Kinetics and mechanisms of reactions catalysed by immobilized lipases

Kinetics and mechanisms of reactions catalysed by immobilized lipases F. Xavier Malcata,* Hector R. Reyes,* Hugo S. Garcia,t Charles G. Hill, Jr.*'t and Clyde H. Amundsont,$ Departments o f * Chemical Engineering, ?Food Science, and SAgricultural Engineering, University o f Wisconsin, Madison, WI, USA

This review focuses on the kinetics and mechanisms of reactions catalysed by immobilized lipases. The effects of pH, temperature, and various substances on the catalytic properties of immobilized lipases and on the processes by which they are deactivated are reviewed and discussed.

Keywords:Immobilizedenzymes;rate equations;factors affectingkinetics

Introduction Glycerol esters of fatty acids, which constitute ca. 99% of the lipids of natural origin, have traditionally been called fats and oils.~ (The distinction between a fat and an oil is made on the basis of whether the material in question is a solid or a liquid, respectively, at room temperature.) A large number of chemicals can be produced from vegetable oils and animal fats. Since these feedstocks are renewable and can be produced in virtually every country of the world, ~ there has recently been a renewal of interest in using enzymes for chemical conversion of these materials. Lipases play a major role in this respect because they are commercially available in large quantities, they are very selective towards their substrates, and they are very efficient catalysts which are effective under mild operating conditions. 3 Among the most promising new processes for chemical modification of lipids are the hydrolysis, ester synthesis, and interesterification reactions of these materials in the presence of lipases. ~ Hydrolysis reactions involve an attack on the ester bond of glycerides in the presence of water molecules to produce both an alcohol functionality and a carboxylic acid. Esterification reactions between polyhydric alcohols and free fatty acids are, in essence, the reverse of the hydrolysis reaction of the corresponding glyceride. The relative rates of the forward and the reverse reactions are usually con-

trolled by the water content of the reaction mixture. The term interesterification refers to the exchange of acyl radicals between an ester and an acid (acidolysis), an ester and an alcohol (alcoholysis), or an ester and another ester (transesterification). The costs of producing the lipases necessary to catalyse the aforementioned reactions are often prohibitive .4,5Hence processes that do not require the physical presence of lipase in the final product and that use feedstocks that are fluids (or that can be treated as such) are subject to dramatic improvements in process economics if the lipase is employed in immobilized form) Studies of the technical feasibility of processes employing immobilized lipases have recently been reviewed by Malcata et al. 6 Insofar as is possible, the design of commercial-scale enzymatic processes should be based on a quantitative understanding of the underlying physical and chemical phenomena. Hence, one critical step in the design of immobilized lipase reactors is a determination of the rate expressions that describe the performance of the enzyme and relate this performance to the major operating conditions under the operator's control (i.e. the concentrations of substrates and/or products, reactor space time, pH, and temperature). After providing general background information relative to the catalytic activity of lipases, this paper critically reviews the kinetic properties of immobilized lipases.

General characteristics of lipases pertinent to their catalytic activity Address reprint requests to Prof. C. G. Hill, Chairman,Department of Chemical Engineering, University of Wisconsin-Madison, 1415 Johnson Dr., Madison, WI 53706 Received 7 May 1991;revised 27 December 1991 426

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The activation of lipases by interfaces, the effects of various structural properties on the catalytic action of lipases, the constraints imposed by the physicochemical properties oflipases, and the specificity characteris© 1992 Butterworth-Heinemann

Immobilized lipase kinetics: F. X. Malcata et al. tics of lipases are described in the subsections that follow.

Activation of lipases by interfaces Acylglycerol hydrolases (EC 3.1.1.3), or lipases, are enzymes that catalyse the reversible hydrolysis of tri-, di-, and monoglycerides.7 However, this definition may be deemed too narrow because lipases are also able to bring about the hydrolysis of a variety of compounds containing carboxylic ester moieties which are not acylglycerols.8 Although lipases can act on soluble monomeric substrates, practical utilization of lipase-catalysed reactions is restricted almost exclusively to situations where the overall substrate concentration is higher than its solubility in the natural solvent, water. 9'1° This characteristic restriction provides a convenient criterion for differentiating lipases from conventional esterases, which ordinarily act only on soluble monomeric substrates. High concentrations of glycerides and other organic substrates lead to phase separation and concomitantly to the formation of an oil/water interface. The observed rates of lipase-catalysed reactions are strongly influenced by the available interfacial area. 1i Theoretical interpretations of the activation of lipases by interfaces have been attempted by a number of authors. These interpretations can be divided into two major groups: (i) those which assume that the substrates are activated by the presence of an oil/water interface, and (ii) those which assume that the lipase undergoes a change to an activated form upon contact with an oil/water interface.12 Explanations for the former type of activation involve higher concentrations of substrates in the vicinity of the interface than in the bulk of the oil, ~3more suitable conformations or orientations of the lipid molecules for chemical reaction, 14-17 and poor hydration of lipid molecules, thereby avoiding ester-bond shielding in the neighborhood of oil/water interfaces) 8:9 Explanations for the latter type of activation involve the existence of separate adsorption and catalytic sites for the lipase such that the lipase becomes catalytically active only after binding to the interface, 2°-22 or a conformational change of the lipase upon approaching the oil/water interface that might be due to the high degree of order in such regions 22-25 or to reorientation of the short one-turn or-helix that covers the active s i t e . 26 As Malcata et al. 6 have emphasized, when lipases are immobilized on porous solid supports, the highest retention of lipolytic activity is usually observed when the carrier possesses hydrophobic characteristics. When contacted with oil or melted fat, hydrophobic supports tend to be selectively permeated by these substances, thus stripping most water molecules away from the immediate vicinity of the lipase. Hence, the true oil/water interface is not likely to be in direct contact with the immobilized lipase. This fact seems to contradict the generally accepted dogma that an oil/ water interface is needed in order for lipases to exhibit full catalytic activity. Consequently, it is preferable

to formulate the condition for existence of lipolytic activity in more general terms as the requirement for

a continuous ordered hydrophobic microenoironment. The surfaces of contact of two phases provide a continuum irrespective of whether these phases are two liquid phases (e.g. oil and water) or a liquid and a solid phase (i.e. oil and the support for lipase). At oil/water interfaces, the lipid molecules orient themselves in a manner that gives rise to an ordered packing in which virtually all the polar heads are exposed to the water phase and virtually all the hydrocarbon tails are exposed to the organic phase. 27 (The lipid molecules in true solution orient themselves randomly because they are completely surrounded by water molecules.) This ordered continuum can also be achieved on the hydrophobic surface of a solid carrier. Only the immediate vicinity of the lipase should possess a hydrophobic character z8 because the overall conformation of the enzyme and the local conformation of the active site depend primarily on the intramolecular forces and the short-range interactions of the protein with the solvent. The water molecules used as substrate species in the hydrolysis reaction are more likely to have reached the active site by diffusion from the bulk of the oil phase than by direct contact of the water phase with the active site. 29 If diffusional limitations on the reaction rate are to be avoided, a high degree of saturation of the oil phase must be maintained. 3°'31

Proteinaceous structure o f lipases Lipases can be obtained from mammals, 32-35 plants, 36 and microbes .4,37The number of amino acid residues of the lipases studied varies from 270 to 641. The primary structures of several lipases obtained from mammalian, 26,38-54 bacterial, 55-58 and funga141'59 sources have been determined from either amino acid or nucleic acid sequences. All lipases sequenced to date share sequence homologies including a significant region, His-X-Y-Gly-Z-Ser-W-GIy or Y-Gly-His-Ser-W-Gly (W, X, Y, and Z denote unspecified amino acid residues) that is conserved in all such enzymes. 6° The presence of a histidine residue at the active site is consistent with results obtained for the activity of lipases upon removal of His,61'62 photooxidation,63 combination with ethoxyformate :2 and treatment with diethylpyrocarbonate. 64 The pK values for amino acid residues relevant for catalysis in the typical range corresponding to His have been reported by a number of authors. 13.65,~sThe argument for the presence of a serine residue at the active site is based on results for the activity of lipases upon derivatization with diethyl p-nitrophenylphosphate,67 use of specific inhibition reactants, 68reaction with organophosphates 64'69'7°or diisopropylfluorophosphate, 6° transformation with alkylisocyanates,71 labeling with phenylmethylsulfonylfluoride,72and contacting with butylboronic acid prior to assay by differential x-ray crystallography.26 The three-dimensional structure of the lipase produced by Geotrichum candidum has been determined by x-ray analysis of its crystals. This enzyme is ellipsoi-

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Review dal in shape with dimensions 50 x 50 x 70 A, and it possesses a large cleft in the vicinity of the molecular center (where the active site is believed to be located).73 The results obtained from x-ray crystallography of pancreatic lipase 26and lipase from Mucor miehei 72indicate that serine is the nucleophilic residue essential for catalysis. This residue is located in the larger N-terminal domain at the C-terminal edge of a doubly wound parallel/3-sheet and is part of an Asp-His-Ser triad, which is chemically analogous, but structurally different from that in the serine proteases. 26 This putative hydrolytic site is buried under a "lid" composed of a short helical fragment of a long surface loop stabilized by extensive hydrophobic and electrostatic interactions. 26,72 Hence it is inaccessible to solvent. The catalytic action of lipase can thus be viewed as a two-stage process: the aforementioned "lid" is removed or displaced (possibly through interfacial activation), and the substrate ester bond is subsequently hydrolysed by a mechanism very similar to that involved in the action of serine proteases. 72 The "lid" may also serve as a device to inhibit the proteolytic activity of the triad, thereby protecting the enzyme itself. The interaction of the lipase with the oil/water interface could be controlled by the serine residue, whereas the carboxyl residue would be of importance in adjusting or stabilizing the best active conformation of the lipase. 33 As for the quaternary structure, experimental evidence exists indicating that some lipases are composed of more than a single subunit: examples include two, 6~ four, TMand six 56 subunits. Some lipases are also associated with glycosidic 21,26and lipid 75residues. The molecular weights of the known lipases in their active, native forms range from 27 kilodaltons for a lipase from Penicillium cyclopium 76and 29 kilodaltons for a lipase from Pseudomonas aeruginosa 77 to over 500 kilodaltons for human lingual lipase. 33

Physicochemical properties o f lipases Lipases are generally soluble in water. Most animal lipases exhibit pH optima on the alkaline side (say, 8 to 9). 7 However, depending upon the substrate used, the presence of salts, and the kind of emulsifiers present, the optimum may be shifted to the acidic range .78,79 Acid lipases have been found in the lysosomes of a variety of mammalian tissues. 8°,8~ Most microbial lipases display maximum activity at pH values ranging from 5.6 to 8.57 and maximum stability in the neutral pH range. 37 With respect to temperature, most lipases are optimally active between 30 and 40°C. 82 The thermostability of lipases varies considerably according to their origin: animal and plant lipases are usually less thermostable than microbial extracellular lipases. 83 It has been reported that an extremely thermophilic strain of Pseudomonas releases a lipase which is heat-stable at 100°C. 84'85 The heat stability of lipases depends on whether or not substrate is present, 86-88 probably because substrate removes excess water from the immediate vicinity of the enzyme and thus restricts its overall conformational mobility.

