Partial characterization of pyloric-duodenal lipaseof gilthead seabream (Sparus aurata)

Hector Nolasco • Francisco Moyano-Lopez •

Fernando Vega-Villasante

Received: 11 August 2009 / Accepted: 16 June 2010

� Springer Science+Business Media B.V. 2010

Abstract In the present study, we report the

isolation and characterization of seabream Sparus

aurata pyloric caeca-duodenal lipase. Optimum

activity was found at pH 8.5 and salinity of 50 mM

NaCl. Lipase activity was sensitive to divalent ions,

and extreme pH values (4, 5, and 12), being more

stable at alkaline than acid pH. Optimum temperature

was found at 50�C, but lipase was stable at temper-

atures below 40�C. Lipase has a bile salt sodium

taurocholate requirement for increased activity. Gra-

dient PAGE electrophoresis revealed the presence of

four isoforms with apparent molecular masses of 34,

50, 68, and 84 KDa, respectively. Pyloric-duodenal

lipase was able to hydrolyze emulsified alimentary

oils. Results confirm the presence of true lipases in

Sparus aurata digestive tract.

Keywords Digestive enzymes �Fish digestive physiology � Fish nutrition �Lipase � Seabream � Sparus aurata

Introduction

The gilthead seabream Sparus aurata is an important

species in the mediterranean finfish aquaculture

(Fernandez et al. 2001; Cara et al. 2003; Deguara

et al. 2003; Venou et al. 2009). As in most cultured

species, protein is the major ingredient in its feeds,

then, a number of studies have been focussed to the

characterization of its digestive proteases (Moyano

et al. 1998) and to the evaluation of their use within in

vitro digestibility assays (Moyano and Savoie 2001).

Nevertheless, although feeds routinely used in

ongrowing of this species contain as much as 20%

fat, the main functional aspects of its digestive lipases

have not been similarly studied. The evaluation of

digestive lipases has been carried out in different fish

species like Pagrus major (Iijima et al. 1998),

Pseudoplatystoma corruscans (Lundstedt et al.

2004), Oreochromis spp. (Jun-Sheng et al. 2006),

Thunnus orintalis (Matus de la Parra et al. 2007),

Oncorhynchus tshawytscha and Macruronus novaez-

elandiae (Kurtovic et al. 2010), or Glyptosternum

maculatum (Xiong et al. 2010. Other aspects, like its

changes with larval development (Izquierdo and

Henderson 1998; Cahu et al. 2000), effect of diet on

H. Nolasco (&)

Centro de Investigaciones Biologicas del Noroeste, S.C.,

Mar Bermejo No. 195, Col. Playa Palo Santa Rita,

23000 La Paz, BCS, Mexico

e-mail: hnolasco04@cibnor.mx

F. Moyano-Lopez

Universidad de Almerıa, Carr. a Sacramento sn,

04120 La Canada de San Urbano, Almerıa, Spain

F. Vega-Villasante

Centro Universitario de la Costa, Universidad de

Guadalajara, Puerto Vallarta, Jalisco, Mexico

123

Fish Physiol Biochem

DOI 10.1007/s10695-010-9414-7

lipase activity (Debnath et al. 2007; Hansen et al.

2008; Chatzifotis et al. 2008), or the effect of

supplementary lipase on growth and body composition

(Samuelsen et al. 2001) have been also assessed.

Although a comprehensive revision on their main

functional features has been recently carried out by

Kurtovic et al. (2009), studies focused to the charac-

terization of fish lipases are scarce (Gjellesvik et al.

1989; Iijima et al. 1997; Taniguchi et al. 2001; Degerli

and Akpinar 2002). Specific reports on Sparus aurata

lipases include only the study of the insulin regulation

of serum lipoprotein lipase (LPL) activity and expres-

sion (Albalat et al. 2007). Taking this into account,

partial characterization of lipase of Sparus aurata was

considered an interesting objective for a better under-

standing of its digestive physiology, as well as a tool

for further development of in vitro assays for lipid

digestibility in this species.

Methods

Preparation of pyloric-duodenal lipase extract

Thirty live specimens of seabream (Sparus aurata)

ranging from 250 to 300 g were provided by a local

farm (Piagua S.L. Almerıa Spain). Fish were routinely

fed on a commercial diet (45% protein), three times

per day (09:00, 14:00, and 19:00 h) to reach a total

amount of feed representing 3% of the body weight.

