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Food Science and Technology

Print version ISSN 0101-2061

Ciênc. Tecnol. Aliment. vol.31 no.3 Campinas July/Sept. 2011

http://dx.doi.org/10.1590/S0101-20612011000300009 

ORIGINAL PAPERS

 

Lipase and esterase - to what extent can this classification be applied accurately?

 

Lipases e esterases: como definir e classificar?

 

 

Danielle Branta LopesI; Laira Priscila FragaI; Luciana Francisco FleuriII, *; Gabriela Alves MacedoI

IDepartamento de Ciência de Alimentos, Faculdade de Engenharia de Alimentos - FEA/DCA, Universidade Estadual de Campinas - Unicamp, Campinas, SP, Brasil
IIDepartamento de Química e Bioquímica, Instituto de Biociências de Botucatu - IBB/DQB, Universidade Estadual Paulista "Júlio de Mesquita Filho" - UNESP, Distrito de Rubião Jr., s/n, CEP 18618-970, Botucatu, SP, Brasil, e-mail: luciana@ibb.unesp.br

 

 


ABSTRACT

Enzyme technology is an ever-growing field of knowledge and, in recent years, this technology has raised renewed interest, due to the search for new paradigms in several productive processes. Lipases, esterases and cutinases are enzymes used in a wide range of processes involving synthesis and hydrolysis reactions. The objective of this work was to investigate and compare the specific lipase and esterase activities of five enzymes - four already classified as lipases and one classified as cutinase - in the presence of natural and synthetic substrates. All tested enzymes presented both esterase and lipase specific activities. The highest specific esterase activity was observed for Aspergillus 1068 lipase in natural substrate and for F. oxysporum cutinase in synthetic substrate, while the highest specific lipase activity was observed for Geotrichum sp. lipase in natural substrate and for F. oxysporum cutinase in synthetic substrate. These results display some interface-independent lipolytic activity for all lipases tested. This is in accordance with the rationale that a new and broader definition of lipases may be necessary.

Keywords: lipase; esterase; cutinase; lipase specific activity; esterase specific activity.


RESUMO

Tecnologia enzimática é um campo de conhecimento cada vez maior, e, nos últimos anos, esta tecnologia tem apresentado um interesse crescente, devido à busca de novos paradigmas em diversos processos produtivos. Lipases, esterases e cutinases são as enzimas usadas em uma ampla gama de processos que envolvem reações de síntese e hidrólise. O objetivo deste trabalho foi investigar e comparar as atividades de lipase e esterase de quatro enzimas já classificadas como lipases e uma como cutinase, na presença de substratos naturais e sintéticos. Todas as enzimas testadas apresentaram atividades específicas tanto de esterase como de lipase. A maior atividade específica de esterase foi observada para a lipase de Aspergillus 1068 em substrato natural e a cutinase de F. oxysporum em substrato sintético, enquanto a atividade específica de lipase mais alta foi observada para a lipase de Geotrichum sp. em substrato natural e a cutinase de F. oxysporum em substrato sintético. Estes resultados permitem visualizar algumas interfaces independentes de atividade lipolítica para todas as lipases testadas. De acordo com estes estudos, uma nova e mais ampla definição de lipase pode ser necessária.

Palavras-chave: lipase; esterase; cutinase; atividade específica de lipase; atividade específica de esterase.


 

 

1 Introduction

Esterases (EC 3.1.1.X) comprise a diverse group of hydrolases that catalyze the cleavage and formation of ester bonds. They are widely distributed in animals, plants and microorganisms. Many of them show a wide range of possible substrates, leading to the assumption that they evolved to enable access to carbon sources or to be involved in catabolic pathways. These enzymes also display high regio- and stereo-specificity, making them attractive biocatalysts in the production of optically pure compounds in fine chemical synthesis.

