Çörekotu (nigella Sativa L) Lipaz Enziminin Stabilizasyonuna Şekerlerin Etkisi

Abstract

Son yularda lipaz enzimlerinin, deterjan ve deri sanayiinde yağların biyolojik hidrolizinde; yağ sanayiinde istenilen özelliklerde sıvı yağ ve margarin üretimi için yağların modifikasyonunda ve kimyasal yolla başarılamayan çeşitli organik sentezlerin gerçekleştirilmesinde kullanılabilir olmasının ortaya çıkması, endüstriyel lipaz enzimlerinin üretimi ve stabilizasyonu üzerine çalışmaların hızlanmasına neden olmuştur. Literatürde, mikrobiyal lipazların stabilitesi üzerinde çok sayıda çalışma olmasına karşılık, tahıllar ve tohumlar daha ucuz lipaz kaynakları olmalarına rağmen, bitkisel kökenli bu lipazların stabilizasyonu üzerinde çok az sayıda çalışmaya rastlanmıştır. Bu çalışmada, çörekotu lipaz enziminin şeker bileşikleri ile stabilizasyonu amaçlanmıştır. Bu amaçla, çörekotu lipazına 50-1500 mM konsantrasyonlarında sakkaroz ve glukoz katılmış ve şeker konsantrasyonuna bağlı olarak 30-60°C'larda lipazın termostabilitesinin değişimi incelenmiştir. Çörekotu lipaz enzimine en yüksek stabilite kazandıran optimum sakkaroz ve glukoz konsantrasyonu, sırasıyla 300 ve 500 mM olarak saptanmıştır. Optimum konsantrasyonlarda sakkaroz ve glukoz ilave edilmiş lipaz enzimlerinin incelenen her sıcaklıkta inaktivasyon hız sabitleri küçülmüş ve yanlanma ömürleri uzamıştır. 60°C'da yanlanma ömrü 8.2 dakika olan doğal enzim, sakkaroz katkısıyla 6.2 kat, glukoz katkısıyla ise 3.8 kat daha uzun yarılanma ömrüne sahip olmuştur. Doğal enzimin inaktivasyon ve hidroliz aktivasyon enerjileri 120.1 ve 24.1 kJ/mol iken. sakkaroz ve glukoz katkılı enzimlerde bu değerler sırasıyla 84.8 ve 21.1 kJ/mol: 97.7 ve 19.2 kJ/mol değerlerine düşmüştür. Doğal enzimin Km ve Vmax değerlerinde de ortamdaki şeker etkisiyle küçük mertebelerde değişim gözlenmiştir. Çalışmanın sonucunda, glukoz ve sakkaroz katkısıyla çörekotu lipaz enzimine, çözelti ortamında, karakteristik değerlerininde çok az değişimle stabilite kazandırabileceği ortaya çıkarılmıştır. v

Lipases are one of the most important class of industrial enzymes, which are used intensively in the biotransformation of fats and oils, in household detergents, and especially in the synthesis of very important pharmaceuticals and agrochemicals. To date, most of the industrial lipase enzymes have been produced from microbial resources and they are generally stabilized by modification geneticelly or immobilization against heat denaturation. Cereal grains and oilseeds are cheap and alternative sources of lipases, however they have not yet been stabilized and used as industrial purposes. Most of the enzymes become unstable and rapidly inactivate (or denature) when they are isolated from their natural environment in vivo. In living organism nascent polypeptide chains of enzymes emerging from ribosomes fold spontaneously to acquire the conformation of native, biologically active proteins. Although very little is known about how this happens, available evidence indicates that two large and opposing forces drive the transition from the primary structure to the tertiary structure. The first one, hydrophobicity, stems from the propensity of nonpolar residues to remain buried in the protein core in order to avoid contact with water. The second one arises from the impossibility of two polypeptide segments simultaneously occupying the same volume. This perfectly folded, fully functional protein can lose its biological activity in vitro by unfolding of its tertiary structure to a disordered polypeptide in which key residues are no longer aligned closely enough for continued participation in functional interactions. Such unfolding is termed denaturation. Enviromental conditions, such as pressure, pH. ionic strength, mechanical forces, the presence of a variety of denaturing agents and particularly temperature, effect on the extent of denaturation. Denaturation is usually cooperative and may be reversible if the denaturing influence is removed. Enzyme proteins are also subject to chemical changes leading to an irreversible loss of activity or inactivation, particularly following unfolding. These chemical changes may be listed as subunits dissociation, proteolysis, loss of essential cofactor, aggregation, incorrect refolding, deamidation of asperagine residues, rupture of peptide bonds on the C-terminal side of aspertic acid residus and destruction of disulfide bonds. The stabilization of enzymes against inactivation is of great interest either from a theoretical point of view, i.e., a better understanding of the mechanisms of protein inactivation, and from a practical point of view when enzymes are to be employed in industrial processes. In fact, the large-scale use of a thermostable biocatalyst, rather than a thermosensitive one, has several advantages such as: vm 1. The possibility to operate at high temperature, which affords high temperature rates, high