Lipase from liver of seabass (Lates calcarifer): Characteristics and the use for defatting of fish skin

Food Chemistry 240 (2018) 9–15

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Lipase from liver of seabass (Lates calcarifer): Characteristics and the use for defatting of fish skin Thanasak Sae-leaw, Soottawat Benjakul

MARK

⁎

Department of Food Technology, Faculty of Agro-Industry, Prince of Songkla University, Hat Yai, Songkhla 90112, Thailand

A R T I C L E I N F O

A B S T R A C T

Keywords: Seabass Viscera Lipase Liver Enzyme Purification Defatting Skin

Lipase from liver of seabass (Lates calcarifer), with a molecular weight of 60 kDa, was purified to homogeneity using ammonium sulfate precipitation and a series of chromatographies, including diethylaminoethyl sepharose (DEAE) and Sephadex G-75 size exclusion columns. The optimal pH and temperature were 8.0 and 50 °C, respectively. Purified lipase had Michaelis–Menten constant (Km) and catalytic constant (kcat) of 0.30 mM and 2.16 s−1, respectively, when p-nitrophenyl palmitate (p-NPP) was used as the substrate. When seabass skin was treated with crude lipase from seabass liver at various levels (0.15 and 0.30 units/g dry skin) for 1–3 h at 30 °C, the skin treated with lipase at 0.30 units/g dry skin for 3 h had the highest lipid removal (84.57%) with lower lipid distribution in skin. Efficacy in defatting was higher than when isopropanol was used. Thus, lipase from liver of seabass could be used to remove fat in fish skin.

1. Introduction Lipases are the enzymes that catalyze the hydrolysis of ester bonds in substrates, such as triacylglycerol (Wong & Schotz, 2002). Lipases have been known to show their detrimental effects on the quality of food products, via liberating free fatty acids prone to oxidation. On the other hand, there is a wider range of actual and potential applications of lipases, ranging from cleaning products to modified foods, flavour development, biodiesel production, and synthesis of structured lipids (Kurtovic, Marshall, Zhao, & Simpson, 2010). Important applications of lipases in foods include modification of flavour and preferential hydrolysis of ethyl esters of polyunsaturated fatty acids (PUFAs) for enrichment of eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) in marine oils (Shahidi & Wanasundara, 1998). Lipases have been used intensively in the dairy industry for the hydrolysis of milk fat, contributing to flavour enhancement in cheeses, creams and other milk products, and accelerating cheese ripening. In addition, lipases can also be used to prevent the generation of trans-fats in margarines (Kurtovic, Marshall, Zhao, & Simpson, 2009). The attachment of a fatty acid to the bioactive compounds, such as antioxidants using lipases provide the improved solubility in food and cosmetic applications (Kurtovic & Marshall, 2013). For pharmaceutical application, lipases have been used to synthesize lovastatin, a drug that lowers serum cholesterol level (Choudhury & Bhunia, 2015). Recently, lipase from the Pacific white shrimp hepatopancreas has been applied as a biocatalyst for the biodiesel production (Kuepethkaew, Sangkharak,

⁎

Corresponding author. E-mail address: [email protected] (S. Benjakul).

http://dx.doi.org/10.1016/j.foodchem.2017.07.089 Received 21 February 2017; Received in revised form 26 May 2017; Accepted 18 July 2017 Available online 19 July 2017 0308-8146/ © 2017 Elsevier Ltd. All rights reserved.

