or in nonaqueous media (interesterification and esterification). The stability of the biocatalysts was quite low when triglycerides participated in reactions as lipase substrates. For example, the biocatalysts lost 90% of initial activity during six reaction cycles of tributyrin hydrolysis [19]. Another example, the stability of adsorbed on MCA r*Pichia*/lip was determined during interesterification of linseed oil with ethyl acetate in order to produce the valuable product, vitamin F—ethyl esters of ω-3 fatty acids. The conversion of triglycerides was 87 and 66% in the 1st and 3–5th reaction cycles, respectively, and then biocatalysts inactivated due to rapid dehydration.

As mentioned above, esterification of acid with alcohols was accompanied by the formation of water and the corresponding ester, and, as a result, accumulation of essential water molecules inside biocatalysts in the vicinity of the adsorbed lipase occurred. So, the stability of the prepared biocatalysts was much higher in esterification than in hydrolysis and interesterification. And the biocatalysts operated during esterification without a loss of activity for more than several hundred hours in the nonaqueous media of anhydrous organic solvents. As seen in **Figure 9**, the

of the biocatalyst, 75,000 U g<sup>1</sup>

*DOI: http://dx.doi.org/10.5772/intechopen.89411*

**Acknowledgements**

A17-117041710075-0.

**19**

, was measured at the adsorption corresponding to

the formation of the 1st dense adsorptive layer. The activity and stability of the prepared biocatalysts decreased during the periodic process of tributyrin hydrolysis because a displacement of essential water via efficient adsorption of triglyceride on CNTs occurred, which was followed by progressive dehydration of biocatalysts. In esterification of fatty acids with alcohol (nonaqueous reaction media), hydrophobic MCA did not prevent the accumulation of the produced essential water in the vicinity of adsorbed lipase. The lipase-active biocatalysts possessed high stability in the synthesis of esters of fatty acids with *iso*-pentanol and operated

*Heterogeneous Biocatalysts for the Final Stages of Deep Processing of Renewable Resources…*

for a few hundred hours under mild condition of the synthesis of *iso*-pentyl

As a general conclusion based on all results of adsorption/desorption and activity/stability, we can convincingly note that texture and chemical nature of supports such as mesoporous silica and macroporous carbon aerogel greatly affect the biocatalytic properties of the adsorbed recombinant *T. lanuginosus* lipase. This influence was due to not only hydrophilic-hydrophobic properties of the adsorbents but also specific molecular features of lipase and peculiarities of biocatalysis carried out in aqueous and nonaqueous reaction media. The main reason of inactivation of the prepared biocatalysts during bioconversion of hydrophobic triglycerides, such as hydrolysis and interesterification, was their dehydration. The accumulation of essential water inside the lipase-active biocatalysts during esterification prevented their inactivation. Analyzing our own and literature data, it was concluded that due to the prominent catalytic properties, such as enzymatic activity and operational stability, in combination with simplicity of the immobilization method, the biocatalysts prepared by adsorption of recombinant lipase onto mesoporous silica and macroporous carbon aerogel are promising for practical implementation for reactions of organic synthesis, in particular for production of fatty acids' esters.

The authors are grateful to Anatoly Beklemishev and Maria Pykhtina for design strain of r*Pichia*/lip producing recombinant *T. lanuginosus* lipase for this research. This work was conducted within the framework of the budget Project No. AAAA-

caprinate in binary (hexane and diethyl ether) solvent at 40°C.

#### **Figure 9.**

*Operational stability of the biocatalyst in periodic esterification of various fatty acids such as butyric (C4), capric (C10), and stearic (C18) with iso-pentanol as a function of the acid conversion depending on a number of reaction cycle. Conditions of esterification: 40°C, 0.10 M acid, 0.40 M alcohol, hexane:diethyl ether = 1:2, agitation 70 rpm, content of biocatalyst in reaction medium 3.3 wt.%. The amount of adsorbed lipase is 206.0 mg g<sup>1</sup> .*

esterification activity of dried biocatalysts sharply increased during the 1st reaction cycle; a pre-conditioning stage was observed as described above. The esterified activity depended on the structure of fatty acid, namely of a number of carbon atoms in molecule. Similar to LipoSil biocatalysts described above, the rate of esterification of butyric C4 acid was the lowest compared to capric (C10:0) and stearic (C18:0) acids (**Figure 9**). In the process of the synthesis of *iso*-pentyl caprinate in binary (hexane and diethyl ether) solvent at 40°C, the prepared biocatalysts demonstrated high operational stability and operated without losing activity for a few hundred hours [19].

#### **3.4 Conclusion for the part 3.2**

Macroporous carbon aerogel (MCA) obtained by *in situ* catalytic synthesis of the carbon nanotubes was studied as the efficient promising support for adsorptive immobilization of enzymes, in particular recombinant *T. lanuginosus* lipase, due to the rigid 3D-framework of granules and high adsorption ability of MCA. Heterogeneous lipase-active biocatalysts were prepared by adsorption of the enzyme on carbon nanotube (CNT)-forming aerogel, and the processes of bioconversion of substrates into valuable products were organized in periodic regimes under mild ambient conditions.

Recombinant *T. lanuginosus* lipase was adsorbed on macroporous carbon aerogel very firmly exclusively due to hydrophobic interactions between enzyme and carbon nanotubes. Two dense adsorptive layers were formed by lipase molecules; the amount of the lipase in each layer was nearly 100 mg/g.

