Lipases catalytic activity

Lipases from different sources have shown different substrate specificity and catalytic activity. Lipases with narrow specificity are not suitable for biodiesel production. The performance of regiospecific lipases can possibly improve when used with nonspecific lipases in combination. Also, some lipases show more hydrolytic activity while others... 

Lipases have proven to be effective in prespotters and other liquid detergent formulations when used in undiluted form for pretreatment of tough fatty stains. The low water content on the fabric in this situation is believed to be responsible for the high catalytic activity (50). 

Lipases have also been used as initiators for the polymerization of lactones such as /3-bu tyro lac tone, <5-valerolactone, e-caprolactone, and macrolides.341,352-357 In this case, the key step is the reaction of lactone with die serine residue at the catalytically active site to form an acyl-enzyme hydroxy-terminated activated intermediate. This intermediate then reacts with the terminal hydroxyl group of a n-mer chain to produce an (n + i)-mer.325,355,358,359 Enzymatic lactone polymerization follows a conventional Michaelis-Menten enzymatic kinetics353 and presents a controlled character, without termination and chain transfer,355 although more or less controlled factors, such as water content of the enzyme, may affect polymerization rate and the nature of endgroups.360... 

Aqueous solutions are not suitable solvents for esterifications and transesterifications, and these reactions are carried out in organic solvents of low polarity [9-12]. However, enzymes are surrounded by a hydration shell or bound water that is required for the retention of structure and catalytic activity [13]. Polar hydrophilic solvents such as DMF, DMSO, acetone, and alcohols (log P<0, where P is the partition coefficient between octanol and water) are incompatible and lead to rapid denaturation. Common solvents for esterifications and transesterifications include alkanes (hexane/log P=3.5), aromatics (toluene/2.5, benzene/2), haloalkanes (CHCI3/2, CH2CI2/I.4), and ethers (diisopropyl ether/1.9, terf-butylmethyl ether/ 0.94, diethyl ether/0.85). Exceptionally stable enzymes such as Candida antarctica lipase B (CAL-B) have been used in more polar solvents (tetrahydrofuran/0.49, acetonitrile/—0.33). Room-temperature ionic liquids [14—17] and supercritical fluids [18] are also good media for a wide range of biotransformations. 

Enzymatic synthesis of aliphatic polyesters was also achieved by the ringopening polymerization of cyclic diesters. Lactide was not enzymatically polymerized under mild reaction conditions however, poly(lacfic acid) with the molecular weight higher than 1 x 10" was formed using lipase BC as catalyst at higher temperatures (80-130°C). Protease (proteinase K) also induced the polymerization however, the catalytic activity was relatively low. 

Several enzymes like lipases, esterases, and dehydrogenases have been active in hydrophobic environments. Thermodynamic water activity is a good predictor of the optimal hydration conditions for catalytic activity [51]. Enzyme preparation can be equilibrated at a specific water activity before the reaction [52]. When water concentration is very low, enzyme is suspended in the solid state in the water-immiscible organic solvent [46]. Enzymes are easily recovered after the reaction by the method of filtration. 

Vacuum was applied to shift the equilibrium forward by removal of the activated alcohol formed [30, 31, 37, 38]. In the enzymatic polycondensation of bis(2,2,2-trifluoroethyl) sebacate and aliphatic diols, the polymer with Mw of more than 1 x 104 was obtained using lipases CC, MM, PPL, and Pseudomonas cepacia lipase (lipase PC) as catalyst and lipase MM showed the highest catalytic activity [37]. Solvent screening indicated that diphenyl ether and veratrole were suitable for the production of the high molecular weight polyesters under vacuum. In the PPL-catalyzed reaction of bis(2,2,2-trifluoroethyl) glutarate with 1,4-butanediol in veratrole or 1,3-dimethoxybenzene, periodical vacuum method improved the molecular weight (Mw 4 x 104) [38]. 

Lipase-catalyzed polymerization of divinyl adipate or divinyl sebacate with a, co-glycols with different chain length has been reported [40]. Lipases CA, MM, PC, and PF showed high catalytic activity toward the polymerization. A combination of divinyl adipate, 1,4-butanediol, and lipase PC afforded the polymer with number-average molecular weight (Mn) of 2.1 x 104. The yield of the polymer from divinyl sebacate was higher than that from divinyl adipate, whereas the opposite tendency was observed in the polymer molecular weight. 