Benjakul, & Klomklao, 2017a, 2017b). Additionally, it could be used as the detergent, in which efficacy was comparable to the commercial laundry detergents (Kuepethkaew et al., 2017a, 2017b). By-products from fish processing industries, especially digestive organs, are the potential sources of numerous enzymes, such as lipases and proteases. Fish lipases exhibit some characteristics and properties that complement those of lipases from mammalian and microbial sources due to the evolutionary pathways, diets and habitats of fish (Kurtovic et al., 2009). In general, fish lipases have cold-adapted properties, which are known to show more catalytic activity and stability than mammalian lipases, thereby making them suitable for applications in food processing at low temperatures (Morrissey & Okada, 2007). Moreover, they are salt tolerant and can be advantageous in certain food applications (Haard, 1998). Extraction, purification and properties of lipases from fish viscera have been reported. Nayak, Viswanathan Nair, Ammu, and Mathew (2003) prepared crude lipase extracts from the stomach and pyloric caeca, liver, intestine and red muscle of four species of fish, including rohu, oil sardine, mullet and Indian mackerel. Moreover, lipases from different digestive organs of various fish species were reported for golden grey mullet (Smichi, Gargouri, Miled, & Fendri, 2013), cod (Gjellesvik, Lombardo, & Walther, 1992), sardine (Smichi et al., 2010), red sea bream (Iijima, Tanaka, & Ota, 1998), carp (Görgün & Akpınar, 2012), and Chinook salmon and New Zealand hoki (Kurtovic et al., 2010). Seabass (Lates calcarifer) has become popular in many dishes in Thailand and other countries in tropical and subtropical regions of Asia

Food Chemistry 240 (2018) 9–15

T. Sae-leaw, S. Benjakul

2.3. Preparation of ammonium sulfate fraction

and the Pacific. During dressing, by-products, such as skin, bone and viscera, are generated (Sae-leaw, O'Callaghan, Benjakul, & O'Brien, 2016). Seabass skin can be used as a good source for production of collagen, gelatin and hydrolyzed collagen (Sae-leaw, Benjakul, & O'Brien, 2016a, 2016b; Sinthusamran, Benjakul, & Kishimura, 2013). However, seabass skin contains high amounts of fat or lipids, which can be oxidized during extraction at high temperatures. This phenomenon is associated with the development of undesirable odour/flavour, especially fishy odour in the final products, including gelatin (Sae-leaw, Benjakul, & O'Brien, 2016a, 2016b). To alleviate such a problem, defatting of skin has been employed. Sae-leaw et al. (2016a, 2016b) reported that defatting of seabass skin using isopropanol could reduce the fat content and fishy odour in gelatin to some degree. The use of lipase, particularly from fish viscera, can be another promising approach to remove the lipids from fish skin. Also, the solvents are omitted, thereby reducing the problem associated with the cost of solvents and their residues in the products. Moreover, a lipase-catalyzed reaction is a more efficient, selective and environment-friendly process, compared to chemical treatment (Kumar, Dhar, Kanwar, & Arora, 2016). Lipase could induce the hydrolysis of ester bonds, thereby releasing free fatty acid and glycerol (Wong & Schotz, 2002). Those small substances could be leached out during washing effectively. As a consequence, the remaining lipid was drastically reduced. Although extraction of lipases from fish viscera has been reported, no information regarding the characteristics and application of fish lipase, especially from seabass liver, for defatting of fish skin exists. Therefore, the objectives of this study were to extract, purify and characterize lipase from seabass liver and to use crude lipase for defatting of seabass skin.

CLE (200 ml) from internal organ with the highest lipase activity (liver extract) was subjected to ammonium sulfate precipitation at 40–60% saturation at 4 °C for 30 min. The mixture was centrifuged at 9000×g for 15 min at 4 °C and the pellet was dissolved using extraction buffer (20 ml). The solution was dialyzed with extraction buffer for 12 h at 4 °C. The dialysis buffer was changed every 4 h. The dialysate was referred to as “ammonium sulfate fraction, ASF.” 2.4. Purification of lipase All purification steps were conducted at 4 °C. ASF was purified using a series of chromatographies, including DEAE-sepharose anion exchange and Sephadex G-75 gel filtration columns as described by Senphan et al. (2015). Fractions of 3 ml were collected and those with the highest lipase activity were pooled and further lyophilized using a freeze-dryer. 2.5. Protein determination Protein concentration was determined as per the method of Lowry, Rosebrough, Farr, and Randall (1951). Bovine serum albumin was used as a standard. 2.6. Lipase activity assay Lipase activity was determined using p-nitrophenyl palmitate (pNPP) as a substrate according to the method of Kurtovic et al. (2010) with some modifications. p-NPP was dissolved in isopropanol to obtain a concentration of 15 mM and used as stock solution. Substrate working solution was 0.25 mM p-NPP in 20 mM Tris–HCl buffer (pH 8.0) containing 20 mM CaCl2, 5 mM Na cholate and 0.01% gum arabic. The substrate solution was left at 50 °C for 20 min before the assay. Enzyme solution with an appropriate dilution (20 µl) was added into 3.0 ml of substrate working solution. The reaction mixture was incubated at 50 °C for 5 min. The release of p-nitrophenol (p-NP) was measured at 410 nm using a UV-1601 spectrophotometer (Shimadzu, Kyoto, Japan). One unit (U) of activity was defined as the amount of enzyme producing 1 µmol p-NP per min under the assay condition.

