Ind. Eng. Chem. Res. 2008, 47, 9651–9657
9651
Response Surface Methodology for Optimization of Lipase Production by an Immobilized Newly Isolated Penicillium sp. Elisangela Wolski,† Elisangele Menusi,† Marcio Mazutti,‡ Geciane Toniazzo,† Elisandra Rigo,§ Roge´rio Luiz Cansian,† Altemir Mossi,† J. Vladimir Oliveira,† Marco Di Luccio,† De´bora de Oliveira,† and Helen Treichel*,† Programa de Mestrado em Engenharia de Alimentos, UniVersidade Regional Integrada do Alto Uruguai e das Misso˜es, Campus de Erechim, AV. Sete de Setembro, 1621, 99700-000, Erechim, RS, Brazil, Departamento de Engenharia de Alimentos, UniVersidade Estadual de Campinas, UNICAMP, SP, Brazil, and Departamento de Engenharia Quı´mica e de Alimentos, UniVersidade Federal de Santa Catarina, UFSC, SC, Brazil
This work reports the use of response surface methodology to optimize the lipase production by submerged fermentation using immobilized cells of a newly isolated Penicillium sp. For this, experimental designs were employed to evaluate the best condition for microorganism immobilization. The optimum experimental conditions for immobilized microorganisms were determined as 2.5% (w/v) of calcium alginate, 0.2% (w/v) of CaCl2, and 1.5% (v/v) of glutaraldehyde and 6 h of curing time, yielding a lipase activity as high as 20.96 U/mL in 120 h of fermentation. The activity obtained is higher than those observed with free cells of this microorganism and comparable to results presented in the literature for lipase production with immobilized cells from several microorganisms. The crude enzymatic extract presented optimum pH and temperature of 7.0 and 37 °C, respectively, and maintained its activity for 60 days storage at -10 °C. Introduction Lipases are enzymes belonging to the group of the hydrolases, whose main biological function is to work as the catalyst of the hydrolysis of insoluble triacylglycerols to generate free fatty acids, mono- and diacylglycerols, and glycerol. Besides its natural function, lipases can catalyze esterification, interesterification, and transesterification reactions in nonaqueous media.1,2 Microbial lipases are biocatalysts that have interesting characteristics, such as action under mild conditions, stability in organic solvents, high substrate specificity, and regio- and enantioselectivity.3-5 The potential for industrial applications of lipases comprises the industry of additives (modification of aromas), fine chemistry (esters synthesis), detergents (hydrolysis of fats), wastewater treatment (decomposition and removal of oleaginous substances), leather (fat removal from animal skin), and pharmaceutical and medicinal areas (medicines, digestive, and enzymes for diagnosis).6,7 However, the high costs of production of these biocatalysts often restrict their use.3 Microorganisms able to produce lipases can be found in several habitats, including wastes of vegetable oil and dairy product industries and soils contaminated by oils, seeds, and deteriorated food.8 The soil has a great variety of microorganisms that can be isolated and evaluated for their potential as enzyme producers.9 The isolation and screening of microorganisms can lead to the production of lipases with desirable properties (stability to high temperatures and wide pH ranges, high specificity to certain fatty acids, and enatioselectivity). Furthermore, it can open promising scientific and commercial perspectives.10 Several methods can be used for microorganism screening based on the determination of the presence of extracellular * To whom correspondence should be addressed. Tel: +55 54 5209000. Fax: +55 54 5209090. E-mail:
[email protected]. † Universidade Regional Integrada do Alto Uruguai e das Misso˜es. ‡ Universidade Estadual de Campinas. § Universidade Federal de Santa Catarina.
lipases. The use of a solid medium with inducer substrates such as vegetable oils, standard triglycerides (tributirin, triolein), Tween 80, and dyes has been widely described in the literature.9,11,12 However, some of these substrates may not be adequate for lipase detection, turning of extreme importance the verification of the specificity of the analysis for lipases and esterases.13 Both enzymes catalyze the hydrolysis and synthesis of the ester bound. The esterases hydrolyze esters with carboxylic acids of short chain, while lipases prefer long chain fatty acids.14 Thus, investigations on the use of different microorganisms, supplements and substrates for lipase production can contribute in finding ideal combinations to obtain high-value lipases, using substrate and operational conditions that help developing largescale cost-effective systems.15-17 In this context, enzyme and microorganism immobilization has been of great interest recently. The immobilization of whole cells, in addition to make easy extraction and purification steps of desirable enzyme, may also provide higher yields of enzyme activity, higher operational stability, and low and efficient production costs.18,19 Immobilization systems are employed to enzymes, microorganisms, and plant and animal cells, aiming at involving the biocatalysts in a proper matter where they can retain their activity and may be used repeatedly for a long period of time.20 Compared to free cells, the microorganism immobilization presents many advantages, including high growth rate, ease of operation in continuous mode and process scale-up, easy purification of the products, and good catalytic stability, as well as better tolerance to high concentration of toxic compounds.19,21 For cell immobilization, natural polymers such as alginate, carrageen, agar, and chitosan and synthetic ones like polyacrylamide, polyacrilate, and polyurethane have been widely used as encapsulating agents.19 Sodium alginate is one of the most used hydrocolloids for encapsulation. The main advantage in alginate gels is their thermostability, and thus they can be stored in room temperature.22 In spite of all these aspects, the literature is scarce regarding microbial cell immobilization for lipase production.23 For
10.1021/ie800658j CCC: $40.75 2008 American Chemical Society Published on Web 10/29/2008
9652 Ind. Eng. Chem. Res., Vol. 47, No. 23, 2008