Lactide was polymerized by lipase PC in bulk at high temperature (80-130°C) to produce poly(lactic acid) with Mw up to 2.7 x 105 [64, 65]. The molecular weight of the polymer from the D,L-isomer was higher than that from the d,d- and L,L-ones. Protease (proteinase K) also induced the polymerization of lactide, however, the catalytic activity was relatively low. 

Lipase catalyzed the ring-opening polymerization of medium-size lactones, d-valerolactone (<5-VL, six-membered) and -caprolactone (c-CL, seven-mem-bered). Lipases CC, PF and PPL showed high catalytic activity for the polymerization of <5-VL [74,75]. The molecular weight of the polymer obtained in bulk at 60 °C was relatively low (less than 2000). 

Nine-membered lactone, 8-octanolide (8-OL), was also enzymatically polymerized [ 84]. Lipases CA and PC showed the high catalytic activity for the polymerization. 

Four macrolides, 11-undecanolide (12-membered,UDL) [85,86], 12-dodeca-nolide (13-membered,DDL) [86,87], 15-pentadecanolide (16-membered, PDL) [85,86,88,89], and 16-hexadecanolide (17-membered, HDL) [90], were subjected to the lipase-catalyzed polymerization. For the polymerization of DDL, lipases CC, PC, PF, and PPL showed the high catalytic activity and the activity order in the bulk polymerization was as follows lipase PC > lipase PF > lipase CC> PPL. These enzymes were also active for the polymerization of other macrolides. NMR analysis showed that the terminal structure of the polymer was of carboxylic acid at one end and of alcohol at the other terminal. 

Enzyme activity for the polymerization of lactones was improved by the immobilization on Celite [93]. Immobilized lipase PF adsorbed on a Celite showed much higher catalytic activity than that before the immobilization. The catalytic activity was further enhanced by the addition of a sugar or poly(ethylene glycol) in the immobilization. Surfactant-coated lipase efficiently polymerized the ring-opening polymerization of lactones in organic solvents [94]. 

The concept of zeolite action was tested in a particular reaction where the enzyme is exposed from the beginning to an acidic environment the esterification of geraniol with acetic acid catalyzed by Candida antarctica lipase B immobilized on zeolite NaA [219]. Lipases have been used for the hydrolysis of triglycerides and due to their ambivalent hydrophobic/hydrophilic properties they are effective biocatalysts for the hydrolysis of hydrophobic substrates [220]. When water-soluble lipases are used in organic media they have to be immobilized on solid supports in order to exhibit significant catalytic activity. 

Table 2. Catalytic activity of the immobilized lipase in the transesterification reaction of triolein with methanol (18 hours of reaction, 40°C, trioleimmethanol molar ratio 1 3).

The packaging of triacylglycerol into chylomicrons or VLDL provides an effective mass-transport system for fat. On a normal Western diet, approximately 400 g of triacylglycerol is transported through the blood each day. Since these two particles cannot cross the capillaries, their triacylglycerol is hydrolysed by lipoprotein lipase on the luminal surface of the capillaries (see above). Most of the fatty acids released by the lipase are taken up by the cells in which the lipase is catalytically active. Thus the fate of the fatty acid in the triacylglycerol in the blood depends upon which tissue possesses a catalytically active lipoprotein lipase. Three conditions are described (Figure 7.23) ... 

DKR of secondary alcohol is achieved by coupling enzyme-catalyzed resolution with metal-catalyzed racemization. For efficient DKR, these catalyhc reactions must be compatible with each other. In the case of DKR of secondary alcohol with the lipase-ruthenium combinahon, the use of a proper acyl donor (required for enzymatic reaction) is parhcularly crucial because metal catalyst can react with the acyl donor or its deacylated form. Popular vinyl acetate is incompatible with all the ruthenium complexes, while isopropenyl acetate can be used with most monomeric ruthenium complexes. p-Chlorophenyl acetate (PCPA) is the best acyl donor for use with dimeric ruthenium complex 1. On the other hand, reaction temperature is another crucial factor. Many enzymes lose their activities at elevated temperatures. Thus, the racemizahon catalyst should show good catalytic efficiency at room temperature to be combined with these enzymes. One representative example is subtilisin. This enzyme rapidly loses catalytic activities at elevated temperatures and gradually even at ambient temperature. It therefore is compatible with the racemization catalysts 6-9, showing good activities at ambient temperature. In case the racemization catalyst requires an elevated temperature, CALB is the best counterpart. 