2. Materials and methods 2.1. Chemicals β-mercaptoethanol (β-ME) was obtained from Sigma Chemical Co. (St. Louis, MO). Coomassie brilliant blue R-250, sodium dodecyl sulfate (SDS) and N,N,N′,N′-tetramethylethylenediamine (TEMED) were purchased from Bio-Rad Laboratories (Hercules, CA). Glycine, tris (hydroxymethyl) aminomethane (Tris–HCl) and ammonium sulfate were procured from Merck (Darmstadt, Germany). Diethylaminoethyl (DEAE)-Sepharose Fast Flow, Sephadex G-75 and low molecular weight protein markers were procured from GE Healthcare Life Sciences (Uppsala, Sweden).

2.7. SDS-PAGE and native-PAGE

2.2. Preparation and extraction of crude lipase from different internal organs of seabass

SDS-PAGE was performed according to the method of Laemmli (1970). Enzyme solution was mixed with the SDS-PAGE sample buffer (0.125 M Tris–HCl, pH 6.8; 4% SDS; 20% glycerol; 10% βME) and boiled for 3 min. The sample was loaded onto the 4% stacking and 12% separating gels and subjected to electrophoresis. The gels were subsequently stained and detained (Senphan et al., 2015). Native-PAGE was also done as described by Senphan et al. (2015).

Whole viscera of seabass were purchased from a local market in Hat Yai (Songkhla, Thailand). Samples were packed in polyethylene bag, stored in ice using a sample/ice ratio of 1:2 (w/w) and transported. Thereafter, different internal organs, including pyloric caeca, intestine, liver and stomach, were separated. Each sample was chopped into small pieces, rapidly frozen in liquid nitrogen and powdered using a blender (Model MX-898N, Panasonic, Panasonic Sdn. Bhd., Kuala Lumpur, Malaysia). The samples were defatted using acetone as described by Senphan, Benjakul, and Kishimura (2015). The defatted powder named “acetone powder” was used for lipase extraction. Crude lipase extract was prepared according to the method of Bouchaâla et al. (2015) with some modifications. Acetone powder (50 g) was mixed with extraction buffer (25 mM Tris–HCl, pH 8.0 containing 2.5 mM benzamidine and 1 mM CaCl2) at a ratio of 1:50 (w/ v) and stirred at 4 °C for 3 h. To remove the tissue debris, the suspension was centrifuged for 10 min at 4 °C at 10,000×g using a refrigerated centrifuge (model CR22N, Hitachi, Hitachi Koki Co., Ltd., Tokyo, Japan). The supernatant was filtered through a Whatman No.1 filter paper. The filtrate obtained was referred to as “crude lipase extract, CLE.”

2.8. pH and temperature profiles The pH profile of purified lipase was performed over the pH range of 5.0–9.0. For the temperature profile study, the lipase activity was measured at various temperatures (30, 40, 45, 50, 55, 60 and 70 °C) for 10 min at pH 8.0. 2.9. Kinetic study Kinetic study of purified lipase from seabass liver was carried out using p-NPP (0.1 to 1.0 mM) as the substrate. The kinetic parameters, including the maximal velocity (Vmax) and Michaelis–Menten constant (Km), were measured at pH 8.0 and 50 °C using a Lineweaver–Burk double-reciprocal plot (Lineweaver & Burk, 1934). Catalytic constant (kcat) was calculated from the following equation: kcat = Vmax/[E], 10

Food Chemistry 240 (2018) 9–15

T. Sae-leaw, S. Benjakul

Table 1 Purification of lipase from liver of seabass. Purification steps

Total protein (mg)

Total activity (unit)*

Specific activity (unit/mg)

Yield (%)

Purification fold

Crude lipase extract (NH4)2SO4 (40–60%) Diethylaminoethyl-sepharose Sephadex G-75

2775.00 400.40 10.19 0.53

59.93 11.85 2.43 0.48

0.02 0.03 0.24 0.92

100.00 19.77 4.06 0.81

1.00 1.37 11.06 42.39

* Lipase activity was assayed at pH 8.0 and 50 °C for 5 min using p-nitrophenyl palmitate as a substrate.