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Lipases are strongly adsorbed at air/water interfaces. 89 A lipase from Candida cylindracea was easily

inactivated at an air/water interface due to its high interfacial tensiong°; this effect is enhanced by shear forces and increases with temperature. A lipase from Candida cylindracea has been reported to lose activity as a function of shearing time and shear rate. 9°

Specificity o f lipases The specificities of lipases have classically been divided into five major types8: (i) lipid class, (ii) positional, (iii) fatty acid, (iv) stereochemical, and (v) combinations thereof. Lipid class selectivity has been observed in animal plasma which apparently contains separate lipoprotein lipases for the hydrolysis of triglycerides, diglycerides, and monoglycerides. 8 A lipase produced by a strain of Penicillium cyclopium 91 has been shown to display its highest activity on monoglycerides, and much lower activities toward di- and triglycerides. This type of selectivity is dependent on temperature for a lipase from Pseudomonas fluorescens.92"93 Lipases obtained from natural sources can be positionally nonspecific or display one of two kinds of positional specificity: sn-l,3 specific or sn-2 specific. Nonspecific lipases hydrolyse all three ester bonds of triglycerides equally well. Nonspecificity has been observed for lipases from Chromobacterium o i s c o s u m , 94 Pseudomonas fluorescens, 95 Candida cylindracea, 96 Geotrichum candidum, and Penicillium eyelopium, 97'98 and also for hepatic lipase. 99 Specificity of the sn-l,3 type is associated with the preferential release of fatty acid residues from the terminal positions of the glycerol backbone rather than from the central carbon atom, whereas sn-2 specificity refers to preferential release from the central carbon atom. The sn-l,3 type of specificity has been observed for pancreatic 1~ and adipose tissue 1°° lipases, Rhizopus arrhizus lipase,l°l:°2 Aspergillus niger lipase, 97.98.1°2Rhizopus delemar lipase, 97,98 and Mucor miehei lipase. J0z The sn-2 specificity is extremely rare, and it has been ascribed to a lipase from Geotrichum candidum which has a particular ability to hydrolyse oleic and linoleic acids from the sn-2 location. 8 A more general classification states that the positional specificity of lipases is not divided clearly into the above categories; instead it changes continuously from highly specific sn-l,3 activity to a very weakly specific or completely nonspecific activity. 83 Lipases often exhibit a particular ability to release fatty acids whose chain lengths or degrees of unsaturation fall within well-defined ranges. This situation has been explored in the lipase-catalysed production of cheese-type flavors. ~03 Early reports by Brockerhoff ~4 and Desnuelle and Savary 1°4 indicated a preference of pancreatic lipase for C4 over other saturated fatty acid moieties. The lipase from Mucor rniehei exhibits similar activities for release of C4 and C6 acids at either pH 5.3 or 8.0; however, longer chain fatty acids are released more slowly at the acidic pH.~°5 Experimental

Immobilized lipase kinetics: F. X. Malcata et al. data for the fatty acid specificities of lipases from Chromobacterium viscosurn 94 and Geotrichum candidum 1o6 indicate that the rate of attack by these lipases follows a bell-shaped distribution in the number of carbon atoms in the hydrocarbon backbones of unsaturated fatty acid residues. The distribution is centered at C8-C10. Experimental data for the alcohol chain length specificities of lipases from Candida rugosa, Aspergillus niger, and Rhizopus arrhizus 1°7 in ester synthesis reactions also follow a bell-shaped distribution in the number of carbon atoms centered at C4, Ca, and C2, respectively. Bimodal distributions consisting of two superimposed bell-shaped distributions centered at C 3 - C 4 and C8-C10 for lipases from Penicillium cyclopium, Aspergillus niger, and Rhizopus delemar, 106or at C2 and C5 for a lipase from Mucor miehei, 107were also observed. For the same chain length of the fatty acid residue, the rate of attack by lipase seems to increase with the number of double bonds in the hydrocarbon backbone, 1o6although more extensive studies have demonstrated that similar bell-shaped variations of the rate of attack versus the number of double bonds in the hydrocarbon backbone exist. 94One explanation for this type of specificity involves the concept of induced fit. Although a great many substrates can bind at the active site, only a few can release the proper amount of binding energy required for the change in the conformation of the lipase to a form which is a much more efficient catalyst.l°8 Substrates which are too small or possess too few double bonds are not able to release enough binding energy. In such cases the change in conformation of the native lipase to the desired catalytically active conformation does not occur or is, at best, incomplete. Hence, the reaction will proceed slowly. Substrates which are too long or possess too many double bonds are able to release an amount of binding energy which would in principle be sufficient to effect the desired conformational change. However, some of this energy becomes unavailable for this purpose because it is required to change the conformation of the substrate so as to make it fit into the active site. Hence only a small fraction of the energy released by the binding process will actually be available to drive the conformational change of the enzyme. Consequently, optimal activity will not be achieved. The existence of a bimodal distribution of activity may be explained by the existence of more than one active site in the overall iipase molecule26: one of these active sites is a typical lipase active site (which acts preferentially on insoluble substrates), and the other is a typical esterase active site (which acts preferentially on soluble substrates). The distribution of activities of lipases relative to various triglycerides changes with temperature; as temperature is increased, the rates of release of long-chain fatty acids increase faster than those of the corresponding short-chain acids) °9 By taking advantage of the different fatty acid (and positional) specificities pertaining to different lipases, it has been possible to effect synergistic hydrolysis of soybean oil using mixtures of

lipases from Penicillium sp. and Rhizopus niveus, or from Penicillium sp. and Rhizopus delemar, n0 and synergistic production of monoglycerides by partial hydrolysis of palm oil and tallow using combinations of lipases from Penicillium camembertii and Humicola lanuginosa.111 Immobilization of Mucor rniehei lipase on a Duolite ion-exchange resin hydrolysed more C14:0 and C16 : 0 than the free enzyme. 112In the case of lipases from Mucor miehei, the interesterification activity increases markedly from acetic to octanoic acid, but is almost independent of chain length between octanoic and stearic acids, n3a 14A special kind of fatty acid specificity has been reported for the hydrolytic action of a lipase from Geotrichum candidum on fatty acids with cis-9 unsaturation. 115,116 Although early studies did not report evidence of any kind of stereospecificity for the catalytic action of lipases on fats and oils, a rather large body of literature dealing with the preparation of chiral esters and alcohols via lipase-mediated kinetic resolution of racemic (nontriglyceride) substrates is currently available. 102A number of researchers have observed stereoselectivity for the catalytic action of lipase on such substrates as straight-chain secondary alcohols, 117,118 aceton i d e 1°2:19'12° and butyric acid optically active esters, 12),122oxazolidones, ~23glycidyl ethers, 120cyclohexanols,)24'125 2-benzylglycerol ether, 126sugar alcohols, 127 and several esters of ibuprofen. 128Stereoselectivity has also been observed with serum lipoprotein lipase for hydrolysis of the enantiomeric esters at the sn-1 position, 129 and with h u m a n 129 and rat 13° lingual lipases for the hydrolysis of enantiomeric esters at the sn-3 position. Finally, combinations of fatty acid and stereoselectivities have been found with rat lingual TMand a human lipoprotein lipase preparation. 129

Effects o f inhibitors and promoters on the activity o f lipases It is generally recognized that free fatty acids 13z and alcohols )32,133tend to inhibit lipase-catalysed hydrolysis reactions. The precise mechanism by which this inhibition occurs is still unclear, but fatty acid molecules are thought to accumulate at the lipid/water interface, thereby blocking access of the enzyme to unreacted triglyceride moleculesm; low-molecular-weight alcohols are believed to disrupt the three-dimensional architecture of the lipase. Many light metal cations are known to influence the action of lipases on their substrates. Calcium ions usually play important roles in the mechanisms of both the hydrolysis and ester synthesis reactions. The presence of these ions usually leads to increases in reaction rates. 134-143Sodium ions have been reported to increase the activities of soluble pancreatic and Aspergillus wentii lipases, 144,145but to partially inhibit the lipolytic activity of two different lipases from Aspergillus niger. 141,146