example, Ellaiah et al.24 investigated the lipase production by immobilized cells of Aspergillus niger in different encapsulating agents, such as alginate, k-carragen, and polyacrilamide. The maximum lipase production was obtained with the use of free microorganism (4.29 U/mL) and with cells immobilized in 3% (w/v) of alginate. Elibol and Ozer23 evaluated the immobilization of Rhizopus arrhizus cells in polyurethane for lipase production and obtained a production compared to the use of free cells. Hemachander et al.18 evaluated the immobilization of Ralstonia pickettii cells for lipase production in different solid matrices (agar, alginate, and polyacrilamide). The best results were obtained when using 20% (w/v) of polyacrilamide, 24.75 U/mL. Benjamin and Pandey25 used immobilized cells of Candida rugosa for lipase production in a packed bed reactor, and after a series of experiments, the optimum lipase activity achieved was 17.94 U/mL. Yang et al.26 studied the lipase production by immobilized mycelium of Rhizopuz arrhizus in polyurethane through submerged fermentation. The authors obtained lipase productivity of 17.6 U/(mL h) in repeated groups of fermentation. Usually the method for determining optimal conditions in fermentation processes is varying one parameter while keeping others at a constant level. This is a time and cost ineffective method that presents also the disadvantage of not including the interaction effects among variables. Optimization using factorial design and response surface methodology can overcome such drawbacks. A factorial design technique has been successfully used to optimize and evaluate the effect of process parameters in the production of enzymes and other metabolites.27,28 On the basis of these aspects, the main objective of this work was to report the use a sequential strategy of experimental design for the optimization of lipase production by submerged fermentation using an immobilized newly isolated Penicillium sp. as the microorganism. Partial characterization of the crude enzymatic extract from fermented media was also carried out to determine some properties of the obtained lipases. Materials and Methods Cell Production. Following a standard procedure for isolation, several microorganisms were isolated from different sources as olive oil, cheese, tomato extract, soybean oil, milk cream, meat, soybean bran, and contaminated culture media (data not shown). The most promising strain, isolated from soybean bran, was preliminarily identified as Penicillium sp. and used in this work. This microorganism was maintained in glycerol and potato dextrose agar slants under refrigeration. The propagation of spores prior to fermentation was carried out for 7 days at 27 °C in a medium constituted by potato dextrose agar (PDA) 3.9% (w/v) and distilled water.15 Microorganism Immobilization. Immobilization of free Penicillium sp. cells was preliminarily performed using different entrapment materials with the best results obtained using calcium alginate as carrier. With the aim of optimizing lipase production, this gelificant agent was used in further studies. Spores were immobilized in calcium alginate by a traditional external gelation method.29 About 25 mL of sodium alginate and 10 mL of spore suspension (2 × 106 spores/mL) were mixed well, and this slurry was added dropwise to CaCl2 solution at room temperature. The beads (∼4 mm) formed were then cured in a refrigerator at 5 °C for maturation. Afterward, the beads were filtrated, washed two to three times with sterile distilled water and kept in a glutaraldehyde solution for 15 min, then washed again with distilled water, and added to the fermentation medium. The effects of sodium alginate, glutaraldehyde and calcium chloride
Table 1. Ranges of the Factors Investigated in the Four Experimental Designs -1.41
Level
-1
0
+1
+1.41
First Experimental Design sodium alginate (%, w/v) CaCl2 (%, w/v) glutaraldehyde (%, v/v) curing time (h)
2 0.2 0 6
3 1.1 1 15
4 2 2 24
Second Experimental Design sodium alginate (%, w/v) CaCl2 (%, w/v) glutaraldehyde (%, v/v) curing time (h)
2 0.2 0.5 1
2.5 0.6 1 6
3 0.8 1.5 11
Third Experimental Design glutaraldehyde (%, v/v) curing time (h)
0.8 1.7
1 3
1.5 6
2 9
2.2 10.7
2.6 10.1
3 11.75
Fourth Experimental Design glutaraldehyde (%, v/v) curing time (h)
0 1.25
0.4 1.9
1.5 6
concentrations and curing time were evaluated by sequential strategy of experimental designs. Culture Conditions. Fermentation media consisted of peptone 2% (w/v), NaCl 0.5% (w/v), yeast extract 0.5% (w/v), and olive oil 1% (w/v). The temperature was fixed at 28 °C. These culture conditions were optimized earlier and defined as the control experiment, using the free microorganism. Cultivation was carried out in 250 mL Erlenmeyer flasks covered with cotton plugs. The medium (100 mL) was sterilized at 121 °C for 15 min, cooledm and inoculated with a suspension of spores to a final count of 2 × 106 spores/mL. Samples were collected and filtered through Whatmann qualitative paper. The filtrate was considered as crude enzyme and was used for analytical assays. Lipase Activity. Lipase activity was assayed by reaction using olive oil as the substrate followed by alkali titration. Olive oil (10% (w/v)) was emulsified with arabic gum (5% (w/v)) in sodium phosphate buffer 100 mM, pH 7.0. A 2 mL sample of crude enzyme was added to 18 mL of this emulsion. After incubation in a shaker for 15 min at 37 °C and 150 rpm, the reaction was interrupted and the fatty acids were extracted by the addition of 20 mL of an acetone/ethanol solution (1:1 v/v). The amount of fatty acids liberated was then titrated with 0.05 M NaOH until pH 11. Reaction blanks were run in the same way, but the sample was added after addition of acetone/ethanol solution. Lipase activity measurements were carried out in duplicate. A unit of lipase activity was defined as the amount of enzyme that yields 1 µmol of fatty acids per minute in the assay conditions.30 Optimization of Lipase Production by Immobilized Microorganism. Table 1 presents the variables and range of study used in the sequential strategy of experimental designs for optimization of lipase production by immobilized by Penicillium sp. In the first and second experimental designs (full 24), the effect of sodium alginate, glutaraldehyde, and calcium chloride concentrations and curing time were evaluated. Temperature and agitation were fixed at 28 °C and 180 rpm, respectively, on the basis of previous tests conducted for free microorganism. After statistical analysis, a third experimental design (full 22) was performed keeping constant the sodium alginate and calcium chloride concentrations. With the objective of optimizing the lipase production, a fourth, full 22 experimental design was conceived, making use of the same variables evaluated in the third experimental design.