Two major classes of hydrolases are of utmost importance: 'true' esterases (EC 3.1.1.1, carboxyl ester hydrolases) and lipases (EC 3.1.1.3, triacylglycerol hydrolases) (BORNSCHEUER, 2002). Lipases are mainly active against water-insoluble substrates, such as triglycerides composed by long-chain fatty acids, whereas esterases preferentially hydrolyze 'simple' esters and usually only triglycerides composed by fatty acids shorter than C6 (HELISTÖ; KORPELA, 1998; KULKARNI; GADRE, 2002).

Lipases can be distinguished from carboxyl esterases by their substrate spectra, using p-nitrophenyl palmitate (cleaved by lipases) versus p-nitrophenyl butyrate (cleaved by esterases). Lipases can also be distinguished from esterases by the phenomenon of interfacial activation, which is only observed for lipases. Both enzymes remain stable in organic solvents, but this property is more noticed for lipases (Bornscheuer, 2002).

According to Brenda (2007), cutinases can be classified as EC 3.1.1.3, the recommended name being triacylglycerol lipase, and the systematic name triacylglycerol acyl hydrolyse. The same source also classifies it as 3.1.1.74, the recommended name being cutinase and the systematic name cutin hydrolyse. It is first considered as an esterase that hydrolyzes cutin, but can also works as and be classified as lipase.

Cutinases are characterized by the ability to degrade cutin, the biopolyester that forms the cuticle of higher plants. They were first described in phytopathogenic fungi that grow on cutin as the sole carbon source, cleaving the ester bonds in the cutin polymer (KOLATTUKUDY, 1984; ETTINGER; THUKRAL; KOLATTUKUDY, 1987; EGMOND; VLIEG, 2000). Cutinases are also able to hydrolyze a great variety of synthetic esters, being as efficient as pancreatic lipases on short and long chains of emulsified triacylglycerols (DE GEUS; LAWEREYS; MATTHYSSENS, 1989). Cutinases are active regardless of the presence of a lipid-water interface, either in soluble and emulsified triglycerides (VERGER; HAAS, 1976; LONGHI; CAMBILLAU, 1999; MACEDO; PIO, 2005; PIO; MACEDO, 2007).

Comparisons between carboxylesterases and lipases reveal remarkable sequence similarities in spite of radically different substrate specificities or physiological functions (SCHRAG; CYGLER, 1993; LIN; WU; CHEN, 2001; BENCHARIT et al., 2002; 2003a,b).

The comparative analysis of three-dimensional structures of esterases and lipases reveals a feature known as a/b hydrolase fold, which consists of a central b-sheet surrounded by a variable number of a-helices, and accommodates a catalytic triad composed of serine, histidine, and a carboxylic acid (OLLIS et al., 1992; CYGLER et al., 1993; HOLMQUIST, 2000). The database ESTHER- Esterases, a/b Hydrolase Enzymes and Relatives (INLAND NORTHWEST RESEARCH ALLIANCE, 2007) lists the three-dimensional structures of all known a/b hydrolase fold proteins.

Many lipases and esterases have their active site buried under secondary structural elements that must change conformation to allow substrate access to the active site. These secondary structure elements have been called caps, lids or flaps, and have important roles in regulating accessibility of substrates into the catalytic device.

Hydrolytic enzymes such as lipases and esterases can also catalyze synthesis reactions, once the chemical equilibrium is shifted into the direction of the synthesis reaction (STAMATIS et al., 1998).

Teng and Xu (2007) compared two methods: colorimetric and CG-FID, to test the lipase activity for many commercial lipases. They concluded that using p-NPP the measurement time was greatly shortened and contributed to evaluate the lipase activity in organic solvent.

Titrimetric methods tend to be time consuming, which may render them unsuitable for large-scale screening of lipolytic activity (THOMSON; DELAQUIS; MAZZA, 1999).

 Many literature citations report the use of synthetic substrates for the measurement of lipase activity. In particular, p-nitrophenyl derivatives of fatty acids (generally lauric or palmitic acids) have been popular (BECKER et al., 1997; LABUSCHAGNE; VAN TONDER; LITTHAUER, 1997).