In combination with free fatty acids and the corre-

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Review sponding calcium soaps, bile salts (which act to emulsify fats in the digestive tract of mammals in the presence of the pancreatic lipase/colipase complex 147,148) form soluble, mixed micelles which accelerate the diffusion of products away from the hydrophobic interface. 147Hence the activity of some lipases is enhanced by bile salts (e.g. Phycomyces nitens 149). Bile salts have also been shown to shift the optimum pH of pancreatic lipase. 150The lipase from Rhizopus arrhizus 151is inhibited by bile salts, but lipases from pancreas, m Pseudomonas, 85 Chromobacterium, ~53,~54Streptococcus, 143 and Mucor 155are not. Although the activity of a lipase from Pseudomonas fluorescens towards various natural substrates depends strongly on the type of emulsifier used, 156 pancreatic lipase is inhibited by detergents in general, irrespective of structure and charge. 157'158 Reversible inhibition of lipolytic activity by heavy metal cations has been observed upon short exposures to low concentrations of these materials. 159Irreversible inhibition occurs on long exposure to high concentrations of these cations. 139 Ferrous ions were observed to increase the hydrolytic activity of a lipase from Aspergillus niger TM and Streptococcus faecalis, 143 whereas ferric ions inactivated or inhibited lipases from Aspergillus niger, TM Chromobacterium lipase, 153'154'16° Pseudomonas, 95 and Streptococcus faecalis. 143 The positive effects of metal ions could be due to the formation of complexes with ionized fatty acids which change their solubilities and behavior at interfaces, whereas negative effects can be attributed to competitive binding at the active site) 7 Often the lost activity can be restored via the addition of metal-chelating agents. 139 Nonaqueous solvents can have a variety of effects on enzymes: they may bind specifically, compete with substrate binding, dissociate multimers, shift an equilibrium between two enzyme conformations, alter the amount of helix, react with the enzyme, or affect the rate of the catalytic reaction in a number of other ways. 161 Hydrolysis of olive oil by lipases has been reported to be inhibited by the presence of organic solvents. This inhibition has been attributed to competition of the organic molecules with the triglycerides for adsorption at sites at the oil/water interface. 162Lipases from porcine pancreas and Candida cylindracea were reported to be stable for several hours at high temperatures (-100°C) when the reacting medium was a mixture of tributyrin and heptanol with a water content below 0.4%. 163 This enhanced thermostability can be attributed to the fact that the conformational mobility of the enzyme becomes more restricted when very small amounts of water are present164: conformational mobility is necessary for partial unfolding of the lipase. This unfolding is the first step in thermal inactivation. 165 When dehydrated, lipases also lose their ability to catalyse interesterification reactions involving bulky alcohols, probably because they begin to lose the conformational mobility needed to bind these species in the active center. A layer of bound water (or water hydration shell) plays a key role in maintaining the structural integrity and catalytically active conformation of lipases 166since it affects intramolecular salt bridges and

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hydrophobic interactions.167 Therefore, the existence of trace amounts of water in the immediate vicinity of the lipase is a prerequisite for successful functioning of a lipase in a microaqueous system, lu This requirement explains why lipases are not active in solvents which are miscible with water: these solvents impart changes in conformation leading to inactivation of lipases because they extract bound water from the proteinaceous backbones of these enzymes. 114'169 In microaqueous systems in which lipase-catalysed interesterification or ester synthesis reactions are occurring, the state of water binding in the lipase preparation is of much higher technological significance than the total water content thereof. 170For lipases from Mucor miehei and Rhizopus sp., the water binding capacity depends not only on temperature but also on the hydrophilic character of the support on which the lipases are adsorbed 17° and of the additives employed in the immobilization procedure. TM Since hysteresis is observed in the adsorption/desorption behavior of water/lipase systems, special care must be exercised to obtain lipase preparations with a specified degree of water binding.17°

Kinetics of reactions catalysed by immobilized lipases In general, lipases are able to cleave ester bonds of insoluble acylglycerols; this process is termed hydrolysis. Lipases can also catalyse the reverse reaction, i.e. the formation of ester bonds from an alcohol moiety contributed by a polyhydric alcohol (e.g. glycerol), or a derivative thereof, and a carboxylic acid moiety contributed by a long-chain acid (e.g. a fatty acid). This process is termed ester synthesis. In such processes, water molecules are consumed (hydrolysis) or released (ester synthesis). Lipases can also catalyse the cleavage and formation of ester bonds in a sequential fashion (transesterification). In these processes, water molecules are sequentially consumed and released with no net consumption or formation of water. The aforementioned processes are depicted in Figure 1. In hydrolysis and ester synthesis, only two types of hydrocarbon moieties are considered (say, RI and Rz). In alcoholysis and acidolysis, three types of hydrocarbon moieties are considered (say, RI, Rz, and R3): alcoholysis consists of the exchange of the hydrocarbon moiety linked to the ether group (i.e. R z) with another hydrocarbon moiety contributed by an alcohol (i.e. R3), whereas acidolysis consists of the exchange of the hydrocarbon moiety linked to the carbonyl group (i.e. R 0 with another hydrocarbon moiety contributed by an acid (i.e. R3). In interesterification, four types of hydrocarbon moieties are involved (say, RI, R2, R3, and R4): interesterification consists of the mutual exchange of the hydrocarbon moieties linked to the ether groups (i.e. Rz and R4) or, equivalently, of the hydrocarbon moieties linked to the carbonyl groups (i.e. RI and R3). I f N different types of fatty acid residues are considered, the general form of the rate equation describing the change with time of the concentration of any re-

Immobilized lipase kinetics: F. X. Malcata et al.

R1COOR2 R1COOH

RICOOH R1COOR2

R2OH HOH Hydrolysis

HOH R,2OH Ester synthesis RlC O O ~ C O O R 2

R1C O ~ C O O R 3

R~COOH~ , O H / R 3COOH N / Acidolysis

R2OH~ R 3 O H Alcoholysis

R3COOR2NNN~CO~'/R3COOR4 X

/

Transesterification Figure 1 Schematic representation of the reactions catalysed by lipases. Ri (i = 1, 2, 3, 4) denotes a hydrocarbon moiety. The diamonds enclose the processes that take place in the active site of the enzyme but do not contribute to the overall stoichiometry. The compounds located outside the diamonds denote reactants or products for the reaction in question

actant or product species includes the concentrations of N free fatty acids, glycerol, water, N monoglycerides of each of three different types (i.e. sn- 1, sn-2, and sn-3), N 2 diglycerides of each of three different types (i.e. sn-l,2, sn-l,3, and sn-2,3), and N 3 triglycerides (i.e. sn-l,2,3). Hence the total number of species that should be included in the rate expression is given by (N + 1)[1 + (N + 1)2]. For three of the most common natural sources of lipids--olive oil, beef tallow, and butter fat--the total number of fatty acid residues existing in non-negligible molar concentrations (i.e. larger than 0.5%) is 5, 6, and 11, respectively.l These numbers lead to total numbers of 222, 350, and 1,740, respectively, for the different species to be included as either reactants and/or products in the corresponding rate expression! To the aforementioned functionality of the reactive species, one must still add the dependence on the operating temperature of the system, the actual pH of the solution (in the case of macroaqueous systems) or the pH at the time of water removal by a hydrophobic organic solvent (in the case of microaqueous solu-

tions), 172 the type of solvent used (if any), and the concentrations of species which, though chemically inert in terms of hydrolysis or ester synthesis, interact with reactants, products, or the immobilized enzyme, thus changing the reactivity of the system (e.g. calcium cations which complex the free fatty acids). Finally, the form of the rate expressions may be further complicated by the occurrence of spontaneous (nonenzymatic) side reactions that affect the overall rate of the chemical transformation catalysed by the lipase (e.g. thermal deactivation of the enzyme 173and intramolecular acyl migration97). The search for rate expressions containing so many parameters presents an intractable problem because enormous amounts of experimental data would be required and the numerical work necessary to conduct the nonlinear regression analyses would be prohibitive. Consequently, most kineticists have elected to either focus their attention on model systems which contain only a few different chemical species or analyze naturally occurring complex systems using approaches that lump the various chemical species into a few representative groups. Each of these approaches is discussed below. Several mechanisms have been proposed for lipasecatalysed hydrolysis reactions. 27 Most of these mechanisms have been developed for soluble lipases acting on insoluble substrates (e.g. oil droplets dispersed in water). In many cases these mechanisms may be easily extended to encompass the case where the lipase is present in an immobilized state. The most commonly accepted mechanisms are described next in order of increasing degree of complexity.

Approaches assuming the substrate possesses independent labile ester bonds Since most of the naturally occurring substrates for lipases contain more than one ester bond which is susceptible to hydrolysis, a problem arises as to the order in which the various ester bonds will be attacked. In free states, pancreatic l i p a s e 174-176 and a lipase from Mucor miehei 177 hydrolyse triglycerides much more rapidly than diglycerides and monoglycerides, whereas lipases from swine and rat adipose tissue, ~°° and from bovine adrenal c o r t e x 178 and corpus l u t e u m , 179 hydrolyse diglycerides much more rapidly than triglycerides and monoglycerides. Lipases from Rhizopus delemar and Aspergillus niger 97 and from lingual and insect tissues ~8° attack the sn-3 position first, and then the sn-1 position. Release of the fatty acid esterified at the sn-2 position is possible only after spontaneous intramolecular transfer of this residue to one of the terminal positions via an isomerization reaction. 97 On the other hand, lipases from Geotrichum candidum and Penicillium cyclopium attack the sn-3 and sn-2 positions equally well. 97 The orders of attack are reversed for esterification reactions. 98 Relative rates of interesterification catalysed by a lipase from Mucor miehei immobilized by ion exchange on a resin were in the order: long-chain alcohol > fatty acid > triacylglycerol

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Review > methyl ester > glycerol. ~8~ The relative activities of pancreatic lipase towards soluble substrates remain essentially unchanged upon immobilization (i.e. triglycerides are hydrolysed faster than diglycerides, and diglycerides faster than monoglycerides); however, the relative activities of the same lipase on insoluble substrates exhibit an inversion upon immobilization (i.e. monoglycerides are hydrolysed faster than diglycerides, and these species are hydrolysed faster than triglycerides). 144A lipase from Pseudomonasfluorescens adsorbed on polypropylene catalysed ester synthesis reactions of glycerol and oleic acid to give a final product composed of mono-, di-, and triglycerides in an approximate molar ratio of 3 : 4 : 1.182 The hydrolysis of tributyrin in aqueous solutions as catalysed by a number of immobilized lipases has been described by a sequence of irreversible pseudo-firstorder reactions. 183The values of the pseudo-first-order rate constants for the conversion of dibutyrin to monobutyrin (kz) were two orders of magnitude greater than those associated with the hydrolysis of tributyrin (k) and monobutyrin (k3) for the lipase from Candida cylindracea immobilized by covalent attachment on Sepharose or by adsorption on Celite, and for the lipase from C. antarctica immobilized by adsorption on a macroporous acrylic resin. 183 For lipase from Mucor miehei immobilized by anion exchange on a phenolformaldehyde macroporous basic resin, these rate constants were of the same order of magnitude. 183For the free lipase from Candida cylindracea, Otero et al.183 reported that k~ > k 2 but that after immobilization k 1 becomes smaller (2- to 150-fold) than k 2. This effect was attributed to the use of a porous support that hampers the diffusion of dibutyrin out of the pores, thus facilitating its hydrolysis because of its higher local concentration. This result is consistent with the results of Mattiason and Mosbach j84 who found that when three enzymes, each of which catalyses one of a sequence of three consecutive reactions, are coimmobilized on the same support, they become more efficient than the corresponding combination of soluble enzymes. Otero et a1.~83also reported that the ratio k2/k 3 increases with the hydrophobic character of the support chosen to immobilize the lipase.