Ind. Eng. Chem. Res., Vol. 47, No. 23, 2008 9653 2
Table 2. Variables and Levels Studied in the Full 2 Experimental Design for Evaluation of Optimum Values of Temperature and pH
a
level
temperature (°C)
pH
-1.41 -1 0a +1 +1.41
30 32 37 42 44
4.9 5.5 7.0 8.5 9.1
Central point.
Partial Characterization of Crude Enzymatic Extract. To determine the optimum values of temperature and pH in terms of lipase activity, a full 22 experimental design was accomplished using the crude enzymatic extract obtained by immobilized cells of Penicillium sp. Sodium phosphate buffer at different pH values was added to the emulsion for measurement of lipase activity, and the samples were incubated in a shaker at different temperatures for 15 min and 150 rpm. The ranges of pH and temperature studied is presented in Table 2. All the experiments were executed in duplicate. The stability temperature of the crude enzymatic extract was determined by incubation at a fixed pH (7.0) and different temperatures: 25, 35, 45, 55, and 65 °C. The stability pH was determined by incubation of the extract at 50 °C at the following pH (sodium phosphate buffer 100 mM): 4.8, 5.5, 6.5, 7.5, and 8.5. All runs were carried out in duplicate, and samples were withdrawn at regular time intervals. The stability of the crude enzymatic extract at low temperature was evaluated by storing the extract at 4 and -10 °C and determining the enzyme activity periodically. Results and Discussion First Experimental Design. Table 3 presents the matrix of the 24 full experimental design performed in the first step of this work with real and coded values for independent variables (sodium alginate, calcium chloride, and glutaraldehyde concentrations and curing time) and the response in terms of lipase activity during the fermentation with the immobilized microorganism. The last line of this table presents the lipase activity obtained with the free microorganism at previously optimized experimental conditions. This point was named as the control experiment and was performed so as to allow comparison with the results obtained with immobilized Penicillium sp. From Table 3 it can be observed that the highest lipase activity was obtained at 96 h of fermentation (12.81 U/mL) using 4% (w/v) of sodium alginate. It is worth mentioning that capsules obtained with calcium alginate 2% (w/v) presented lipase activities similar to that obtained with 4% (w/t) of this component. However, after 48 h of fermentation, the granules dissolved in the production medium, not an interesting fact when using immobilized cells. The lipase production presented good results at curing time of 6 h (5 °C, in CaCl2 solution), time possibly enough to give resistance to granule capsules. The data obtained in the first experimental design were statistically treated in 48, 72, and 96 h of fermentation. In 48 h of fermentation one can observe that, within a confidence level of 95%, the CaCl2 and calcium alginate concentrations presented a negative effect on lipase production. The interaction between glutaraldehyde concentration and curing time also presented a negative effect. For 72 h of fermentation it could be observed that all variables presented a significant effect (p < 0.05) on lipase production. The glutaraldehyde concentration presented a positive effect. The same behavior was observed at 96 h of fermentation. At
this step, it may be opportune to emphasize that, besides good lipase activities, one may require an experimental condition that results in morphologically perfect granules and suitable diffusivities that can allow a good interaction of nutrients in the fermentation medium with microbial cell. Second Experimental Design. On the basis of the results obtained in the first experimental design, a second full experimental design was performed. Table 4 presents the matrix of the 24 full experimental design accomplished in this step with real and coded values for the same independent variables evaluated previously and the response in terms of lipase activity. From this table one can observe that higher lipase activities were achieved at 96 h of fermentation, with activity values around 13.18 U/mL in the central point. When using a calcium alginate concentration of 3% (w/v) (run 4), a lipase