solubility of substrates, a lower viscosity of reaction medium and reduced risk of microbial contemination; 2. The increase the storage and operational stabilities; and 3. The eventual increase of the resistance to protein denaturating agents. Methods for minimizing inactivation can be divided into four main categories: synthesis of more stable enzymes by genetic engineering, chemical modifications, immobilization, and use of water-miscible stabilizing additives. In many instances the latter represents the simplest and most effective approach to hindering enzyme inactivation. Though selection of additives depends on the nature of the enzyme, increases in the thermostabilities of the enzymes are frequently observed by the addition of sugar or polyol. Recently, the stabilizing effect of adding polyol and sugar was quantitatively analyzed by a series deactivation model for glucose oxidase, a-amylase, papain, a-chymotrypsin, and penicillin G acylase. However, the stabilization of lipases, particularly plant lipases have not been throughly investigated in this respect. In recent studies conducted at Chemical Technologies in the Chemical Engineering Department, Nigella sativa (Black cumin) seeds have been shown a good source of lipase enzyme which can be used as effective biocatalyst in hydrolysis and esterification reactions of triglycerides. In this study, the thermostability of black cumin lipase was investigated in the presence of sugar compounds (sucrose and glucose). To select the optimum concentration of each sugar that provides the highest thermostabilization of enzyme, the effects of then- different concentrations on the stability of Nigella sativa lipase were examined. Samples of Denizli origin black cumin seeds were purchased from a local market in Istanbul. Chemical composition of the seeds was determined according to the standart AOCS methods. Moisture content was found by oven-drying at 105°C. Total nitrogen content was determined using the standard Kjeldahl method. Crude protein was expressed as 6.25xN. Crude fiber was determined by calculating the loss in weight of dried residue remaining after digestion of a fat-free sample with 0.25 M H2SO4 and 0.6 M NaOH under specified conditions. Ash content was determined by incineration of the sample in a muffle furnace at 600°C for 16 h. Total fat content was obtained by the soxhlet extraction method using hexane as described by AOCS method. Carbohydrate content was obtained by subtracting the stun of protein, fat, ash and moisture from 100. The proximate composition of Nigella sativa seeds indicated that seeds were composed of 23.3% protein, 35.4% oil, 9.4% moisture, 4.0% ash, 6.7% crude fiber and the rest being composed of other carbohydrates. The lipase extracts were obtained as follows: 10 gram seeds were first soaked in water IX overnight at 25°C, then were homogenized for 20 min using a blender with 200 mL phosphate buffer solution (pH 6) at 4°C. The homogenate was filtered through nylon cloth. The homogenate was then centrifuged for 30 min at 10.000 g, yielding supernatant liquid and crude particulate fraction. The supernatant liquid is called as "lipase extract" throughout this text. The protein content of lipase extract was measured spectrophotometrically according to the Biuret method using a Randox Total Protein Reagent. Bovine serum albumin was used as the standard. The hydrolytic activity of the Nigella sativa lipase was assayed using 58 mM olive oil emulsion. For preparation of this emulsion, 5 g olive oil, 5 g gum arabic and 95 mL 0.89% (w/v) NaCl solution were emulsified with a blender at room temperature two times for 5 minutes. Emulsions were always prepared immediately before use. The assay mixture contained 5 mL olive oil emulsion, 0.5 mL 0.1 M CaCb solution, 3 mL phosphate buffer (pH 6) was incubated at 37°C for 15 min. 1 mL of lipase extract (containing 5- 10 mg of protein) was then added, and the reaction proceeded at 37°C in a shaker for 15 min. The reaction was terminated by adding 20 mL of acetone: ethanol (1:1, v/v). The liberated free fatty acid was measured by titration with 0.05 N NaOH using thymolphthalein as an indicator. Corrections were made for endogenous fatty acid production (assay mixture without olive oil emulsion) and nonen2ymatic fatty acid production (assay mixture without enzyme extract). One unit of specific activity is equivalent to 1 umol of free acid liberated/min. mg protein at 37°C. To investigate the effect of sugar compounds on the stability of Nigella sativa lipase, aqueous solutions of the sugar (sucrose and glucose at various concentrations) were added to the lipase solution. The mixtures were incubated at 30, 40, 50 and 60°C for 30 min. The samples were cooled in ice immediately after heat treatment. Later, residual activity was measured at 37°C and pH 6 according to the above mentioned activity determination method. Thermostability was