Approaches assuming that the labile ester bonds o f the substrate can be treated in a lumped fashion An assumption which is widely used in the analysis of kinetic data for the hydrolysis of glycerides is that the rate of attack on each labile ester bond in the glycerol backbone is equally probable. This assumption implies that the total molar concentrations of substrates such as triglycerides (CTI ,3), diglycerides (CDI 2' CDI 3' and CD2~, corresponding t/J sn-1,2, sn-1,3, and sn-2,3 diglycericles, respectively), and monoglycerides (CMj, Cry:, and CM3, corresponding to sn-1, sn-2, and sn-3 monoglycerides, respectively) can, in principle, be replaced in lumped rate equations by the total molar concentration of the corresponding ester bonds susceptible to 432

EnzymeMicrob. Technol., 1992, vol. 14, June

attack by lipase. That is to say, the effective concentration of labile ester bonds is (3CTI23 + 2CD12 + 2CD13 + 2CD~.3 + Cra~ + CM2 + CM3) for'iionspeciflc lipases', (2CTl23 +CD~z + 2CD~3+CD23 + CM~+CM3) for sn-l,~' specifiE lipases, ~nd (CT123 + CDj2 + CD23 + CM2) for sn-2 specific lipases, respectively. In the' absence of spontaneous intramolecular migration of the acyl moiety (as is often the situation under common experimental conditions ~85)and for the case where the initial concentrations of diglycerides and monoglycerides are both negligible (as is often the case for natural feedstocks1), the above expressions for sn-l,3 and sn-2 specific lipases can be simplified to yield (2CT~ 23 + CDI 2 + CD2 3) and CTI 23, respectively. This is equivalent to ~aying ffiat the afo?ementioned substrates are replaced by a lumped substrate which can be simply termed a glyceride. Most of the mechanisms described below lead to rate expressions which are based on this simplifying assumption.

Mechanisms based on single unimolecular enzyme-substrate complexes. The simplest kinetic model of lipasecatalysed reactions of ester bonds can easily be derived from the classic Michaelis-Menten mechanismS86: kl

E+S,

k- I

'ES

kcat

~E+P+Q

(1)

where E denotes the immobilized enzyme, S the substrate (glyceride), ES the enzyme-substrate complex, P a product species containing one more alcohol moiety than reactant S, and Q a free fatty acid. The rate of disappearance of glyceride per unit volume of reacting fluid (-dCs/dt) can be represented in terms of this mechanism as

dCs dt

_

Umax C S Km + Cs

(2)

The kinetic parameters appearing in this expression are defined as Vmax --- kcatCEtotand K m --- (kcat + k O/k 1. The parameter Vmaxis the rate observed when the lipase is saturated with substrate, Km is the Michaelis-Menten constant, and the subscript tot denotes the total amount of enzyme present in either the E or ES form. Values reported in the literature for Vmax and K m for immobilized lipases are tabulated in Table 1. The apparent value of the parameter K m should decrease as the physical dimensions of the emulsion droplets in suspension decrease. The value of the parameter Vmaxshould increase as the lipase concentration in the bulk fluid increases. 199 However, if the substrate concentration is expressed in terms of interfacial area/ volume rather than in terms of weight/volume, the plots of the reaction rate vs. substrate "concentration" for different emulsions of the same substrate coincide, and one obtains a single value for K m which is independent of the degree of dispersion of the substrate.~99 Simplifications of the above Michaelis-Menten model have been claimed for the case of some shortchain substrates, e.g. tripropionin.13 These simplifications arise when the concentration of substrate in the

Immobilized lipase kinetics: F. X. Malcata et al. Table 1

A p p a r e n t Vr.ax and Km values for lipolytic reactions catalysed by immobilized lipases

Km

T (°C)

0.026-0.15'" 0.896'" 2.5" 2.5" 2.5" 6.0" 141' 20' 50' 0.043" 0.043" 1.6-3.9" 9.2-13.2" 4.4-6.9" 63.6' 2.62' 0.50' 0.48' 0.0266" 140' 92' 390' 420' 2.6"

25 25 45 45 37 37 35 35 35 35 35 20-50 20-50 20-50 50 50 35 35 37 30 30 30 30 37

Vm~x 0.208-2.52* 0.282* n.a. n.a. 0.0434" 0.0306" 0.06476** n.a. n.a. 41.5"** 62.5*** 19.7-23.4"** 64.28-81.88"** 23.24-32.72*** 220* 160" n.a. n.a. 0.6**** 182"** 85*** 3*** 32*** 430-1470"**

n.a.: not available * p,mol/(min mgimmobilized protein) **/~mol/(min Cm~uppo~) ***/~mol/(min g,uppo,) ****/.¢mol/(min mlsoDport)

'/~mol/ml " % w/v

pH

Source of lipase

9.0 Pancreatic 9.0 Pancreatic 8.5 Microbial 8.0 Microbial 8.2 Microbial 7.5 Microbial 7.0 Aspergillus niger 7.5 Candida cylindracea 7.5 C. cylindracea 6.0 C. rugosa 6.0 C. rugosa n.a. Rhizopus arrhizus n.a. Ft. arrhizus n.a. Ft. arrhizus 8.7 R. oryzae 8.7 Ft. oryzae 8.5 Geotrichum candidum 8.5 G. candidum 7.0 Mucorjavanicus n.a. M. miehei n.a. M. miehei n.a. M. miehei n.a. 114.rniehei 5.5 Pseudornonas

BB: butyl butyrate CMC: carboxymethylcellulose DEAEC: DEAE-cellulose DIPE: di-isopropyl ether

EB: ethyl butyrate IA: isopentyl acetate IB: isopentyl butyrate IO: isooctane MBBA: 4-methoxybenzilidene-4'-n-butylaniline PA: polyacrylamide

'" % v/v

vicinity of the immobilized lipase is low, thus leading to kinetics which are pseudo-first-order in both the enzyme and the substrate. For studies employing a lipase from Candida cylindracea, Yamane et al. 20°-2°2 have used a true reversible Michaelis-Menten kinetic expression to relate the rate of release of free fatty acids to the molar concentrations of ester bonds and free fatty acids in the oil phase (initially consisting of pure olive oil). This approach was possible because they employed continuous phases of water and oil partitioned by a polypropylene, hydrophobic membrane. In the case of feedstocks of natural origin which contain multiple chemical species susceptible to the action of lipase (e.g. butter fat), the Michaelis-Menten mechanism denoted as equation (1) must be extended to incorporate competitive inhibition by every substrate, S~.19° For extents of hydrolysis below 75%, the following rate expression results: dCs i .

.

dt

Omax,i Cs i .

.

iv

Y, csj

j=l

(3)

Method of

Solvent or continuous

Support

immobilization

phase

SS PA Agarose Agarose Agarose Agarose PP Agarose PA PEG PPG Mycelia Mycelia Mycelia Alumina Alumina CMC PABC DEAEC Resin Resin Resin Resin MBRA

Cross-linking Covalent Covalent Covalent Covalent Covalent Adsorption Covalent Entrapment Entrapment Entrapment Cell binding Cell binding Cell binding Covalent Covalent Covalent Covalent Ion exchange Ion exchange Ion exchange Ion exchange Ion exchange Entrapment

W W W W W W W W W IO + 10% W I0 + 10% W DIPE + 0.07% W DIPE + 0.17% W DIPE + 0.37% W W W W W W Hexane Hexane Hexane Hexane TMAC

Substrate

Ref.

TB TB Olive oil Butter oil Olive oil Butter oil Butter oil Olive oil Olive oil Olive oil Olive oil Olive oil Olive oil Olive oil TB TB + PVA Olive oil Olive oil Olive oil EB BB IA IB SMPOE

187 187 188 188 189 189 190 191 191 192 192 193 193 193 194 194 195 195 196 197 197 197 197 198

PABC: p-aminobenzylcellulose PEG: polyethylene glycol PP: polypropylene PPG: polypropylene glycol PVA: polyvinyl alcohol SMPOE: sorbitan monolaureate poly(oxyethylene)ether SS: stainless steel TB: tributyrin TMAC: tetramethylammoniurn chloride, aqueous solution W: water

where the constants are defined by /)max,/~ kcat,i CE,ot• The above rate expression is based on the assumptions that: (i) the Michaelis-Menten constants, Km,;, associated with every substrate S; (out of N possible substrates) are approximately equal to one another, and (ii) all Cs/Kmi are very large compared to unity. 19° Values of the several individual Vmax/ were correlated m terms of the number of carbon atoms in the hydrocarbon moiety of the corresponding fatty acid residue according to an empirical expression based on a F-type probability distribution• 190 For a simple enzymatic reaction in solution, the maximum intrinsic rate of reaction is limited by the rate at which lipase and substrate come together in the proper orientation. ~°3 For the case of hydrolysis reactions, the substrate is often part of the disperse phase of an emulsion, a micelle, or a monolayer which contacts water. These structures may be orders of magnitude larger in size than the supported enzyme for the case where the carrier exists in powdered form. Thus, the maximum attainable rate is limited by the amount of lipase which can interact with the substrate continuum. 173 This phenomenon, which is similar in nature to

Enzyme Microb. Technol., 1992, vol• 14, June

433

Review Table 2

Half-lives o f i m m o b i l i z e d lipases

Half-life (h)

Type of reaction

1 0.12 477 36O 144-360 110-1040 223-816 5,680 4,737 1,157 167-326 184 50O 87.6 1,019 504-1248 0.21-4.81 1,367 1,600 1,296 552 240-480 33-138 768 120 93.6-180 26.4 93-231 120 288 10 6 17.5-362 101-1,512