Activity and stability of the prepared biocatalysts strongly depended on the type of reaction media, namely aqueous or nonaqueous. In the hydrolysis of triglycerides (aqueous reaction medium), the specific activity of the *T. lanuginosus* lipase significantly (in 20 times) decreased upon adsorption on MCA, because orientation of adsorbed lipase on the carbon surface was probably incorrect. The maximal activity

### *Heterogeneous Biocatalysts for the Final Stages of Deep Processing of Renewable Resources… DOI: http://dx.doi.org/10.5772/intechopen.89411*

of the biocatalyst, 75,000 U g<sup>1</sup> , was measured at the adsorption corresponding to the formation of the 1st dense adsorptive layer. The activity and stability of the prepared biocatalysts decreased during the periodic process of tributyrin hydrolysis because a displacement of essential water via efficient adsorption of triglyceride on CNTs occurred, which was followed by progressive dehydration of biocatalysts.

In esterification of fatty acids with alcohol (nonaqueous reaction media), hydrophobic MCA did not prevent the accumulation of the produced essential water in the vicinity of adsorbed lipase. The lipase-active biocatalysts possessed high stability in the synthesis of esters of fatty acids with *iso*-pentanol and operated for a few hundred hours under mild condition of the synthesis of *iso*-pentyl caprinate in binary (hexane and diethyl ether) solvent at 40°C.

As a general conclusion based on all results of adsorption/desorption and activity/stability, we can convincingly note that texture and chemical nature of supports such as mesoporous silica and macroporous carbon aerogel greatly affect the biocatalytic properties of the adsorbed recombinant *T. lanuginosus* lipase. This influence was due to not only hydrophilic-hydrophobic properties of the adsorbents but also specific molecular features of lipase and peculiarities of biocatalysis carried out in aqueous and nonaqueous reaction media. The main reason of inactivation of the prepared biocatalysts during bioconversion of hydrophobic triglycerides, such as hydrolysis and interesterification, was their dehydration. The accumulation of essential water inside the lipase-active biocatalysts during esterification prevented their inactivation. Analyzing our own and literature data, it was concluded that due to the prominent catalytic properties, such as enzymatic activity and operational stability, in combination with simplicity of the immobilization method, the biocatalysts prepared by adsorption of recombinant lipase onto mesoporous silica and macroporous carbon aerogel are promising for practical implementation for reactions of organic synthesis, in particular for production of fatty acids' esters.

## **Acknowledgements**

esterification activity of dried biocatalysts sharply increased during the 1st reaction cycle; a pre-conditioning stage was observed as described above. The esterified activity depended on the structure of fatty acid, namely of a number of carbon atoms in molecule. Similar to LipoSil biocatalysts described above, the rate of esterification of butyric C4 acid was the lowest compared to capric (C10:0) and stearic (C18:0) acids (**Figure 9**). In the process of the synthesis of *iso*-pentyl caprinate in binary (hexane and diethyl ether) solvent at 40°C, the prepared biocatalysts demonstrated high operational stability and operated without losing

*Operational stability of the biocatalyst in periodic esterification of various fatty acids such as butyric (C4), capric (C10), and stearic (C18) with iso-pentanol as a function of the acid conversion depending on a number of reaction cycle. Conditions of esterification: 40°C, 0.10 M acid, 0.40 M alcohol, hexane:diethyl ether = 1:2, agitation 70 rpm, content of biocatalyst in reaction medium 3.3 wt.%. The amount of adsorbed lipase is*

Macroporous carbon aerogel (MCA) obtained by *in situ* catalytic synthesis of the carbon nanotubes was studied as the efficient promising support for adsorptive immobilization of enzymes, in particular recombinant *T. lanuginosus* lipase, due to the rigid 3D-framework of granules and high adsorption ability of MCA. Heterogeneous lipase-active biocatalysts were prepared by adsorption of the enzyme on carbon nanotube (CNT)-forming aerogel, and the processes of bioconversion of substrates into valuable products were organized in periodic regimes under mild

Recombinant *T. lanuginosus* lipase was adsorbed on macroporous carbon aerogel very firmly exclusively due to hydrophobic interactions between enzyme and carbon nanotubes. Two dense adsorptive layers were formed by lipase molecules; the

Activity and stability of the prepared biocatalysts strongly depended on the type of reaction media, namely aqueous or nonaqueous. In the hydrolysis of triglycerides (aqueous reaction medium), the specific activity of the *T. lanuginosus* lipase significantly (in 20 times) decreased upon adsorption on MCA, because orientation of adsorbed lipase on the carbon surface was probably incorrect. The maximal activity

amount of the lipase in each layer was nearly 100 mg/g.

activity for a few hundred hours [19].

**3.4 Conclusion for the part 3.2**

ambient conditions.

**18**

**Figure 9.**

*206.0 mg g<sup>1</sup>*

*.*

*Molecular Biotechnology*

The authors are grateful to Anatoly Beklemishev and Maria Pykhtina for design strain of r*Pichia*/lip producing recombinant *T. lanuginosus* lipase for this research. This work was conducted within the framework of the budget Project No. AAAA-A17-117041710075-0.

*Molecular Biotechnology*