expressed as percent residual activity of sugar containing lipase enzyme after heat treatment at temperatures in range of 30-60°C. To select the optimal concentration of sucrose and glucose that provides the highest thermostabilization of enzyme at all temperatures, the variations of the residual activity values with the concentration of sugar compounds at 30, 40, 50 and 60°C were plotted in same figure. From the examination of the curves, the optimum concentrations of sucrose and glucose were found to be 300 and 500 mM, respectively. The inactivation kinetics of the native and sugar added (at optimum concentrations) lipase enzymes were studied in the 30-60°C temperature interval. Inactivation rate constants (ki) of Nigella sativa lipase samples were estimated according to the first order inactivation kinetics. A mixture of 1 mL lipase extract and 3 mL phosphate buffer (pH 6) was incubated for different time intervals at various temperatures. The heat-treated enzyme solutions were rapidly cooled. Thereafter, 5 mL olive oil emulsion (58 mM) and 0.5 mL 0.1 M CaCh solution were added and residual lipase activity was immediately assayed as described previously. Inactivation rate constants (ki) at different temperatures were calculated from the slope of the linear equation of first order inactivation kinetics Ln [At/Ao]= -kj.t [At: Activity of enzyme after heat treatment; Ao: Activity of enzyme before heat treatment; t: time (min)]. The level of thermostabilization of sucrose and glucose added lipase enzymes was expressed as a stabilization factor (SF) and calculated using the equation: SF=- half - life of native enzyme half - life of sugar added enzyme The ki, to.5 (half-life) and SF values of sugar-containing lipase together with ki and to.: values of native enzyme are given in the following table. Inactivation rate constants obtained by sugar containing enzyme preparations were always lower than those of the native enzyme. On the other hand the half-life values for inactivation of sugar added enzymes were significantly higher than the values obtained for native enzyme at all temperatures. The greatest enhancement of thermostability was observed with sucrose, as a nearly 6-fold increase at 60°C. The apparent activation energy for inactivation of native and sugar added Nigella sativa lipase was calculated from the slope of the linearized Arrhenius equation: Ln ki= LnA-(Ei/RT) when it is plotted as Ln ki versus 1/T [ki= inactivation rate constant (min"1); Ei= activation energy of inactivation (J/mol); R= gas constant (8.3 14 J/mol.K): T= absolute temperature (K)]. Activation energies for inactivation were calculated as 120.1 kJ/mol for nativa lipase, and 84.8 and 97.7 kJ/mol for sucrose and glucose containing enzyme preparations, respectively. To see the effect of sugar compsunds on the kinetic parameters (Km and Vmax) of Nigella sativa lipase, olive oil was used as substrate and the hydrolysis reactions of olive oil with all enzyme samples investigated were carried out at 37°C in phosphate buffer (pH 6), changing the oil concentration from 19 to 76 mM. The initial rates were determined by measuring the amount of liberated free fatty acids. The variation of the initial rates with substrate concentration was plotted by using Lineaweaver-Burk equation for native and sugar containing lipase enzymes. Km ve Vmax values of native and sugar added enzyme preparations were estimated from the slopes of the lines. The Km values were found to be 40.8, 28.6 and 33.9 mM for the native, sucrose added and glucose added lipase preparations, respectively. The Vmax values were established as 39.2, 26.7 and 32.3 umol/min for the native, sucrose added and glucose added Nigella XI sativa lipases, respectively. In the presence of sugar compounds the reactions follow Michaelis-Menten kinetics with uncompetitive inhibition. Finally, the effect of sucrose and glucose on the optimal temperature and activation energy of Nigella sativa lipase was investigated. The hydrolysis reactions of olive oil were conducted with native and sugar added enzymes at 30, 35, 40, 45 and 50°C. The initial rates were determined by measuring the amount of liberated free fatty acids. The addition of sugar compounds did not cause any change on the optimal temperature of native Nigella sativa lipase. The optimum temperature for olive oil hydrolysis in all cases was 40°C. However, slight decreases were observed in the activation energies of sugar containing lipases preparations. The activation energies of native and sucrose and glucose added enzyme preparations were calculated from the Arrhenius plot. Estimated values in kJ/mol, were 24.1, 21.1 and 19.2 for native, sucrose and glucose added lipases, respectively. As conclusion, sucrose and glucose compounds can be used for stabilization of Nigella sativa lipase. Materials used for the stabilization are easily available and safe for applications. xn