Hydroiysts Hydrolysis Hydrolysts Hydrolysm Hydrolys=s Hydrolysis Hydrolysm Hydrolysts Hydrolysis Hydrolysm Hydrolysis Hydrolysis Hydrolysis Hydrolysts Hydrolysis Ester synthesis Hydrolysis Interestification Ester synthesis Ester synthesis Interesterification Ester synthesis Ester synthesis Hydrolysis Hydrolysis Hydrolysis Hydrolysis Hydrolysis Interesterification Interesterification Hydrolysis Hydrolysis Hydrolysis Hydrolysis

Source of lipase Pancreas Porcine pancreas Aspergillus niger Candida cylindracea C. cylindracea C. cylindracea C. cylindracea C. cylindracea C. cylindracea C. cylindracea C. rugosa C. rugosa C. rugosa C. rugosa C. rugosa Chromobacteriurn viscosum Mucor javanicus Mucor miehei M. rniehei M. miehei M. miehei M. rniehei M. miehei Rhizopus Rhizopus Rhizopus Rhizepus R. arrhizus R. delemar t7. delemar R. microsperus-EC R. rnicrosporus-IC R. oryzae Thermomyces lanuginosus

Support Spherosil Cellulose Polypropylene Polypropylene Polypropylene Polypropylene Polypropylene Polypropylene ENTP ENTP PVC Chitin Chitosan Agarose Cellulose Polypropylene DEAE-cellulose Celite Resin Polypropylene Resin MIR Resin PVC PVC PTFE PTFE Fungal mycelia Celite ENTP AE-cellulose AE-cellulose Alumina Acrylic

Method of binding Adsor )tion Adsor }tion Adsor )tion Adsor :)tion Adsor ~tion Adsor )tion Adsor ation Adsor :)tion Entra )ment Entra )ment Covalent Covalent Covalent Covalent Adsorption Adsorption Ion exchange Adsorption ion exchange Adsorption Ion exchange Ion exchange Ion exchange Entrapment Adsorption Cross-linking Adsorption Cell binding Adsorption Entrapment Adsorption Adsorption Covalent Adsorption

T (°C)

pH

Ref.

51 50 35 40 40 40 40 35 30 30 40 40 40 40 30 40 40-60 40 60-70 40 60 40 20-70 37 37 37 37 30 50 50 35 35 35-70 50

8.6 9.0 7.0 7 7.0 7 7 7 7.0 7.0 5.4 5.4 5.4 5.4 6.0 7 5.0 n.a. n.a. n.a. n.a. n.a. n.a. 8.0 8.0 8.0 8.0 n.a. 6.5 6.5 8.5 8.5 8.7 5.5-6.5

234 235 190 201 202 236 236-238 236 239 239 240 240 240 240 241 135 196 113 242 182 243 244 245 28 28 246 246 193 247 247 205 205 194 248

n.a.: not available ENTP: polypropylene glycol EC: extracellular IC: intracellular MIR: microporous anion exchange resin PTFE: polytetrafluoroethylene PVC: polyvinylchloride

an adsorption process, can be schematically represented by equation (1) provided that k 1 and k_l are interpreted as adsorption (kads) and desorption (kdJ constants, respectively. In the case of lipases immobilized on continuous supports, or on discrete supports larger in size than the individual droplets of substrate, the above reasoning remains valid provided that the spacing between neighboring molecules of immobilized lipase is still larger than the area of contact between the droplet and the lipase carrier. For this situation, in contrast to the classic Michaelis-Menten rate expression where the reaction rate increases linearly with the total enzyme concentration, a limiting rate is approached as the lipase concentration is increased. In the present case a balance on the total number of adsorption sites is more relevant than a balance on the total number of active

434

Enzyme Microb. Technol., 1992, vol. 14, June

sites. 173 This approach leads one to the following rate expression:

dCs = Umax C E dt Km+ CE

(4)

where the constants are defined as V~ax--- kcat Cs and K m =- kdes/kads, and where the physical interpretations of the constants are as follows: Vmaxis the rate when the adsorption sites on the surface of the fat globule are saturated with lipase, and Km is a pseudo-Michaelis-Menten constant for the above rate expression. When a lipase from Candida rugosa was immobilized by adsorption on cellulose, values for V-ax and K m changed from ca. 6.0 to 2.7 ttmol m-2s-t'i and from ca. 0.12 to 0.0040 g m -:, respectively. 2°4 The primary mechanistic distinction between this mechanism and the simple Michaelis-Menten mechanism is that in the

Immobilized lipase kinetics: F. X. Malcata et al. Table 3

pH optima for reactions catalysed by immobilized lipases as compared with those for the corresponding free lipase

Optimum pH before immobilization

Optimum pH after immobilization

8.5 8.5

8.5 8.0 7.5-8.5 8.0-8.5 6.62 7.5 7.5 4.0 7.0 7.0 6.0 8.5 8.5 7.5 8.6 7.0 8.75 8.5 6.3 6.3 6.4 5.0 7.0 7.5 10 10 7.0 6.5 6.5 6.5 7.5 8.5-8.6 8.8

n.a.

7.5-8.0 n.a. 7.5 7.5 4.0 7.0 5.5 5.5 7.5 7.5 7.5 7.5 n.a. 8.5 8.5 6.5 6.5 6.5 7.0 n.a. 7.5 10 10 5.5 5.5 n.a. n.a. 7.5 8.5-8.6 7.5

Source of lipase

Support

Method of binding

T (°C)

Ref.

Pancreas Pancreas Cotton plant Microbial Aspergillus niger Candida cylindracea C. cylindracea C. cylindracea C. cylindracea C. rugosa C. rugosa C. rugosa 6". rugosa C. rugosa C. rugosa C. rugosa Geotrichurn candidum G. candidum Humicola lanuginosa H. lanuginosa H. lanuginosa Mucor Pseudomonas P. fluorescens Rhizopus Rhizopus R. arrhizus R. arrhizus R. delemar R. delernar R. delernar R. microsporus R. oryzae

Stainless steel Polyacrylamide n.a. Agarose Polypropylene Agarose Polyacrylamide HDPE Silica gel ENT ENTP Chitosan Chitin Agarose PVC Sephadex CMC PABC Alginate ENTP Amberlite MBBA-collagen MBBA Porous glass PVC PVC ENT ENTP Celite ENTP Porous glass AE-cellulose Alumina

Cross-linking Covalent n.a. Covalent Adsorption Covalent Entrapment Adsorption Adsorption Entrapment Entrapment Covalent Covalent Covalent Covalent Adsorption Covalent Covalent Entrapment Entrapment Adsorption Entrapment Entrapment Covalent Entrapment Adsorption Entrapment Entrapment Adsorption Entrapment Covalent Adsorption Covalent

n.a. n.a. 36 n.a. 35 35 35 42 n.a. 35 35 37 37 37 37 30 35 35 45 45 45 37 37 37 37 37 35 35 40 40 37 35 50

187 187 144 188, 189 190 191 191 236 249 192 192 240 240 240 240 263 195 195 264 264 264 265 198 266 28 28 192 192 247 247 266 205 194

n.a.: not available CMC: carboxymethylcellulose ENT: polyethylene glycol ENTP: polypropylene glycol HDPE: high-density polyethylene MBBA: 4-methoxybenzilidene-4'-n-butylaniline PABC: p-aminobenzylcellulose PVC: polyvinylchloride

present case adsorption of lipase at the fluid solid interface (i.e. contact with the substrate molecules) is independent of catalysis in the interracial plane. Observed Km values for lipases may thus reflect the extent of adsorption of the lipase at the lipid/water interface rather than the affinity between enzyme and substrate at the active site. 34 It has been reported that the substrate may also function as a competitive inhibitor of the lipase. 2°5 In this situation, the following mechanism has been proposed:

E+S~ ES + S¢

Km

) E S k~"t~E + P + Q

Ks

>SES

flkcat)

ES + P + Q

The corresponding rate equation is

(5)

dC s dt

_

kcat CEtot [1 + (fl Cs/Ks)] Cs Km+ Cs + (C2s/Ks)

(6)

The values of the kinetic parameters were reported to depend on a number of experimental conditions, such as the size of the emulsified particles. 2°5 Experimental results obtained by a number of authors12'13'206'207 have demonstrated the existence of a lag (or induction) period between the addition of a lipase to an emulsion and the start of substrate depletion. It has been suggested that this period can be attributed to diffusion-controlled adsorption of the lipase at the lipid/water interface. 13 Longer induction periods may be interpreted as a consequence of a necessity for pseudoactivation of the lipase by improvement of the dispersion of the substrate or of promotion of enzyme/ substrate interactions through charge effects. 33 A sim-

Enzyme Microb. Technol., 1992, vol. 14, June

435

Review Table 4 Optimum operating temperatures for reactions catalysed by immobilized lipases as compared with their free counterparts Optimum T before binding (°C) 32 n.a. 35 35 35 30 36 35-40 37 37 37 37 n.a. 35 35 51 51 51 n.a. n.a. 62 n.a. n.a. 35-40 n.a. 37 39 42 40

Optimum T after binding (°C)

Type of reaction

40 50-55 45 35 35 40 36 45 60 48 43 52 30-35 35 35 51 61 60 37-40 50 62 37 37 45 40 37 37 35 48

Hydrolysis Interesterification Hydrolysis Hydrolysis Hydrolysis Hydrolysis Hydrolysis Hydrolysis Hydrolysis Hydrolysis Hydrolysis Hydrolysis Interesterification Hydrolysis Hydrolysis Hydrolysis Hydrolysis Hydrolysis Hydrolysis Ester synthesis Hydrolysis Hydrolysis Hydrolysis Hydrolysis Interesterification Hydrolysis Hydrolysis Hydrolysis Hydrolysis

Source of lipase Pancreas Cotton plant Microbial Candida cylindracea C. cylindracea C. cylindracea C. cylindracea C. rugosa C. rugosa C. rugosa C. rugosa C. rugosa C. rugosa Geotrichum candidum G. candidum Humicola lanuginosa H. lanuginosa H. lanuginosa Mucorjavanicus M. miehei Pseudomonas fluorescens Rhizopus Rhizopus R. arrhizus R. arrhizus R. delernar R. microsporus-EC R. microsporus-IC R. oryzae

Support

Method of binding

pH

Ref.