activity of 13.08 U/mL was obtained. Results obtained here are, for example, higher than those obtained by Ellaiah et al.24 (4.29 U/mL) using immobilized Aspergillus niger as the microorganism. The statistical analysis of experimental data of the second experimental design revealed that CaCl2 concentration presented a significant negative effect (p < 0.05) on lipase production while the glutaraldehyde concentration presented a positive one (p < 0.05). Third Experimental Design. Table 5 presents the matrix of the full 22 experimental design of 11 experiments, where the levels of sodium alginate and CaCl2 were fixed at 2.5% (w/v) and 0.2% (w/v), respectively. The ranges of investigation for glutaraldehyde concentration and curing time were established as 0.8 and 2.2% (v/v) and 1.7 and 10.2 h, respectively. After 120 h of fermentation, the highest lipase activity was obtained at the central point (19.82 U/mL) at 6 h of curing time and 1.5% (v/v) of glutaraldehyde concentration. Related to the first experimental design it is possible to observe a considerable improvement on lipase activity, from 12.81 U/mL to 20.09 U/mL, which means an increase of nearly 100%. The statistical analysis of the data obtained showed that all evaluated variables presented a positive effect (p < 0.05) on lipase production. Fourth Experimental Design. After analyzing the results obtained in the third experimental design, the next step toward optimizing lipase production was to perform a fourth 22 full experimental design. Table 6 presents the matrix of the experimental design with the immobilized microorganism and the response in terms of lipase activities. The glutaraldehyde concentration and curing time were evaluated from 0 to 3% (v/v) and 1.25 to 11.75 h, respectively. It can be observed that the highest enzymatic activity (20.96 U/mL) was achieved at 1.5% (v/v) of glutaraldehyde and curing time of 6 h. In this experimental condition the granules kept their morphology until the end of the fermentation. Ellaiah et al.24 evaluated the lipase production by immobilized cell of Aspergillus niger in three different supports. The submerged fermentation was performed at 200 rpm and 28 °C for 120 h. The results obtained by the authors in 96 h of fermentation were 4.09 U/mL using free cells and 4.22 U/mL when immobilized cells in 3% (w/v) of calcium alginate were used. When K-carragen and polyacrilamide were used as supports, 3.85 U/mL and 4.07 U/mL were obtained, respectively. Equation 1 presents the coded optimized model for lipase production as a function of glutaraldehyde concentration and curing time. The ANOVA analysis for lipase activity showed high correlation coefficient (R ) 0.97) and a good performance of the F-test for regressions. The model generated the response surface depicted in Figure 1.
9654 Ind. Eng. Chem. Res., Vol. 47, No. 23, 2008 Table 3. Matrix of the First Experimental Design (Coded and Real Values) with the Responses in Terms of Lipase Activity run
X1
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17a 18a 19a
-1(2) +1(4) -1(2) +1(4) -1(2) +1(4) -1(2) +1(4) -1(2) +1(4) -1(2) +1(4) -1(2) +1(4) -1(2) +1(4) 0(3) 0(3) 0(3)
X2
X3
-1(0) -1(0.2) -1(0) -1(0.2) +1(2) -1(0.2) +1(2) -1(0.2) -1(0) +1(2) -1(0) +1(2) +1(2) +1(2) +1(2) +1(2) -1(0) -1(0.2) -1(0) -1(0.2) +1(2) -1(0.2) +1(2) -1(0.2) -1(0) +1(2) -1(0) +1(2) +1(2) +1(2) +1(2) +1(2) 0(1) 0(1.1) 0(1) 0(1.1) 0(1) 0(1.1) control experimentb
X4 -1(6) -1(6) -1(6) -1(6) -1(6) -1(6) -1(6) -1(6) +1(24) +1(24) +1(24) +1(24) +1(24) +1(24) +1(24) +1(24) 0(15) 0(15) 0(15)
LA (U/mL) 24 h
LA (U/mL) 48 h
LA (U/mL) 72 h
LA (U/mL) 96 h
LA (U/mL) 120 h
0.97 1.63 0.97 0.92 0 0 0.81 0 0.91 0 0.59 0 0 0 0 0 1.63 1.60 1.69 4.27
0.67 2.03 7.79 1.16 0 0 0.97 0 4.16 0 1.45 0 0 0 0 0 2.40 2.38 2.93 5.41
0 2.93 11.36 9.44 0 0 5.12 0 5.12 0 2.14 0 0 0 0 0 1.02 1.12 1.24 6.20
0 3.01 5.19 12.81 0 0 4.12 0 4.2 0 6.2 0 0 0 0 0 0 0 0 6.31
0 2.23 3.36 3.98 0 0 2.95 0 0 0 0 0 0 0 0 0 0 0 0 3.54
a Central point. b Control experiment, free microorganism (peptone 2% (w/v), NaCl 0.5% (w/v), yeast extract 0.5% (w/v), olive oil 1% (w/v), and temperature 28 °C), where X1 ) sodium alginate (%, w/v), X2 ) glutaraldehyde (%, v/v), X3 ) CaCl2 (%, w/v), X4 ) curing time (h); and LA) lipase activity (U/mL).