It is worth noting that the enzyme can be withdrawn and recycled by using supercritical CO2. The success of the polymerizations carried out in organic solvents stems directly from the sustained activity of several lipases in organic solvents. In this respect, it must be noted that water has a manifold influence on the course of the polymerization. On the one hand, water can initiate the polymerization. On the other hand, a minimum amount of water has to be bound to the surface of the enzyme to maintain its conformational flexibility, which is essential for its catalytic activity [94]. Lipase-mediated polymerization cannot therefore be achieved in strictly anhydrous conditions. 

T. Maruyama, T. Kotani, H. Yamamura, N. Kamiya, M. Goto, Poly(ethylene glycol)-lipase complexes catalytically active in fluorous solvents, Org. Biomol. Chem. 2 (2004) 524-527. 

We will first focus on the implications of the catalytic activation mechanism of lipases, namely monomer activation, on the attainable chains lengths and polydis-persity when cyclic esters are converted into polyesters. Moreover, the control of... 

The influence of surfactant on the catalytic activity of lipases in water is well known. The addition of surfactants can enhance the activity and enantioselectivity of these enzymes in aqueous solutions [94] due to the interfacial activation and due to the emulsification of hydrophobic substrates. 

Itmnobilization of lipases in silica gels by gel entrapment has recently received interest (Sata et al, 1994 Reetz, 1997 Reetz et al, 1996). This so-called sol-gel process involves hydrolysis of SilOR) in the presence of enzyme. The formed Si-monomers crosslink in the presence of an acid or base catalyst into an amorphous Si02-network entrapping the enzyme. The use of alkylsilanes RSilOCHjlhas been reported to give lipase preparations with unusually high catalytic activities (Reetz, 1997). 

Lipases exhibit high catalytic activity in water, an even higher activity in a two-phase system, such as water/water-immiscible organic solvent, and in water-immiscible organic solvents of low water content86-88,90. This allows for the attainment of favorable equilibria in asymmetric hydrolysis and esterification reactions catalyzed by lipases. They are used to their greatest... 

Relatively few detailed studies of enzyme kinetics in organic media have been carried out. Preferably, full kinetics should be studied, allowing the determination of Km and kcat values, but it is much more common to see just reports on the catalytic activity at fixed substrate concentrations as a function of water activity. That such studies can be misleading was shown in an investigation of lipase-catalyzed esterification [26]. When the reaction rate in the esterification reaction was plotted versus the water activity at three different substrate concentrations, maxima were obtained at three different water activities (Figure 1.4). Such maxima should not be used to claim that the optimal water activity of the enzyme was found. Detailed kinetic studies showed that both the kcat and the Km values (for the alcohol substrate) varied with the water activity. The Km value of the alcohol increased with increasing water... 

Flomenbom. O, Velonia, K.. Loos. D.. Masuo. S.. Cotlet, M, Engelborghs, Y., Hofkens. J.. Rowan, AE., Nolte, R.J.M., de Schyver. F.C, and Klafter, J. (2005) Stretched exponential decay and correlations in the catalytic activity of fluctuating single lipase molecules. Proc. Natl. Acad. Sci. U.S.A., 102, 2368-2372. 

Potentiometry is another useful method for determining enzyme activity in cases where the reaction liberates or consumes protons. This is the so-called pH-stat method. pH is kept constant by countertitration, and the amount of acid or base required is measured. An example of the use of this method is the determination of lipase activity. The enzyme hydrolyzes triglycerides and the fatty acids formed are neutralized with NaOH. The rate of consumption of NaOH is a measure of the catalytic activity. 

Porcine pancreatic lipase catalyzes the transesterification reaction between tribu-tyrin and various primary and secondary alcohols in a 99% organic medium (Zaks, 1984). Upon further dehydration, the enzyme becomes extremely thermostable. Not only can the dry lipase withstand heating at 100 °C for many hours, but it exhibits a high catalytic activity at that temperature. Reduction in water content also alters the substrate specificity of the lipase in contrast to its wet counterpart, the dry enzyme does not react with bulky tertiary alcohols. 

R. H. Valivety, P. J. Halling, A. D. Peilow, and A. R. Macrae, Lipases from different sources vary widely in dependence of catalytic activity on water activity, Biochim. Biophys. Acta 1992b, 1122, 143-146. 

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