3. Results and discussion

where [E] is molar concentration of enzyme calculated based on the MW determined by SDS-PAGE and protein concentration.

3.1. Lipase activities of different internal organs 2.10. Study on the use of crude lipase extract (CLE) for defatting of seabass skin

Crude lipase extracts (CLE) from liver of seabass had the highest lipase activity (7.24 units/g tissue) (P < 0.05), followed by intestine (6.34 units/g tissue), pyloric caeca (5.73 units/g tissue) and stomach (2.40 units/g tissue), respectively. The differences in lipase activities between various internal organs of Antarctic fish were reported by Sidell and Hazel (2002). Lipase activity of liver was higher than those of heart, oxidative skeletal muscle and adipose tissue from four species of Antarctic fish. When lipases were extracted from internal organs of rohu, oil sardine, mullet and Indian mackerel, the highest activities were observed in intestine and stomach/caeca (Nayak et al., 2003). Thus, liver of seabass was used for extraction and purification of lipase.

2.10.1. Preparation of seabass skin Pretreatment of seabass skins was performed by alkaline treatment, as described by Sae-leaw et al. (2016a, 2016b), to remove non-collagenous proteins. Alkali-treated skins were washed with tap water until a neutral or slightly basic pH (7.0–7.5) of wash water was obtained. Thereafter, the skins (100 g) were swollen with 1 l of 0.05 M acetic acid, followed by thorough washing until wash water became neutral or slightly acidic in pH (6.5–7.0). 2.10.2. Treatment of seabass skin with CLE Swollen seabass skins (300 g) were treated with CLE at levels of 0.15 and 0.30 unit/g dry skin. The incubation was performed at 30 °C for various times (1, 2 and 3 h) with manual stirring every 15 min. At the designated time, the treated skins were washed with tap water for 15 min. The samples were then analyzed for lipid contents, using the method of Bligh and Dyer (1959). Swollen skins defatted with 30% (v/ v) isopropanol with a skin/solvent ratio of 1:10 (w/v) for 1 h were also prepared and used as the control.

3.2. Purification of lipase from liver of seabass Purification of CLE from the liver of seabass using a series of chromatographies after precipitation with ammonium sulfate (40–60% saturation) are summarized in Table 1. Ammonium sulfate fraction (ASF) showed an increased purity by 1.37-fold with a yield of 19.77%. Ammonium sulfate precipitation is widely used in the initial step of enzyme purification to eliminate other contaminating proteins from the crude enzyme extract (Khantaphant & Benjakul, 2010). Three activity peaks were observed when ASF was loaded onto DEAE-sepharose column (Fig. 1a). Purity of DEAE-sepharose fraction was 11.06-fold. Ion-exchange chromatography is the most common chromatographic method for enzyme purification. It is effective at the earlier stage in the fractionation of enzymes by removal of contaminating proteins and it can separate enzymes into different isoforms (Khantaphant & Benjakul, 2010). The selected fractions with the highest lipase activity were collected and further loaded onto Sephadex G-75 gel filtration column (Fig. 1b). Two lipase activity peaks were found. The highest activity peak was collected and an increase in purity by 42.39-fold was attained, compared to that of CLE. Gel filtration is the second most frequently used for enzymes purification (Vijayaraghavan, Raj, & Vincent, 2016). Based on gel filtration chromatography, MW of lipase was calculated to be in the range of 55.5–67.6 kDa. SDS-PAGE result indicated that CLE contained several protein bands (Fig. 2a, lane C), representing enzymes, as well as other proteins. After being purified using DEAE-sepharose, some protein bands were eliminated (Fig. 2a, lane D). Only one single band was detected in Sephadex G-75 fraction (Fig. 2a, lane S). The results suggested that anion exchange (DEAE-sepharose) and gel filtration (Sephadex G-75) chromatographies could remove the contaminants from the crude enzyme extract effectively. As a consequence, the purified lipase was obtained. MW of lipase was estimated to be 60 kDa. From native-PAGE, only a single protein band was obtained (Fig. 2b, lane S), confirming that lipase was purified to homogeneity. Lipases have been isolated and characterized from different organs of various fish. Pancreatic lipase from stingray had an apparent MW of 55 kDa (Bouchaâla et al., 2015). Lipase from grey mullet viscera with MW 35 kDa was purified using ammonium sulfate precipitation, gel filtration, anion and cation exchange chromatographies (Smichi et al.,