Table 4. Matrix of the Second Experimental Design (Coded and Real Values) with the Responses in Terms of Lipase Activity run
X1
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17a 18a 19a
-1(2) +1(3) -1(2) +1(3) -1(2) +1(3) -1(2) +1(3) -1(2) +1(3) -1(2) +1(3) -1(2) +1(3) -1(2) +1(3) 0(2.5) 0(2.5) 0(2.5)
X2
X3
-1(0.5) -1(0.2) -1(0.5) -1(0.2) +1(1.5) -1(0.2) +1(1.5) -1(0.2) -1(0.5) +1(0.8) -1(0.5) +1(0.8) +1(1.5) +1(0.8) +1(1.5) +1(0.8) -1(0.5) -1(0.2) -1(0.5) -1(0.2) +1(1.5) -1(0.2) +1(1.5) -1(0.2) -1(0.5) +1(0.8) -1(0.5) +1(0.8) +1(1.5) +1(0.8) +1(1.5) +1(0.8) 0(1) 0(0.6) 0(1) 0(0.6) 0(1) 0(0.6) control experimentb
X4 -1(1) -1(1) -1(1) -1(1) -1(1) -1(1) -1(1) -1(1) +1(11) +1(11) +1(11) +1(11) +1(11) +1(11) +1(11) +1(11) 0(6) 0(6) 0(6)
LA (U/mL) 24 h
LA (U/mL) 48 h
LA (U/mL) 72 h
LA (U/mL) 96 h
LA (U/mL) 120 h
6.32 0.81 0.48 0 3.36 0 0 0 0 0 0 1.44 0 0.32 0 0 1.31 1.28 1.61 3.23
6.80 0.32 5.61 3.52 4.64 0 1.12 0 0.32 2.08 5.28 1.60 0 0 0 0 5.28 5.44 5.36 5.34
4.32 0 5.61 9.76 7.84 0 2.24 0 0 2.96 6.88 5.28 0 0 0.16 2.40 11.91 12.10 12.32 7.83
2.31 0 2.46 13.08 3.68 0 0.80 0 0 3.04 8.99 12.48 0 0 0 3.68 12.89 13.18 12.80 8.58
0 0 0 11.02 0 0 0 0 0 0 7.98 11.21 0 0 0 0 11.98 12.0 11.79
a Central point. b Control experiment, free microorganism (peptone 2% (w/v), NaCl 0.5% (w/v), yeast extract 0.5% (w/v), olive oil 1% (w/v), and temperature 28 °C), where X1 ) sodium alginate (%, w/v), X2 ) glutaraldehyde (%, v/v), X3 ) CaCl2 (%, w/v), X4 ) curing time (h); and LA) lipase activity (U/mL).
Lipase activity (U/mL) ) 22.18 - 1.47 · glutaraldehyde(L) 0.69 · glutaraldehyde(Q) + 0.48 · time(L) - 0.38 · time(Q) + 0.27 · glutaraldehyde(L) · time(L) (1) From the response surface and contour curves presented in Figure 1, an optimum region comprising the curing time of 5-7 h and glutaraldehyde concentration of 1-2% (v/v) can be observed, which means the appropriate range for reaching the best results for lipase activity. It is worth noticing that, within the range of the independent variables investigated, at the boundary conditions, a sharp decrease in lipase activity may be observed. With respect to literature data, the work of Hemachander et al. may be cited,18 who obtained for lipase production by submerged fermentation of immobilized cells of Ralstonia pickettii in different supports, in 120 h of fermentation, 8.5 U/mL for immobilized cells in agar, 14 U/mL in 4% (w/v) of calcium alginate, and 24.75 U/mL in polyacrilamide. Benjamin and
Pandey25 used immobilized Candida rugosa for lipase production in a packed bed reactor and obtained 17.94 U/mL with cells immobilized in calcium alginate. Characterization of Crude Enzymatic Extract. After optimizing the lipase production using immobilized cells of a newly isolated Penicillium sp. by the sequential strategy of experimental design, a partial characterization of the crude enzymatic extract was carried out in terms of optimum temperature and pH values, temperature and pH of stability, and stability to storage at low temperatures. A full 22 experimental design with 4 axial points and 3 central points was performed for determining the optimum temperature and pH. Results obtained in this step are presented in Table 7, which shows that higher lipase activities were obtained at the central point (18.2 U/mL) and in experiments 6 and 2, respectively, with pH of 7 and 5.5 and temperature of 44 and 42 °C. However, experiments 1, 5 and 7 with pH of 5.5, 7.0,
Ind. Eng. Chem. Res., Vol. 47, No. 23, 2008 9655 Table 5. Matrix of the Third Experimental Design (Coded and Real Values) with the Responses in Terms of Lipase Activity run
X1 -1(1) 1(2) -1(1) 1(2) -1.41(0.8) 1.41(2,2) 0(1.5) 0(1.5) 0(1.5) 0(1.5) 0(1.5) controlb
1 2 3 4 5 6 7 8 9a 10a 11a
X2 -1(3) -1(3) 1(9) 1(9) 0(6) 0(6) -1.41(1.7) 1.41(10.2) 0(6) 0(6) 0(6)
LA (U/mL) 24 h
LA (U/mL) 48 h
LA (U/mL) 72 h
LA (U/mL) 96 h
LA (U/mL) 120 h
0 0 0.8 0 0 0 0 0 0 0 0 4.64
2.90 0 1.62 1.01 2.08 0 4.48 0.4 0.21 0.16 0.16 5.12
5.80 7.04 5.6 5.28 5.6 4.44 8.96 5.60 5.30 5.12 5.81 10.08
6.72 13.6 11.2 12.48 12.0 10.88 16.96 12.8 14.72 14.48 15.01 9.76
6.70 11.21 14.24 15.52 13.0 19.2 17.0 20.09 19.01 19.61 19.82 5.19
a Central point. b Control experiment, free microorganism (peptone 2% (w/v), NaCl 0.5% (w/v), yeast extract 0.5% (w/v), olive oil 1% (w/v), and temperature 28 °C), where X1 ) glutaraldehyde (%, v/v), X2 ) curing time (h); and LA ) lipase activity (U/mL).