Vorderwulbecke, Kieslich and Erdmann (1992), in a study of Geotrichum candidum lipase, tested the lipolytic activity using p-NPP and the esterase activity using p-NPB. Titrimetric methods for lipase activity determination are widly employed and seem to offer adequate sensitivity in many applications. The method requires little or non-sophisticated equipment and the analysis is straightforward. However, some authors suggest that successful measurement of lipase activity by titration requires stable emulsions and absolute control of pH during hydrolysis (DE MONPEZAT et al., 1990). Method performance may be enhanced by ensuring electrodes are thoroughly cleaned between assays.

Ghamgui, Karra-Chaâbouni and Gargouri (2004) used the titrimetric method to measure the activities of free and immobilized lipases from Rhizopus oryzae, using olive oil emulsion as substrate. The same method was used by Macedo, Park and Pastore (1997) to measure the activity for Geotrichum candidum lipase, achieving 6.22 U.mL-1.

The objective of the present work was to investigate the esterase and lipase specific activities of four lipases and one cutinase, which were isolated in the biochemistry laboratory of the Faculty of Food Engineering at Campinas State University, and one commercial lipase (Lipozyme), in the presence of natural and synthetic substrates.

 

2 Materials and methods

2.1 Enzymes

Geotrichum sp. lipase was obtained according to Macedo (1995). Lipases from Aspergillus sp. strains 1068 and 1099 were produced according to Costa (1996). Rhizopus sp. lipase was produced by Costa (1997). Lipozyme TL IM lipase was obtained from Novozymes® A/S.

Cutinase from Fusarium oxysporum was obtained according to Pio and Macedo (2007).

2.2 Esterase assays

Method 1: Assay using olive oil as substrate

Esterase activity was performed with olive oil, which was prepared as follows: the reaction mixture containing 5 mL of olive oil, 2 mL of 0.1M phosphate buffer (pH 7.0) and 1 mL of the enzymatic extract (10 mg.mL-1) was incubated at 37 °C for 30 minutes with orbital shaking. Immediately after incubation, the system was disrupted by the addition of 15 mL of acetone-ethanol mixture (1:1 v/v) and the liberated free fatty acids were titrated with 0.05M NaOH. All assays were done independently and in duplicate. One unit of esterase activity was defined as the amount of enzyme which liberated 1 µmol of fatty acids per minute.

Method 2: Assay using p-nitrophenyl butyrate (pNPB) as substrate

Esterase activity was determined spectrophotometrically following the hydrolysis of p-nitrophenylbutyrate (pNPB) at 405 nm. An aliquot (0.070 mL) of the enzyme suspension (10 mg.mL-1) was added to 3.43 mL of a reaction mixture with the following composition: 1.12 mM pNPB dissolved in 50 mM phosphate buffer, pH at 7.2, also containing 0.2% (N/P) Triton X-100 and 0.43M tetrahydrofuran. The reaction was monitored for 15 minutes against blank solution (CALADO et al., 2002). All assays were carried out independently and in duplicate. One unit of esterase activity was defined as the amount of esterase required to release 1 µmol of p-nitrophenol in one minute, under the specified conditions. pNPB was purchased from Sigma-Aldrich Brasil (Sao Paulo, Brazil).

2.3 Effect of surfactant concentration on enzyme activity

The esterase activity was determined using pNPB as substrate according to item 2.2 The concentration of the Triton X-100 varied between zero and 0.8% (w/w). The tests were performed twice and the mean values were compared using the Tukey's test, at a confidence interval of 95% and significant level of p < 0.05. The data analysis was carried out using the software "Statistic for Windows" (Microsoft, 1995).

2.4 Lipase assays

Method 1: Assay using olive oil as substrate

Lipase activity was determined using an olive oil emulsion, which was prepared as follows: 25 mL of olive oil and 75 mL of 7% arabic gum solution were emulsified in liquefier for 2 minutes. The reaction mixture containing 5 mL of olive oil emulsion, 2 mL of 0.1M phosphate buffer (pH 7.0) and 1 mL of the enzymatic suspension (10 mg.mL-1) was incubated at 37 °C for 30 minutes with orbital shaking. Immediately after incubation, the emulsion was disrupted by the addition of 15 mL of acetone-ethanol (1:1 v/v) and the liberated free fatty acids were titrated with 0.05M NaOH (MACEDO; PARK; PASTORE, 1997). All assays were carried out independently and in duplicate. One unit of lipase activity was defined as the amount of enzyme which liberated 1 µmol of fatty acids per minute.