2.11. Fourier transform infrared (FTIR) spectroscopy Seabass skin treated with CLE (0.30 unit/g dry skin) for 3 h, defatted with 30% (w/v) isopropanol and skin without any treatment were firstly freeze-dried and subsequently subjected to FTIR analysis. The spectra, were collected in 32 scans at a resolution of 4 cm−1 in the range of 4000–400 cm−1 (mid-IR region) with automatic signal gain, and ratioed against a background spectrum recorded from the clean and empty cell at 25 °C. Analysis of spectral data was performed using the OPUS 3.0 data collection software program (Bruker Co, Ettlingen, Germany.). 2.12. Fluorescence microscopy Lipid distribution in selected seabass skins was examined using a fluorescence microscope (Olympus, BX50, Tokyo, Japan). The skins (10 × 200 × 10 mm3) were suspended in Nile blue A for 5 min in order to stain lipid. Prepared specimens were placed on the microscopy slide. The fluorescence microscope was operated at the excitation wavelength of 533 nm and the emission wavelength of 630 nm. Magnification of 100 × was used. 2.13. Statistical analysis The experiments were conducted in triplicate. The data were analyzed by one-way analysis of variance (ANOVA). The Duncan’s multiple range test was used for comparison of means. Statistical analysis was carried out using the Statistical Package for Social Science (SPSS 11.0 for windows, SPSS Inc., Chicago, IL, USA). 11

Food Chemistry 240 (2018) 9–15

T. Sae-leaw, S. Benjakul

(a)

Lipase activity

0.2

0.050 0.040

A280

0.15

0.030 0.1 0.020 0.05

0.010

0

0.000 0

5

10

15

20

25

30

35

40

45

Relative activity (%)

A280

Lipase activity (unit/ml)

(a)

100 80 60 40 20

50

0

Fraction no.

5 A280

(b)

7

8

9

pH

Lipase activity

0.350

0.200

0.012

0.150

0.008

0.100 0.004

0.050

Relative activity (%)

0.016

0.250

Lipase activity (unit/ml)

0.020

0.300

A280

6

0.000

0.000 0

10

20

30

40

50

60

40 20

30

(a)

40

45

50

55

60

65

70

Fig. 3. pH (a) and temperature profiles (b) of purified lipase from liver of seabass. Bars represent the standard deviation (n = 3).

3.3. pH and temperature profiles The pH and temperature profiles of purified lipase (Sephadex G-75 fraction) from liver of seabass are illustrated in Fig. 3a & b, respectively. The enzyme exhibited the maximal activity at pH 8. The enzyme displayed acidic lability at pH below 6.0, probably owing to the denaturation of enzyme. The activity was also decreased at the alkaline pH (pH 9). The lowered activity at very acidic and alkaline pH was plausibly due to the conformational changes of enzyme under harsh conditions (Sriket et al., 2012). The optimal pH of seabass liver lipase was in agreement with that of lipase from hepatopancreas of red sea bream (pH 7–9) (Iijima et al., 1998). The maximal lipase activity from grey mullet viscera and carp liver was found at pH 8 (Görgün & Akpınar, 2012; Smichi et al., 2013). Lipases from Chinook salmon and New Zealand hoki showed the highest activity at pH 8 and 8.5, respectively (Kurtovic et al., 2010). The optimal temperature of lipase from seabass liver is depicted in Fig. 3b. Lipase from liver of seabass exhibited the highest activity at 50 °C. A sharp decrease in activity was observed at temperature above 55 °C, presumably as a result of thermal denaturation of lipase. The similar result was found in lipase from grey mullet viscera, which showed an optimal temperature at 60 °C. Lipase from pyloric caeca of salmon displayed two optimal temperatures at 35–40 °C and 50 °C (Kurtovic et al., 2010). However, different optimal temperature was found by Lie and Lambertsen (1985), who reported that the highest lipolytic activity from pyloric caeca and anterior ileum of cod was obtained at 37 °C.