Table 6. Matrix of the Fourth Experimental Design (Coded and Real Values) with the Responses in Terms of Lipase Activity run
X1 -1(0.4) 1(2.6) -1(0.4) 1(2.6) -1.41(0) 1.41(3) 0(1.5) 0(1.5) 0(1.5) 0(1.5) 0(1.5) controlb
1 2 3 4 5 6 7 8 9a 10a 11a
X2 -1(1.9) -1(1.9) 1(10.1) 1(10.1) 0(6) 0(6) -1.41(0.25) 1.41(11.75) 0(6) 0(6) 0(6)
LA (U/mL) 24 h
LA (U/mL) 48 h
LA (U/mL) 72 h
LA (U/mL) 96 h
LA (U/mL) 120 h
0 0 0 0 0.64 0 0 0 0 0 0 3.52
1.10 0.61 1.44 0.12 2.41 0.64 3.99 5.28 7.2 7.0 7.8 5.12
1.88 0.94 3.01 1.08 5.10 0.71 7.00 7.78 10.24 10.01 10.66 7.84
9.01 3.91 9.78 5.76 2.86 1.00 16.01 16.90 22.56 22.00 21.98 9.76
9.0 4.98 10.01 5.76 2.98 1.02 16.00 16.02 20.96 20.01 19.51 6.00
a Central point. b Control experiment, free microorganism (peptone 2% (w/v), NaCl 0.5% (w/v), yeast extract 0.5% (w/v), olive oil 1% (w/v), and temperature 28 °C), where X1 ) glutaraldehyde (%, v/v), X2 ) curing time (h); and LA ) lipase activity (U/mL).
Table 7. Matrix of Full Experimental Design for Evaluation of Optima Temperature and pH (Coded and Real Values) with the Responses in Terms of Lipase Activity run
pH
T (°C)
lipase activity (U/mL)
1 2 3 4 5 6 7 8 9a 10a 11a
-1(5.5) -1(5.5) +1(8.5) +1(8.5) 0(7.0) 0(7.0) -1.41(4.88) +1.41(9.11) 0(7.0) 0(7.0) 0(7.0)
-1(32) +1(42) -1(32) +1(42) -1.41(30) +1.41(44) 0(37) 0(37) 0(37) 0(37) 0(37)
14.61 15.58 0.81 0.94 15.08 16.0 15.01 0 18.2 18.08 17.81
a
Central point.
Table 8. Summary of the Thermal Inactivation Parameters for the Crude Enzymatic Extract Obtained from Immobilized Cells of Penicillium sp. temperature (°C)
k (h-1)
half-life time (min)
25 35 45 55 65
0.0048 0.0066 0.0074 0.0081 0.0109
144.4 105.0 93.6 85.5 63.6
and 4.88 presenting similar activities (14.61, 15.08, and 15.01 U/mL), respectively, at temperatures of 32, 30, and 37 °C. One can also observe that at pH 9.11 the enzymatic activity was diminished. On the other hand, it is important to mention that the crude extract obtained from the immobilized microorganism presented a good stability in a large range of pH values. Tan et al.31 determined the optimum pH and temperature for lipase produced by submerged fermentation of free cells of
Candida sp. as being 28 °C and 7.0, respectively. The literature reports that neutral pH is generally found as optimum for lipase activity.6,29-35 The Pareto chart that summarizes the statistical analysis of the results obtained in this step is presented in Figure 2. Temperature and pH presented a positive significant effect on lipase activity (p < 0.05). Some works presented in the literature report that the optimum temperature for lipase activity is approximately 37 °C.6,29,35 In this work, the optimum temperature range found for lipase activity was 37-42 °C, which is in agreement with that presented by Abbas et al.,32 Burkert,6 Kamini and Mala,33 Fadiloglu and Soylemez,34 Freire et al., 30 and Benjamin and Pandey.25 Regarding the optimum pH values obtained in this work, one can verify that they are in agreement with literature data for other lipases (5.0-9.0).6,25,30,32-34 Results obtained for the crude enzymatic extract related to lipase activity as a function of incubation time at different temperatures showed that, within the range investigated, the temperature range from 25 to 35 °C provided the best stability. Kamini and Mala33 concluded that the lipase from Aspergillus niger presented stability when submitted to temperatures up to 40 °C.25 A study performed by Burkert et al.7 showed that the stability of lipase obtained from Geotrichum candidum NRRLY552 reduced with increasing temperature, with 30 °C as the maximum temperature to keep stability. The experimental data obtained in this work were graphically represented as ln A/A0 vs time. From this plot, the thermal deactivation constant (Kd) was obtained for each temperature, as shown in Table 8. With these values, the activation energy for denaturation reaction (Ed) was calculated from eq 2:
9656 Ind. Eng. Chem. Res., Vol. 47, No. 23, 2008
ln Kd ) ln Kd0 -
Ed RT
(2)
By taking the slope of the curve (R2 ) 0.9763), the value of Ed (128 503.26 cal/mol) was obtained. The expression used for Kd determination was Kd (theoretical) ) Kd0 exp(-Ed/RT), with T ) temperature (K).