Spherosil n.a. Agarose Agarose Polyacrylamide ENTP Silica gel ENTP Chitosan Chitin Agarose PVC Sephadex CMC PABC Alginate ENTP Amberlite DEAE-cellulose Resin Porous glass PVC PVC ENTP Celite Porous glass AE-cellulose AE-cellulose Alumina

Adsorption n.a. Covalent Covalent Entrapment Entrapment Adsorption Entrapment Covalent Covalent Covalent Covalent Adsorption Covalent Covalent Entrapment Entrapment Adsorption Ion exchange Ion exchange Covalent Entrapment Adsorption Entrapment Adsorption Covalent Adsorption Adsorption Covalent

8.6 7.5-8.5 n.a. 7.5 7.5 n.a. 7.0 6.0 8.0 8.0 8.0 8.0 n.a. 8.5 8.5 7.0 7.0 7.0 5.0 n.a. 7.5 10 10 6.0 n.a. 7.5 8.5 8.5 8.7

234 144 188,189 191 191 239 249 192 240 240 240 240 263 195 195 264 264 264 196 245 266 28 28 192 230 266 205 205 194

n.a.: not available CMC: carboxymethylcellulose EC: extracellular ENTP: polypropylene glycol IC: intracellular PABC: p-aminobenzylcellulose PVC: polyvinylchloride

pier explanation of the induction period is based on a requirement that the lipase first be brought into contact with the lipid substrate and then orient itself and/or acquire a more suitable conformation at the oil/water interface before catalysis can occur. 12'20'206,208-212 This mechanism basically consists of two successive equilibria. The first describes the reversible binding of the free lipase to the interface. This binding is believed to be characterized by a Langmuir isotherm. 199 This process has been denoted as p e n e t r a t i o n . 2° It is assumed to convert the lipase to an activated conformation. This conversion to a much more active conformation also agrees with evidence from independent experiments involving a lipase from P s e u d o m o n a s cepacia. 213 After pretreatment with chilled acetone, a lag period was observed when the lipase was brought into contact with substrate dissolved in an organic solvent. The aforementioned penetration step is followed by a second process in which the activated lipase binds a single substrate molecule at the catalytic site. This step is the two-dimensional analog of the Michaelis-Menten 436

E n z y m e M i c r o b . T e c h n o l . , 1992, vol. 14, J u n e

equilibrium established in classical enzyme kinetics. 12'2°6 In terms of this model, all products of the reaction are assumed to (i) be soluble in the aqueous phase, (ii) rapidly diffuse away from the interface, and (iii) induce no change in the enzyme with time. Numerous modifications of the aforementioned mechanism have been proposed. For example, adsorbed lipolytic enzymes may irreversibly denature at the interface 24,65,2°8,214-217 or adsorption of the lipase at the interface may be enhanced by interactions with protein cofactors or soluble ligands. 218

Mechanisms based on single multimolecular, or multiple unimolecular, enzyme-substrate complexes. Since lipases catalyse multisubstrate-multiproduct reactions, the aforementioned rate expressions (which are derived from true or pseudo-Michaelis-Menten mechanisms) are not likely to provide accurate molecular descriptions of the reactions taking place. In order to overcome this modeling difficulty, more complex kinetic mechanisms have been proposed.

Immobilized

l i p a s e k i n e t i c s : F. X. M a l c a t a et al.

Table 5 Optimum water activity for reactions catalysed by immobilized lipases Optimum water concentr, 4"*

1.1"* 0.75** 1.7"* 11"* 1.3"** 12.5"*** 23.3**** 12.5"*** 0.083** 0.042** 0.047** 0.050** 0.091"* 0.093** 0.079** 0.098** 0.082** 2.2** 0.17"* 0.75* 3.0** 3.0** 1"* * aw ** %(w/v) *** %(v/v) **** tool %

Type of reaction

Source of lipase

Ester synthesis Ester synthesis Ester synthesis Ester synthesis Interesterification Ester synthesis Ester synthesis Ester synthesis Ester synthesis Ester synthesis Ester synthesis Ester synthesis Ester synthes=s Ester synthesis Ester synthesis Ester synthesis Ester synthesis Ester synthesis Ester synthesis Hydrolysis Hydrolysis Interesterification Interesterification Ester synthesis

Chromobacterium viscosum C. viscosurn C. viscosurn C. viscosurn Mucor miehei M. rniehei M. rniehei M. miehei M. miehei Pseudornonas fluorescens P. fluorescens P. fluorescens P. fluorescens P. fluorescens P. fluorescens P. fluorescens P. fluorescens P. fluorescens P. mephitica Rhizopus arrhizus R. arrhizus R. delernar R. delemar R. niveus

Support

Method of binding

Solvent or continuous phase

T (°C)

Ref.

Polypropylene Dowex Spherosil DEA DEAEc Resin MIR Resin Resin Resin Celite Celite Celite Celite Celite Celite Celite Celite Celite Dowex Fungal mycelia Celite Celite ENTP Dowex

Adsorption Covalent Covalent Covalent Ion exchange Ion exchange Ion exchange Ion exchange Ion exchange Adsorption Adsorption Adsorption Adsorption Adsorption Adsorption Adsorption Adsorption Adsorption Covalent Cell binding Adsorption Adsorption Entrapment Covalent

Glycerol/OA Glycerol/RBA Glycerol/RBA Glycerol/RBA Olive oil Olive oil Butanol Butanol Butanol Benzene Benzene/A Benzene/S Benzene/E Benzene/P Benzene/L Benzene/BSA Benzene/C Benzene/D Glycerol/RBA DIPE n-Hexane n-Hexane n-Hexane Glycerol/RBA

40 60 60 60 60 40 20 50 70 40 40 40 40 40 40 40 40 40 60 30 36.5 40 40 40

135 270 270 270 242 244 271 271 271 171 171 171 171 171 171 171 171 171 270 193 230 247 247 270

A: arabitol BSA: bovine serum albumin C: casein D: dextran DEAEc: DEAE cellulofine DIPE: di-isopropyl ether E: erythriol

By analogy with the mechanisms associated with catalysis by other serine hydrolases, it has been suggested69,~ 14that the hydrolytic action of lipases follows a two-step reaction mechanism, usually referred to as Ping Pong Bi-Bi2~9: the first step is a nucleophilic attack of the serine hydroxyl group on the ester bond resulting in formation of an acyl enzyme and release of the alcohol moiety of the original substrate; the second step is hydrolysis of the acyl enzyme. 34It is strongly supported by kinetic studies involving pancreatic lipase 22°'221 and lipase from Candida cylindracea. 222 Isolation of an acyl-enzyme intermediate was actually reported for pancreatic lipase. 223These studies have also demonstrated that, depending on the substrates and the acyl acceptors used, either of the two steps (i.e. acylation or deacylation) can be rate-limiting. In the case of a lipase from Chromobacterium, 224 the observed rate expression corresponds to an Ordered Bi-Bi mechanism 2~9 in which the rate-controlling step is acylation of the enzyme. The distinction between Ordered Bi-Bi and Ping Pong Bi-Bi mechanisms may be somewhat artificial because the water molecule, being highly polar, may have difficulty in penetrating the active site where a bulky substrate (i.e. a glyceride molecule with a longchain fatty acid residue) is already bound. Therefore,

ENTP: polypropylene glycol L: lactose MIR: microporous anion exchange resin OA: oleic acid P: phosphatidylcholine RBA: rice bran acid S: sorbitol

a more reasonable picture might correspond to having a water molecule bind to a hydrophilic pocket at the active site (e.g. by hydrogen bonding to the aspartate residue of the active site) before the glyceride molecule binds. The small water molecule might remain in this pocket until it is needed for the deacylation step. In macroaqueous environments, water forms a monolayer in the immediate vicinity of the active site, while in microaqueous systems a pseudo-monolayer (which consists of clusters of water molecules around charged groups of the enzyme) is present. 172Consequently, the concentration of water may be taken as approximately constant (provided that a reservoir of water molecules is available within a small distance of the active site). If this situation prevails, the rate expressions for the aforementioned Ordered Bi-Bi and Ping Pong Bi-Bi mechanisms degenerate to that corresponding to an Ordered Uni-Bi mechanism. ~9 By contrast, Wang 225 has suggested a Random Uni-Bi mechanism in which the release of alcohol and the acyl product occurs via a random release mechanism rather than sequentially, with the alcohol leaving prior to the acyl product. The hydrolytic action of lipases from Aspergillus niger 226 and Candida cylindracea 29 immobilized by adsorption on microporous polypropylene membranes on melted butterfat has been modeled using a generic Ping E n z y m e M i c r o b . T e c h n o l . , 1992, v o l . 14, J u n e

437

Review

Table 6 Effects of different ions on the activity of immobilized lipases Salt or ion

Concentration (mol 1-1)

Percent activity

Source of lipase

pH

T (°C)

Support

Method of binding

Ref.

(NH4)2SO4 NaCI KCI CaCI2 MgCI2 BaCI2 FeCI2 HgCI2 Fe2÷ Fe3+ Cu2+ Fe2÷ Fe3+ Cu2+ FeCI3 NaCI CaCI2 Ca(CH3C02)2

0.025 0.025 0.025 0.025 0.025 0.025 0.025 0.025 0.0001-0.01 0.0001-0.01 0.0001-0.01 0.0001-0.01 0.0001-0.01 0.0001-0.01 0.005-0.03 0.001-10 0.005 0.020 0.020

94.1 94.7 84.4 57.0 117.6 57.4 22.9 0.9 23-39 15-38 26-49 37-63 14-46 29-69 8.1-88 147-500 19-40 41-51.1 36

Candida rugosa C. rugosa C. rugosa C. rugosa C. rugosa C. rugosa C. rugosa C. rugosa Geotrichum candidum G. candidum G. candidum G. candidum G. candidum G. candidum Mucorjavanicus Pseudomonas fluorescens Thermomyces lanuginosus T. lanuginosus T. lanuginosus

7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 8.5 8.5 8.5 8.5 8.5 8.5 7.0 7.0 6.5-7.5 6.5-7.5 6.5-7.5

30 30 30 30 30 30 30 30 35 35 35 35 35 35 20 40 50 50 50

Sephadex Sephadex Sephadex Sephadex Sephadex Sephadex Sephadex Sephadex CMC CMC CMC PABC PABC PABC DEAEC Celite Acrylic Acrylic Acrylic