97.0 kDa 66.0 kDa 45.0 kDa

30.0 kDa

20.1 kDa

S

35

Temperature (ºC)

(b)

D

60

0

Fig. 1. Elution profiles of ammonium sulfate (40–60% saturation) fraction on Diethylaminoethyl (DEAE)-sepharose column (a) and Sephadex G-75 gel filtration chromatography (b). Elution was performed using a gradient (0–1 M NaCl) with a flow rate of 0.5 ml/min for DEAE-sepharose column and 25 mM Tris–HCl, pH 8.0 containing 2.5 mM benzamidine and 1 mM CaCl2 with a flow rate of 0.5 ml/min for the Sephadex G-75 column. Fractions (3 ml) were determined for lipase activity using p-nitrophenyl palmitate as a substrate. ↔ represents pooled fractions.

C

80

70

Fraction no.

M

(b)

100

S

Fig. 2. Sodium dodecyl sulfate-PAGE (a) and native-PAGE (b) of purified lipase from liver of seabass. M: molecular weight standard; C: crude lipase extract; D: DEAE-sepharose fraction; S: Sephadex G-75 fraction.

2013). Lipase from the digestive gut of Indian major carp, Catla catla had a MW of 70 kDa (Kameshwar Sharma, Boora, & Tyagi, 2014). Lipase from pyloric caeca of sardine (Sardinella aurita) had a MW of 43 kDa (Smichi et al., 2010). Iijima et al. (1998) reported that hepatopancreas of red sea bream, Pagrus major, contained lipase with MW of 64 kDa.

3.4. Kinetic study Km and kcat for the hydrolysis of p-NPP were 0.30 mM and 2.16 s−1, respectively. For the p-NPP hydrolysis, lipase from liver of seabass had a higher Km value than that from grey mullet viscera (0.22 mM) (Aryee, Simpson, & Villalonga, 2007). The Michaelis–Menten constant (Km) is associated with the catalytic power of an enzyme (Van, 2010). This 12

Food Chemistry 240 (2018) 9–15

T. Sae-leaw, S. Benjakul

alcohols, resulted in the formation of fishy odours in gelatin from seabass skin. Additionally, aldehydic compounds might induce the protein cross-linking in skin matrix (Domingues et al., 2013). This plausibly caused the lower extractability of gelatin from the skin. The result suggested that lipase from seabass liver effectively removed lipids from seabass skin. Thus, solvent commonly used for defatting could be omitted. Additionally, lipase was shown to have the higher efficiency in defatting of seabass skin.

reflects the affinity for the substrate. Each enzyme has a characteristic Km for a given substrate (Machielsen, Dijkhuizen, & Oost, 2007). Lipase from seabass liver showed higher Km than those from grey mullet viscera (0.22 mM) (Aryee et al., 2007) and carp liver (0.17 mM) (Görgün & Akpınar, 2012), indicating that a higher substrate was required to obtain Vmax/2. Lipase from seabass liver showed the lower catalytic constant (kcat) than those from Chinook salmon (3.70 s−1) and cod pyloric caeca (193.00 s−1) but exhibited higher value than that from New Zealand hoki pyloric caeca (0.71 s−1) (Gjellesvik et al., 1992; Kurtovic et al., 2010). kcat is another essential parameter determining an enzyme efficiency. The higher kcat value of lipase suggested a higher rate of substrate hydrolysis (Van, 2010). Catalytic efficiency (kcat/Km) of lipase from liver of seabass (75.81 s/mM) was higher than those from Chinook salmon (47.44 s/ mM) and New Zealand hoki pyloric caeca (10.44 s/mM) (Kurtovic et al., 2010). Nevertheless, the purified lipase exhibited a lower catalytic efficiency than did lipase from cod pyloric caeca (1,378.57 s/mM) (Gjellesvik et al., 1992). The higher catalytic efficiency, the more efficient the enzyme is in transforming the substrate to product (Van, 2010). Thus, kinetic parameters of lipase from seabass liver were different than those from other fish species.