The half-life time (min) was calculated from eq 3 for the studied temperatures: 0.5 (3) Kd From Table 8 one can observe that the half-life time for the enzymatic extract obtained by immobilized microorganism was higher than that obtained for free microorganism at higher temperatures (55 and 65 °C). The pH of stability for the lipases obtained in this work was assessed varying pH from 4.8 to 8.8 in sodium phosphate buffer 100 mM at 37 °C. Inspection of the results shows that the lipases from Penicillium sp. presented the same rate of activity loss in the whole investigated range. Pastore et al.35 verified that the lipase from Rhizopus sp. showed best stability between pH of 5.0 and 8.5 and 6.5 and 7.5 in tris-HCl buffer for lipase produced by Candida rugosa. Freire et al.30 found a higher stability in the pH range from 7.0 to 8.0 for lipase obtained from Penicillium restrictum. The crude enzymatic extract presented a good stability for 60 days at 4 and -10 °C. t1/2 ) ln
Conclusions The immobilized newly isolated strain of Penicillium sp. used in this work for lipase production yielded good results compared to literature data of free and immobilized cells. The optimized condition for lipase production was determined as 2.5% (w/v) of calcium alginate, 0.2% (w/v) of CaCl2, 1.5% (v/v) of glutaraldehyde, and 6 h as curing time, yielding lipase activity as high as 20.96 U/mL in 120 h of fermentation. The crude enzymatic extract presented optimum pH and temperature of 7.0 and 37 °C, respectively, and maintained its activity for 60 days stored at -10 °C. Acknowledgment The authors thank CAPES/PROCAD, CNPq, and Intecnial S.A. for the financial support of this work and scholarships. Literature Cited
Figure 1. Response surface (a) and contour curve (b) for lipase production. Effect of glutaraldehyde concentration (% v/v) and curing time (h) on lipase activity.
Figure 2. Pareto chart for optimum temperature and pH determination in terms of lipase activity.
(1) Houde, A.; Kademi, A.; Leblanc, D. Lipases and their industrial applications. Appl. Biochem. Biotechnol. 2004, 118, 155. (2) Freire, D. M. G. Selec¸a˜o de Microrganismos lipolı´ticos e estudo da produc¸a˜o de lipases por Penicillium restrictum. Ph.D. thesis, Departamento de Bioquı´mica IQ/UFRJ, Brazil, 1996; p 132. (3) Snellman, E. A.; Sullivan, E. R.; Colwell, R. V. Purification and properties of the extracellular lipase, LipA, of acinetobacter sp. Biochem. Eng. J. 2002, 11, 269. (4) Hasan, F.; Shah, A. A.; Hameed, A. Industrial Applications of microbial lipases. Enzyme Microb. Technol. 2006, 39, 235. (5) Jaeger, K.; Reetz, M. T. Microbial lipases form versatile tools for biotechnology. Trends Biotechnol. 1998, 16, 396. (6) Burkert, J. F. M. Otimizac¸a˜o das Condic¸o˜es de Produc¸a˜o da Lipase por Geotrichum candidum NRRL-Y552. Ph. D. thesis. Departamento de Engenharia de Alimentos DEA/FEA/UNICAMP, Brazil, 2002; p 99. (7) Burkert, J. F. M.; Maugeri, F.; Rodrigues, M. I. Optimization of extracelular lipase production by Geotrichum candidum sp. using factorial design. Bioresour. Technol. 2004, 91, 77. (8) Sharma, R.; Chisti, Y.; Banerjee, U. C. Production, purification, characterization, and applications of lipases. Biotechnol. AdV. 2001, 19, 627. (9) Ko, W. H.; Wang, I. T.; Ann, P. J. A simple method for detection of lipolytic microorganisms in soils. Soil Biol. Biochem. 2005, 37, 597. (10) Bjo¨rkling, F.; Godtfedsen, S. E.; Kirk, O. The future impact of industrial lipases. Tibtech 1991, 9, 360. (11) Cardenas, F.; Alvarez, E.; Castro-Alvarez, M. S.; Sanchez-Montero, J. M. Screening and catalytic activity in organic synthesis of novel fungal and yeast lipases. J. Mol. Catal. B 2001, 14, 111. (12) Wang, Y.; Srivastava, K. C.; Shen, G. J. Thermostable alkaline lipase from a newly isolated thermophilic Bacillus, strain A30-1 (ATCC 53841). J. Ferment. Bioeng. 1995, 79, 433.