Method 2: Assay using p-nitrophenyl palmitate as substrate

A spectrophotometric method was used for fast and routine measurement of lipase activity using p-nitrophenylpalmitate (pNPP) as the substrate (MAHADIK et al., 2002). The assay mixture consisted of 0.9 mL of substrate solution, 0.1 mL of suitable buffer (0.5M) and 0.1 mL of suitable diluted enzyme. The assay mixture was incubated at 37 °C for 30 minutes and the p-nitrophenol released was measured at 410 nm in Spectronic-117 spectrophotometer. All assays were performed independently and in duplicate. One lipase unit (U) was defined as the amount of enzyme needed to release 1 µmol of p-nitrophenol per minute. pNPP was purchased from Sigma-Aldrich Brasil (São Paulo, Brazil).

2.5 Total protein assays

Total protein concentration was determined according to the Bradford (1976) method.

 

3 Results and discussion

3.1 Esterase assays

All enzymes tested, in triplicate, displayed detectable specific esterase activity on both natural and synthetic substrates. The esterase activity was higher in the presence of the natural substrates compared to the synthetic ones for all enzymes tested. Data concerning the specific esterase activity for cutinase and all non-commercial lipases are shown in Figure 1.

Cutinase displayed the highest activity in the presence of the synthetic substrate and the smallest activity in the presence of the natural substrate, among all non-commercial enzymes tested.

In the lipase group, the higher specific esterase activity was registered for the commercial preparation LipozymeTM (specific esterase activity employing the natural substrate was 187.3 U.mg-1 and for the synthetic substrate was 1.8 U.mg-1), followed by Aspergillus 1068 lipase, regardless of the employed method.

3.2 Effect of surfactant concentration on enzyme activity

Enzymatic activities measured, in triplicate, with and without the presence of the surfactant Triton X-100 are displayed in Table 1. With respect to Aspergillus 1068 and Aspergillus 1099 lipases, the addition of Triton X-100 resulted in a decrease in the enzymatic activity regardless of the Triton concentration. The Rhizopus lipase presented the highest activity in the presence of 0.4% Triton X-100, but also presented activity, although smaller, without the surfactant.

 

 

The lipase from Geotrichum sp. showed discrete influence by the presence of Triton X-100, being observed only a tendency of decrease in the enzymatic activity with the increase in Triton concentration.

The LipozymeTM displayed a behavior similar to Rhizopus lipase, although with higher activities overall.

The cutinase activity was statistically the same in the absence or in the presence of Triton X-100 0.2%. However, the enzymatic activity was smaller in the presence of 0.4 and 0.8% surfactant concentrations.

Nozaki, Reynolds and Tanford (1974) described three interaction levels between proteins and surfactants. The first level predicts possible associations involving bonding sites on the protein molecular surface. The second level describes a cooperative association between the protein and a large number of surfactant molecules, without major conformational changes. The third level comprises a cooperative association with protein denaturation.

Unlike anionic and cationic tensoactives, which are generally denaturant to proteins, non-ionic tensoactives like Triton X-100 do not induce conformational changes with loss of biologic properties in proteins.

Triton X-100 has a restrict capacity to break protein-protein bonds, and most proteins retain their quaternary structure even in the presence of high concentrations of this tensoactive. Triton X-100 is well fitted for determination of lipase hydrolytic activity (ROCHA, 1999).

Kim et al. (2002), in a study of characterization of an esterase from Pseudomonas sp. S34 with high stereospecificity to ketoprofen ethyl ester, observed that the enzyme activity was markedly increased (four-fold) with addition of the non-ionic detergent Triton X-100.

Choi et al. (2003) studied the activity of an esterase from E. coli KCTC1767 in the presence of several surfactants, observing that the highest activity occurred in the presence of Triton X-100.