3.6. Fourier transform infrared (FTIR) spectra FTIR spectra of seabass skins without and with defatting using solvent or lipase from seabass liver are illustrated in Fig. 4a. Generally, the ester carbonyl functional group of triglycerides was observed at the wavenumber of 1741–1746 cm−1 (Setiowaty, Che Man, Jinap, & Moh, 2000). Peak at wavenumber of 1744 cm−1 were from triglyceride in the skin of seabass. The result indicated the presence of ester bonds between fatty acids and glycerol backbone of triglycerides. Non-defatted seabass skin had the highest peak amplitude in this region, while seabass skin treated with lipase showed the lowest peak amplitude. This was in accordance with the lowest lipid content in seabass skin defatted with CLE (Table 2). A higher amplitude of peak at wavenumber of 3600–3100 cm−1, representing eOH, eNH, ^CH and ]CeH stretching, was observed in non-defatted skin, compared with defatted counterparts. This was more likely due to the higher amounts of lipids in non-defatted skin. Lipids at high content in non-defatted skin were susceptible to lipid oxidation, thus leading to the formation of hydroperoxides. Hydroperoxide moieties exhibit characteristic absorption bands between 3600 and 3400 cm−1 due to their eOOeH stretching vibration (Van de Voort, Ismail, Sedman, Dubois, & Nicodemo, 1994). A larger peak representing hydroperoxide was found in non-defatted skin than those from defatted skins. Therefore, lipase treatment directly affected the lipid content and distribution in the skin of seabass.

3.5. Defatting of seabass skin using crude lipase extract from liver of seabass Reduction of lipids in swollen seabass skin subjected to treatment with CLE at various concentrations and incubation time is shown in Table 2. The different lipase concentrations and incubation time yielded seabass skin with varying residual lipid contents. The use of solvent (30% isopropanol, v/v) could reduce the lipids by 50.32%, compared to that present in swollen skin (without any treatment). With the same incubation time, lipid content in seabass skin treated with higher lipase level was lower than that of skin treated with lower lipase level (P < 0.05). The efficacy of lipase in lipid removal from seabass skin increased as the incubation time increased (P < 0.05) when the same lipase level was used. The highest lipid reduction was observed in seabass skin treated with lipase at 0.30 unit/g dry skin for 3 h (P < 0.05), in which lipids were removed by 84.57% from swollen skin. The result suggested that lipase from liver of seabass could hydrolyze lipids in seabass skin effectively. Free fatty acids and glycerol could be leached out during the subsequent washing with water. This was evidenced by the lower lipid retained in the treated skin. Lipase is able to hydrolyze ester bonds in substrates, especially triacylglycerols (Wong & Schotz, 2002). Seabass skins contain lipids with high degree of unsaturation (Sae-leaw et al., 2016a, 2016b). Oxidation of PUFAs is a major cause of deterioration, such as nutritional losses, and produces undesirable odour/flavour, colour, and toxic compounds, which makes food less acceptable, or unacceptable, to consumers (Kim & Min, 2008). Sae-leaw and Benjakul (2015) reported that lipid oxidation products decomposed from hydroperoxides, especially aldehydes, ketones and

3.7. Fluorescence micrograph Lipid distribution in seabass skins without and with defatting using solvent or lipase was visualized by fluorescence microscope (Fig. 4b). Seabass skin treated with CLE (0.30 unit/g dry skin) for 3 h and that defatted with 30% (w/v) isopropanol had less lipids than the control (non-defatted skin). It was noted that the lowest amount of lipids was noticeable in skin treated with CLE. The lower lipid distribution of skin treated with CLE was in accordance with the lower lipid content (Table 2). Lipids are generally localized in the layers of dermis and subcutis of fish skin (Mittal, Rai, Banerjee, & Agarwal, 1976; Park, Kim, & Kim, 2003). Lipase from seabass liver might penetrate into the swollen skin and hydrolyze lipids to yield glycerol and free fatty acids. Those glycerol and free fatty acids were leached out during the washing of treated skin. This was evidenced by the lower lipid distribution in CLE treated skin, as shown in fluorescence micrograph. For skin defatted with isopropanol, the lipid distribution was lower than the control but was higher than that treated with lipase. The result reconfirmed the higher efficacy of lipase from seabass liver in removal of lipids from swollen skin.