Adsorption Adsorption Adsorption Adsorption Adsorption Adsorption Adsorption Adsorption Covalent Covalent Covalent Covalent Covalent Covalent Ion exchange Adsorption Adsorption Adsorption Adsorption

263 263 263 263 263 263 263 263 195 195 195 195 195 195 196 171 248 248 248

Na(CH3C02) 2

CMC: carboxymethylcellulose DEAEC: DEAE-cellulose PABC: p-aminobenzylcellulose

Pong Bi-Bi mechanism. After testing different forms of this mechanism corresponding to different rate-controlling steps (i.e. binding of the glyceride, acylation of the enzyme, release of the alcohol moiety, binding of the water, deacylation of the enzyme, and release of the free fatty acid), it was found that the form that best fits the experimental data corresponds to deacylation of the lipase as the rate-controlling step.29 In these circumstances, the following mechanism can be written: kI

E+G<

>EG k- I k2

EG ~

' FP k- 2 k3

FP<

>F+P k_ 3

F + W,

k4

>FW

k- 4

FW

k5

> EQ

k6

EQ,

k- 6

'E + Q

(7)

where G denotes a glyceride molecule and W a water molecule, E and F denote native and acylated forms of lipase, Q and P denote a free fatty acid and a product species containing one less ester bond than the glyceride G, EG and FW are enzyme/reactant complexes,

438

E n z y m e M i c r o b . T e c h n o l . , 1992, v o l . 14, J u n e

and FP and EQ are enzyme/product complexes. The associated rate expression is of the form

dCG dt

OICGCw Cp + O2C G + 03 C GcP + 04C G C w + 05 CoCp

(8) where the various 0i values are relatively simple functions of the rate constants (k,.) for the mechanistic equations and the total concentration of enzyme. Malcata et al. 226-229have modeled their data using a general Ping Pong mechanism and two nested simplifications thereof (i.e. deacylation controlling and acylation controlling). In the pH range 3.0-9.0 at 40°C, and in the temperature range 40-60°C at pH 7.0, the best fit of the kinetic data is provided by a rate expression similar in form to a Michaelis-Menten mechanism with product inhibition. (This rate expression actually results from the assumption of a Ping Pong Bi-Bi mechanism with constant concentration of water and with the rate-controling step being deacylation.) Distortion theory ~°asuggests that the probability that a fatty acid residue with a given length and degree of unsaturation will release the exact amount of energy required for the hydrolysis reaction should be expected to follow a gaussian distribution with respect to the number of carbon atoms contained in the residue. However, a multiplicative factor must be used to correct for the extra degree of hydrophobicity associated with the presence of double bonds. 227 On the other hand, the

Immobilized lipase kinetics: F. X. Malcata et al. Table 7

Effects of different solvents on the activity of immobilized lipases

Solvent

Standard

Percent activity

Source of lipase

pH

T (°C)

Support

Method of binding

Ref.

Cyclohexana Hexane Heptane Octane Nonane Decane Isopropylether Cyclohexane Hexane Heptane Octane Nonane Decane Isopropylether Benzene Toluene Tetrahydrofuran Acetic acid Methylcyanide MIBK Dimethylformamide Dioxane Toluene Chloroform Dioxane Acetone Dimethylformamide Toluene Chloroform TCE Diethyl ether

Isooctane Isooctane Isooctane Isooctane Isooctane Isooctane isooctane Isooctane Isooctane Isooctane Isooctane Isooctane Isooctane Isooctane Hexane Hexane Hexane Hexane Hexane Hexane Hexane Hexane Benzene Benzene Benzene Benzene Benzene Benzene Benzene Benzene Heptane

71.5-84.8 42.9-52.2 54.1-54.3 59.1-52.8 63.4-66.4 60.7-61.9 42.5-47.8 50.5-87.4 24.0-67.0 36.1-67.1 37.4-60.1 44.6-65.7 47.5-66.7 10.5-43.6 61.4 75.7 2.6 5.7 4.3 77.1 2.5 0.86 87 67 23 0 0 24 50 962 516-770

Candida rugosa C. rugosa C. rugosa C. rugosa C. rugosa C. rugosa C. rugosa C. rugosa C. rugosa C. rugosa C. rugosa C. rugosa C. rugosa C. rugosa Mucor miehei M. rniehei M. miehei M. miehei M. miehei M. rniehei M. miehei M. rniehei Pseudomonas fluorescens P. fluorescens P. fluorescens P. fluorescens P. fluorescens P. fragi P. fragi P. fragi Rhizopus arrhizus

7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 n.a. n.a. n.a. n.a. n.a. n.a. n.a. n.a. n.a. n.a. n.a. n.a. n.a. 8.0-8.5 8.0-8.5 8.0-8.5 n.a.

30 30 30 30 30 30 30 30 30 30 30 30 30 30 60 60 60 60 60 60 60 60 n.a. n.a. n.a. n.a. n.a. 25 25 25 37

Sephadex Sephadex Sephadex Sephadex Sephadex Sephadex Sephadex Amberlite Amberlite Amberlite Amberlite Amberlite Amberlite Amberlite Resin Resin Resin Resin Resin Resin Resin Resin PEG PEG PEG PEG PEG PEG/M PEG/M PEG/M Mycelium

Adsor ~tion Adsor ~tion Adsor ~tion Adsor ~tion Adsor ~tion Adsor ~tion Adsor ~tion Adsor 3tion Adsor 3tion Adsor 3tion Adsor ~tion Adsor ~tion Adsor ~tion Adsor ~tion Ion exchange Ion exchange Ion exchange Ion exchange Ion exchange Ion exchange Ion exchange Ion exchange Covalent Covalent Covalent Covalent Covalent Covalent Covalent Covalent Cell binding

169 169 169 169 169 169 169 169 169 169 169 169 169 169 114 114 114 114 114 114 114 114 168 168 168 168 168 168 168 168 107

n.a.: not available M: magnetite PEG: polyethyleneglycol TCE: 1,1,1-trichloroethane

free energy of binding should be a very weak function of both the number of carbon atoms, nc, and the number of double bonds in the hydrocarbon moiety of the corresponding fatty acid residue. Studies of the lipolysis of milkfat by a lipase from Aspergillus niger immobilized on polypropylene fibers indicate that Vmaxchanges with nc according to a bell-shaped distribution centered approximately at C10 : 0. They also indicate that each double bond is approximately equivalent to nearly 6 extra carbon atoms in the hydrocarbon backbone of the fatty acid residue. 227 In spite of an abundance of experimental data for interesterification reactions, modeling of these reactions has lagged far behind that for hydrolysis-ester synthesis reactions. Interesterification reactions are believed to proceed through intermediate stages involving hydrolysis and ester synthesis reactions. The rate of interesterification has been reported to be proportional to the rate of hydrolysis. 222'23° For more extended ranges of operation using lipases from Aspergillus sp. TM and Rhizopus arrhizus 232 immobilized by adsorption on diatomaceous earth, the rate of interes-

terification follows a Michaelis-type dependence on the lipolytic activity. The aforementioned kinetic models may be used as starting points for developing models for interesterification reactions. Recently, the following mechanism has been proposed for the acidolysis of olive or butter oil with caprylic acid as catalysed by a lipase from Pseudomonas cepacia immobilized by precipitation onto a microporous polypropylene powder213: k1

E + G1 ~

EGl,

EQ1-I ~

k2 k3

k4 k5 k6

~ EG1

~ EQI-I

' EQ1 + I

k7

EQI + W ,

k8

~E+Q1

Enzyme Microb. Technol., 1992, vol. 14, June

439

Review *9 >EQ z + W klo

E + Qz'

EQ2 + I<

EQ2-I <

EG2 <

kll k12

>EQ2-I

k13

k14 k15 ki6

> EG 2

' E + G2

(9)

where W denotes a water molecule, I denotes an intermediate species containing one more alcohol moiety than the original reactant glyceride GI and the product glyceride G2, Q~ denotes a generic fatty acid residue of the triglyceride feedstock, and Q2 denotes the free fatty acid supplied to the reactive system. The above mechanism can be described by the following rate expressions:

dCQ 1

Vmax,f,lCGI Cw -- Vmax.r,2 Cl CQI

dt

(1 + KiCl)(CQi "[- CQ2)

dCQ2

Umax,r,lCQ2 C1 -- Umax,f,2 CG2 Cw

dt

(l +

KiCI)(CQI

+

CQ2)

(10)

where the kinetic parameters Umax and K; are simple functions of the elementary rate constants (ki) and the total concentration of enzyme. Parameter values were reported for the kinetic parameters for interesterification of milkfat with caprylic acid and with linolenic acid, as well as for the interesterification of olive oil with caprylic acid. 213

Deactivation processes Lipases are known to lose activity as time elapses. In the simplest models of deactivation phenomena, an active lipase undergoes reversible or irreversible structural changes (temperature-induced conformational transitions) or chemical changes (e.g., deamidation and hydrolysis of side groups of amino acid residues, destruction of disulfide bonds, or isomerizations to form inactive structures) to produce a catalytically inactive f o r m . 233 Such processes are frequently characterized by first-order kinetics. ~7aLiterature values for the halflives of immobilized lipases (defined as ln{2}/kd, where kd is the first-order deactivation constant) are tabulated in Table 2. Half-lives ranging from 7 min. to ca. 7 months have been reported. The large variability is due not only to the different nature of the lipases, supports, and immobilization procedures, but also to the multitude of operating conditions that can be employed (e.g. pH of the buffer, presence or absence of substrate, type of substrate, presence or absence of polyhydric