Prazeres, Cruz and Pastore (2006) tested the influence of different surfactants in the activity of Fusarium oxysporum lipase. It was concluded that the addition of 10% Triton-X had a positive influence on the enzyme activity.

3.3 Lipase assays

All the enzymes tested in triplicate displayed detectable specific lipase activity in the presence of natural and synthetic substrates. On the whole, the highest activities were registered for LipozymeTM (the specific lipase activity employing the natural substrate (olive oil emulsion) was 297.4 U.mg-1, while 45.8 U.mg-1 was registered for the synthetic substrate (pNPP). Figure 2 presents the values of specific lipase activities in the presence of synthetic and natural substrates only for non-commercial enzymes. Among these, the highest activities were observed for Geotrichum sp. lipase in the presence of natural substrate, and for F. oxysporum cutinase in the presence of synthetic substrate.

Cutinase also registered the smallest activity in the presence of the natural substrate. The specific lipase activity profiles of Aspergillus 1099 and Aspergillus 1068 lipases were very similar, both in the presence of natural and synthetic substrates.

According to their current definition, lipases are enzymes that hydrolyze emulsified triglycerides. Due to the broad field of biotechnological applications of these enzymes, this definition is becoming insufficient, and a new and more comprehensive one is necessary.

Lipases are classified by Vorderwulbecke, Kieslich and Erdmann (1992) as enzymes that hydrolyze and also synthesize esters; they can also act regardless of the presence of an interface, under well-established conditions, retaining catalytic activity in either organic or non-organic solvents. This new definition poses a challenge to define lipolytic activity, and to select the standard assay. However, there is no universal assay, and the use of the same method, in the same organism, can result in different values, due to differences in storage time, purification process and source of the microorganism.

The use of water-soluble substrates, such as p-nitrophenyl palmitate (pNPP) (MAHADIK et al., 2002), presents some advantages, due to their short reaction time and easy readings through spectrophotometry. However, these substrates are not the most adequate to determine of true lipase activity, because they are also hydrolyzed by esterases (ERDMANN et al., 1990). Pending this limitation, the use of this substrate in a spectrophotometry assay for determination of lipase activity in organic media is well described by Pencreac'h and Baratti (1996).

The most employed method for lipase activity determination performs the titrating of free fatty acids generated from triglyceride hydrolysis, in an emulsion stabilized by a tensoactive agent. Arabic gum is described as an adequate tensoactive agent, resulting in high values of lipolytic activity, although the effect of emulsifiers in the activity of microbial lipases is not thoroughly understood (VEERARAGAVAN, 1990).

 

4 Conclusions

All enzymes tested displayed both esterase and lipase specific activities. The highest specific esterase activity was observed for Aspergillus 1068 lipase in natural substrate and F. oxysporum cutinase in pNPB substrate. For lipase, the highest activity was observed for Geotrichum sp. lipase in natural substrate and F. oxysporum cutinase in pNPP. F. oxysporum cutinase displayed the highest specific lipase and esterase activities in the presence of synthetic substrate, among all enzymes tested.

Considering the biotechnological applications of lipases, the current definition of these enzymes as being able to hydrolyze emulsified triglycerides is really insufficient, becoming necessary the development of a new and broader definition, which is already being suggested in the literature. The results of this study are in accordance with the rationale that lipases should be classified as enzymes which are able to catalyze reactions of hydrolysis and synthesis of esters and are also able to react, regardless of the presence of an interface under well-defined experimental parameters. The definition of lipolytic activity is a difficult task in the context of this new lipase definition. The enzyme should not be defined through a single method of activity determination and the complete biochemical characterization of the enzyme is essential.

 

Acknowledgements

The authors are grateful to 'Conselho Nacional de Desenvolvimento Científico e Tecnológico' (CNPq) and to 'Fundação de Amparo à Pesquisa do Estado de São Paulo' (FAPESP) their financial support.

 

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Received 26/5/2009
Accepted 24/3/2010 (004215)

 

 

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