Table 2 Effect of lipase from seabass liver on defatting of seabass skin. Lipase concentration (unit/g dry skin)

Incubation time (h)

Lipid reduction (%)

− 0.15

− 1 2 3 1 2 3

50.32 19.33 52.31 62.26 59.20 77.15 84.57

4. Conclusions

0.30

± ± ± ± ± ± ±

0.66f* 0.29g 0.55e 0.83c 0.87d 0.77b 1.04a

Lipase from seabass liver was extracted and purified. Lipase from liver of seabass had a MW of 60 kDa. Lipase exhibited the highest hydrolytic activity towards p-NPP at 50 °C and pH 8.0. Purified lipase had Km and kcat of 0.30 mM and 2.16 s−1, respectively. Lipase from seabass liver reduced the lipids in seabass skin more effectively than isopropanol. The efficacy of defatting depended on the level of lipase and time used. Thus, liver of seabass could serve as a promising source of lipase, which could be used as an alternative processing aid for fat removal of fish skin.

Values are expressed as mean ± SD (n = 3). Different letters within the same column indicate significant differences (P < 0.05). * Seabass skin defatted with 30% (v/v) isopropanol for 1 h at room temperature (30–32 °C).

13

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3600

Absorbance (A.U.)

Absorbance (A.U.)

(a)

NDF DF-I DF-L

3400

3200

3000

1800

NDF DF-I DF-L

1780

1760

1740

1720

1700

1680

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Fig. 4. FTIR spectra (a) and fluorescence micrograph (b) of non-defatted seabass skins (NDF) and those defatted with isopropanol (DF-I) or with seabass liver lipase (DF-L). Magnification; 100×. Red domains represent lipids. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.) purification. Canadian Journal of Biochemistry and Physiology, 37(8), 911–917. Bouchaâla, E., BouAli, M., Ali, Y. B., Miled, N., Gargouri, Y., & Fendri, A. (2015). Biochemical characterization and molecular modeling of pancreatic lipase from a cartilaginous fish, the common stingray (Dasyatis pastinaca). Applied Biochemistry and Biotechnology, 176(1), 151–169. Choudhury, P., & Bhunia, B. (2015). Industrial application of lipase: A review. Biopharm Journal, 1(2), 41–47. Domingues, R. M., Domingues, P., Melo, T., Pérez-Sala, D., Reis, A., & Spickett, C. M. (2013). Lipoxidation adducts with peptides and proteins: Deleterious modifications or signaling mechanisms? Journal of Proteomics, 92, 110–131. Gjellesvik, D. R., Lombardo, D., & Walther, B. T. (1992). Pancreatic bile salt dependent lipase from cod (Gadus morhua): Purification and properties. Biochimica et Biophysica Acta (BBA) - Lipids and Lipid. Metabolism, 1124(2), 123–134. Görgün, S., & Akpınar, M. A. (2012). Purification and characterization of lipase from the liver of carp, Cyprinus carpio L. (1758), living in lake Tödürge (Sivas, Türkiye). Turkish Journal of Fisheries and Aquatic Sciences, 12, 207–215. Haard, N. F. (1998). Specialty enzymes from marine organisms. Food Technology, 53(7), 64–67. Iijima, N., Tanaka, S., & Ota, Y. (1998). Purification and characterization of bile saltactivated lipase from the hepatopancreas of red sea bream, Pagrus major. Fish Physiology and Biochemistry, 18(1), 59–69. Kameshwar Sharma, Y. V. R., Boora, N., & Tyagi, P. (2014). Isolation, purification and characterization of secondary structure and kinetic study of lipase from Indian major

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