Ind. Eng. Chem. Res., Vol. 47, No. 23, 2008 9657 (13) Jaeger, K. E.; Reetz, M. T. Microbial lipases form versatile tools for biotechnology. Tibtech 1998, 16, 396. (14) Kim, J. T.; Kang, S. G.; Woo, J. H.; Lee, J-H.; Jeong, B. C.; Kim, S.-J. Screening and its potential application of lipolytic activity from a marine environment:characterization of a novel esterase from Yarrowia lipolytica CL180. Appl. Microbiol. Biotechnol. 2007, 74, 820. (15) Vargas, G. D.; Treichel, H.; Oliveira, D.; Beneti, S. C.; Freire, D. M. G.; Di Luccio, M. Optimization of lipase production by Penicillium simplicissimum in soybean meal. J. Chem. Technol. Biotechnol. 2007, 83, 47. (16) Carvalho, P. O.; Calafatti, S. A. P.; Marassi, M.; Silda, D. M.; Contesini, F. J.; Bizaco, R.; Macedo, G. A. Potencial de biocata´lise enantiosseletiva de lipases microbianas. Quim. NoVa 2005, 28, 614. (17) Dalla-Vecchia, R.; Nascimento, M. G.; Soldi, V. Aplicac¸o˜es sinte´ticas de lipases imobilizadas em polı´meros. Quim. NoVa 2004, 27, 623. (18) Hemachander, C.; Bose, N.; Puvanakrishnan, R. Whole cell immobilization of Ralstonia pickettii for lipase production. Process Biochem. 2001, 36, 629. (19) Qi, W. T.; Ma, J.; Yu, W. T.; Xie, Y. T.; Wang, W.; Ma, X. Behavior of microbial growth and metabolism in alginate-chitosan-alginate (ACA) microcapsules. Enzyme Microb. Technol. 2006, 38, 697. (20) Peinado, A. R.; Moreno, J. J.; Villaba, M. J.; Gonza´lez-Reyes, J. A. Yeast biocapsules: A new immobilization method and their applications. Enzyme Microb. Technol. 2006, 40, 79. (21) Mariano, G. Q.; Dalton, H. Biotranformation of aromatic compounds by immobilized bacterial strains in bariun alginate beads. Enzyme Microb. Technol. 1999, 24, 232. (22) Orive, G.; Ponce, R. M.; Herna´ndez, A. R.; Gasco´n, M.; Pedraz, I. J. L. Biocompatibility of microcapsules for cells immobilization elaborated with different type of alginates. Biomaterials 2002, 23, 3825. (23) Elibol, M.; Ozer, E. Lipase production by immobilized Rhizopus arrhizus. Process Biochem. 2000, 36, 219. (24) Ellaiah, P.; Prabhakar, T.; Ranakrishna, B.; Taleb, A. T.; Adinarayana, K. Production of lipase immobilized cells of Aspergillus niger. Process Biochem. 2004, 39, 525.
(25) Benjamin, S.; Pandey, A. Enhancement of lipase production during repeated batch culture using immobilized Candida rugosa. Process Biochem. 1997, 32, 437. (26) Yang, X.; Wang, B.; Cui, F.; Tan, T. Production of lipase by repeated batch fermentation with immobilized Rhizopus arrhizus. Process Biochem. 2005, 40, 2095. (27) Muralidhar, R. V.; Chirumamila, R. R.; Marchant, R.; Nigam, P. A response surface approach for the comparison of lipase production by Candida cylindracea using two different carbon sources. Biochem. Eng. J. 2001, 9, 17. (28) Haaland, P. D. Experimental design in biotechnology; Marcel Dekker: New York, 1989. (29) Johnsen, A.; Flink, M. Influence of alginate properties and gel reinforcement on fermentation characteristics of immobilized yeast cells. Enzyme Microb. Technol. 1986, 8, 737. (30) Freire, D. M. G.; Gomes, P. M.; Bom, E. P. S.; Sant´ Anna, G. L., Jr. Lipase production by a new promising strain Penicillium restrictum. ReV. Microbiol. 1997, 28, 6. (31) Tan, T.; Zhang, M.; Wang, B.; Ying, C.; Deng, L. Screening of high lipase producing Candida sp. and production of lipase by fermentation. Process Biochem. 2002, 39, 459. (32) Abbas, H.; Hiol, A.; Deyris, V.; Comeau, L. Isolation and characterization of an extracellular lipase from Mucor sp. strain isolated from palm fruit. Enzyme Microb. Technol. 2002, 31, 968. (33) Kamini, N. R.; Mala, J. G. S. Lipase production from Aspergillus niger by solid-state fermentation using gingerly oil cake. Process Biochem. 1998, 33, 505. (34) Fadiloglu, S.; Soylemez, Z. Kinetics of lipase-catalyzed hydrolysis of olive oil. Food Res. Int. 1997, 30, 171. (35) Pastore, G. M.; Da Costa, V. S. R.; Koblitz, M. G. B. Purificac¸a˜o parcial e caracterizac¸a˜o bioquı´mica de lipase extracelular produzida por nova linhagem de Rhizopus sp. Cienc. Tecnol. Aliment. 2003, 23, 135.
ReceiVed for reView April 22, 2008 ReVised manuscript receiVed September 10, 2008 Accepted September 23, 2008 IE800658J