440

Enzyme Microb. Technol., 1992, vol. 14, June

alcohols, and properties of organic solvent used, if any). It is possible to significantly increase the thermal stability of a lipase by immobilizing it on a solid support. 249Moreover, the stability of the enzyme can often be further increased by several orders of magnitude if it is transferred from a water-rich environment to an organic medium. ~63 This phenomenon has been explained by postulating that when the lipase is transferred to an organic solvent, the process of dehydration causes the lipase to be locked in its native, catalytically active conformation. 172 Frequently, immobilized lipases are more stable than their free counterparts because the enzyme molecule becomes more rigid after multipoint attachment to a solid carrier. Hence unfolding is greatly hindered and disruption of the active center becomes much less likely to o c c u r . 25°251 Activation energies associated with the deactivation of immobilized lipases vary from ca. 75 to 144 kJ mol-1.28,194,196,240 Such values are lower than the range usually associated with thermal deactivation of enzymes 252 because most lipases are stabilized by immobilization. Although these first-order deactivation models are widely employed in the biochemical literature, they are not adequate t6 describe the behavior of enzymes in a number of instances. ~73253Hence, more complex models of enzyme deactivation have been proposed. 254-26~ These models can be viewed as derived from a general multistep model which comprises irreversible or reversible rearrangements between the various active forms of enzyme coupled with first-order irreversible or reversible deactivation of the labile f o r m s . 253 Extra complexity is more likely to be observed in the case of immobilized enzymes. In fact, an enzyme-carrier complex has to be regarded as a heterogeneous preparation in which the individual enzyme molecules may differ from one another by (i) a different number of covalent or noncovalent bonds with the carrier, (ii) different amino acids involved in the enzyme-carrier linkage, and (iii) a different geometry of the matrix at the binding sites. 26z Although stabilities of the individual enzyme molecules should differ, it has been claimed that a great number of immobilized enzyme systems can be characterized by a relatively simple mathematical expression for the residual enzyme activity (the sum of two decaying exponential terms)f162 Malcata et a l . 226-228 have tested three nested models for their ability to describe the change in the activity of a lipase from Aspergillus niger immobilized by adsorption on microporous polypropylene when this lipase is employed to effect the hydrolysis of butteroil: (i) the native lipase is converted to an inactive form; (ii) the native lipase is converted to an inactive form in one of two parallel processes, the other of which leads to another active stable form; and (iii) the native lipase is altered to another active form and both these forms are susceptible to deactivation. Statistical analysis of the rate data indicated that the second model provided the best fit. Similar mathematical expressions for resid-

Immobilized lipase kinetics: F. X. Malcata et al. ual activities were used by Toscano et a l . TM to describe the deactivation of an immobilized hydrolase in the presence of an organic solvent.

Effect of pH on the rates of reactions catalysed by lipases and on the rate of deactivation of lipases Due to the proteinaceous nature of enzymes, lipases may contain basic, neutral, and acidic groups. Consequently, at any given pH the intact lipase may contain both positively and negatively charged groups. Such ionizable groups often constitute part of the active site and are frequently involved in general acid-base catalysis. 173 However, a catalytically active lipase tends to exist in one particular ionization state. Usually, the catalytic activity of the lipase changes with pH in a bell-shaped fashion, thus yielding a maximum rate in the stability range. Shifts in the optimum pH after immobilization for various lipases are tabulated in Table 3. The maxima in the rates of reaction catalysed by immobilized lipases are observed at pH values between 4.0 and 10.0. With very few exceptions, the pH optima for the immobilized lipases are equal to or higher than those for their free counterparts. Hence, the immobilization procedure seems to render catalytically important amino acid residues more basic. An explanation consistent with these results and with the experimental evidence is that upon immobilization the active site becomes more exposed to solvent than it was in the globular, folded soluble lipase form. Hence, proton transfer to the amino acid residues at the active site becomes less hindered. This simplistic view does not lead to rate expressions that are able to accurately fit the experimental data for some systems. For example, Malcata et al. 229 have reported that the rate expression for glyceride hydrolysis in the presence of a lipase from Aspergillus niger immobilized by adsorption on polypropylene is of a form such that two local maxima are observed for the dependence of the rate on pH. Similar behavior was observed for the pH dependence of the activity of a lipase from Rhizopus for the hydrolysis of olive oil at 50oc .267 The pH also affects the stability of enzymes, although this effect is seldom taken into account in the development of mathematical models of the kinetics of enzyme deactivation. For pH values well removed from the optimum pH, enzyme deactivation is usually very rapid. 219 For most proteins minima have been observed in the rate of deactivation. 173In fact, if different enzyme forms arising from different ionization states deactivate at different rates, the overall rate of deactivation will depend on any reaction parameter which influences the relative proportions of the different enzyme forms. 26sImmobilization of a lipase from Candida cylindracea by adsorption on silica gel 249 and immobilization of a lipase from Mucor javanicus on DEAEcellulose by ion exchange 196led to significant increases in their respective stabilities.

Effect of temperature on the rates of reactions catalysed by lipases and on the rate of deactivation of lipases At low temperatures the rate of a lipase-catalysed reaction increases with temperature according to the Arrhenius model (with a low activation energy). However, at elevated temperatures this model breaks down due to extensive irreversible denaturation of the lipase) 73 Values of activation energies reported for the low temperature regimes range from 0.97 to 74.9 kJ mol-1 192.193.196.230,234.245,263These values are typical of enzymatic reactions. 2~9 Very low activation energies 169'193 may indicate that the lipase-catalysed process is controlled by diffusion of substrates. Values of the optimum temperatures for lipase action are tabulated in Table 4. These optima lie between 30 and 62°C. Immobilization almost always leads to positive shifts in the optimum temperature. This result implies that upon immobilization the lipase becomes more resistant to thermal deactivation. The effects of temperature on both the rate of a lipase-catalysed reaction and the rate of deactivation of the lipase must be handled via complex models. Malcata et al. 228 have developed such a model for the hydrolysis of butteroil in the presence of a lipase from Aspergillus niger immobilized by adsorption on polypropylene.

Effects of different substances on the rates of reactions catalysed by immobilized iipases At a molecular level, the mechanism of interesterification reactions involves hydrolysis of the ester molecule followed by an esterification reaction. As a result, in addition to the lipase, water must be present in at least catalytic amounts. 247,269 On the other hand, since the processes in question are reversible, the presence of very large amounts of water promotes the hydrolysis reaction. Hence, one very important operational consideration when one utilizes an immobilized lipase to catalyse ester synthesis and interesterification reactions is the concentration of water necessary to obtain maximum activity. Values reported in the literature are compiled in Table 5. Also included are values for hydrolysis reactions performed in nonaqueous solvents. It is apparent that the optimum water concentration lies in the range from 0.04 to 11% (w/v). These values are comparable in magnitude to the maximum solubility of water in commonly employed feedstocks or solvents. Experiments involving the synthesis of n-butyl oleate from oleic acid and n-butanol catalysed by a lipase from Mucor miehei immobilized on an anion exchange resin TM indicate that the optimal water concentration depends on the operating temperature. Although differences between concentrations and activities are almost always neglected in the literature, it should be emphasized that the hydrolysis equilibrium in the organic phase should be expressed in terms of the activities of the esters, alcohols, fatty acids, and water instead of their molar concentrations.132 The ac-

Enzyme Microb. Technol., 1992, vol. 14, June

441

Review tivity coefficients are strongly dependent on the water content of the organic phase, which in turn is a direct function of the extent of reaction. Tables 6 and 7 summarize the effects of different salts and solvents, respectively, on the activities of immobilized lipases. The effects of these substances on the activities of immobilized lipases should in principle be similar to those observed for free lipases. Inspection of Table 6 indicates that ammonium sulfate and sodium chloride have essentially no effect. Potassium cations have a very small detrimental effect, whereas mercuric ions lead to almost complete inactivation of the lipase in question. Magnesium cations, on the other hand, are strong lipase activators. The ranges of fractional activities of lipases in different solvents are strongly dependent on the standard solvent selected (cf. Table 7); however, solvents such as acetone or dimethylformamide (or, to a less extent, dioxane) should not be employed in a reaction system because they tend to completely inactivate the lipase. On the other hand, diethyl ether and trichloroethane appear to be strong activators. Studies of the effects of surfactants on lipase activity indicate that polyvinyl alcohol TM and bile salts ~96 enhance the catalytic action of lipases from R h i z o p u s o r y z a e and M u c o r j a v a n i c u s bound to alumina or DEAE-cellulose, respectively. By contrast, the detergent Triton 192tends to inactivate a lipase from C a n d i d a r u g o s a entrapped in polyethylene glycol.

products that the oleochemicals industry can produce efficiently using lipases are expanding rapidly. 285 The use of immobilized lipase technology for the production of high value-added final products offers enormous potential for future development. However, the solution of the many engineering problems associated with the use oflipase-catalysed reactions 83'286must first be addressed via generation of the information necessary to understand the fundamental chemical and transport phenomena involved so as to permit process simulation and optimization work to be carried out. We hope that the present review will be a useful step towards this goal.

Acknowledgements Financial support for F. X. Malcata has been provided through funds administered by Portugal's National Board for Scientific and Technological Research (INVOTAN Fellowship), the Institute of Food Technologists (IFT Graduate Fellowship and John V. Luck/ General Mills Fellowship), and the Center for Dairy Research of the University of Wisconsin. Financial support for H. R. Reyes has been provided by AT&T. Financial support for H. S. Garcia has been provided by CONACYT (Mexico) and by the Center for Dairy Research. Overall support of the project has been provided through a grant to the Center for Dairy Research funded by the National Dairy Promotion and Research Board.

Final remarks There is increasing interest in the commercial potential of lipolysed products which can be employed in a number of possible applications. For example, concentrated flavors consisting of esters containing shortchain acid residues 1°7'197 can be produced in this manner. Products with enhanced appeal in certain dietary foods can involve the synthesis of fatty acid acyl esters of carbohydrates 272 and the synthesis of medium chain triglycerides from plant oils or animal fats. 273 Concentrates of polyunsaturated to-3 fatty acids obtained by hydrolysis offish oils,274 concentrates ofy-linolenic and linoleic acids 275'276 or gadoleic, erucic, and nervonic acids 277formed by hydrolysis of seed oils, and concentrates of fl-adrenergic drugs prepared by lipase-catalysed enantioselective hydrolysis278represent products with potential nutropharmaceutical value. Emulsifiers such as monoacylglycerols formed by partial hydrolysis of a variety of o i l s , 279 c o c o a butter substitutes produced by interesterification of cheaper feedstocks, 28°'2s~ waxes synthesized from fatty acids and long-chain monoalcohols, 282 and fatty amides synthesized from alkylamines and triglycerides283 are representative of materials characterized by enhanced functional properties. Most of these products cannot be conveniently prepared using conventional chemical synthesis routes. Hence the use of lipase-mediated reactions is an appropriate approach to the production of these materials. TM The types of high value-added

442

Enzyme Microb. Technol., 1992, vol. 14, June

References 1

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52

53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72

73 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88

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