Prior to sampling, fish were starved for 12 h, then

killed by submersion in ice-cold water (15 min at

2�C). The digestive tract was dissected to separate the

pyloric caeca and anterior duodenal portion. Samples

of this pyloric caeca-duodenal tract were manually

homogenized (potter Eveljhem) with ice-cold water

(1:3 w/v). Isolated lipase extracts, obtained after

centrifugation at 15,680g, 4�C, 10 min (EBA 12R,

Hettich Zentrigugen, Tuttlingen), were stored at

-20�C, and further utilized for enzyme analysis.

Concentration of soluble protein in the lipase extracts

was determined by the method of Bradford (1976).

Lipase activity

Lipase activity was evaluated following the method

described by Versaw et al. (1989), with some

modifications. The detailed procedure was as follows:

10 lL of the enzyme preparation was mixed with

100 lL of taurocholic acid sodium salt hydrate

(100 mM) (Sigma T4009), 920 lL of Tris–HCl

buffer (50 mM, pH 8) (Amresco, Solon, Ohio, USA,

0497). Reaction was initiated by the addition of 10

lL of b-naphthyl caprylate (100 mM, in dimethyl-

sulfoxide, DMSO) (Sigma N-8875) and incubated at

25�C for 10 min, then 10 lL of Fast Blue BB salt

(FB) (100 mM in DMSO) (Sigma F-3378) were

added, just before the reaction was stopped by 100

lL of trichloroacetic acid (TCA) (0.72 N). The

mixture were clarified with 1,350 lL of ethyl

acetate–ethanol (1:1) (Panreac, Barcelona, Spain,

141086.1214, and 321318.1612, respectively), and

the absorbance was read at 520 nm, according to the

absorption spectrum of the coloured reaction mixture

(Ultrospec 3330, Amersham Pharmacia Biotech,

Uppsala, Sweden). A standard curve was prepared

by replacing b-naphthyl caprylate by varying con-

centrations of b-naphthol (Sigma N-1250) dissolved

in DMSO. One unit of activity was defined as the

amount of enzyme required to produce 1 lmol of

b-naphthol per minute.

Optimal pH and temperature ranges

Optimal pH for lipase activity was determined using

Universal Buffer (Stauffer 1989) ranging from 4 to

12. For all treatments, the pH of reaction mixture was

adjusted to values between 8 and 10, just before Fast

Blue reagent was added, to avoid effect of extreme

pH (mainly acid) on colour development.

The effect of pH on stability was determined by

preincubation of lipase extracts at different pH for

3 h, and sampled at 0, 30, 60, 120 and 180 min, to

assay for residual lipase activity at pH 8.

Optimal temperature for lipase activity was deter-

mined by incubating at temperatures ranging from 10

to 80�C at pH 8.0.

The effect of temperature on stability was deter-

mined by preincubation of lipase extracts at different

temperatures ranging from 30 to 60�C for 180 min,

and sampled at 0, 30, 60, 120 and 180 min, respec-

tively, to assay for residual lipase activity at 25�C.

The dual effect of pH and temperature, within

physiological ranges (pH from 6 to 9,5; temperature

from 10 to 35�C), on the activity of the enzyme was

evaluated in a 9 9 6 experimental design.

Fish Physiol Biochem

123

Effect of salinity and metal ions

Optimal salinity for lipase activity was determined

using NaCl ranging from 0 to 1.5 M. The effect of

some divalent ions in the activity (Ca2?, Mg2?,

Mn,2?, Fe2?, Co2?, Cu2?, Hg2?, Pb2? and Zn)2?, all

of them in the form of chloride salts, was evaluated in

the range from 0 to 20 mM, with the exception of

Fe2? and Pb2? (range from 0 to 5 mM).

Bile salt requirements

Optimal bile salt concentration for lipase activity was

determined using sodium taurocholate (Sigma

T4009-5G) ranging from 0 to 40 mM final concen-

tration in the reaction mixture.

Lipase zymograms

Lipase zymograms were obtained using a 4–30%

native gradient PAGE where b-naphthyl caprylate

(200 mM) was copolymerized as described by Alva-

rez-Gonzalez et al. (2008). The gels were equilibrated

for 15 min at 80 V, after the voltage was set to 120 V

for 4.5 h at 4�C. Lipase activity was revealed with the

addition of a Fast Blue (Sigma, D-9805) solution

(100 mM). Purplish bands in the electrophoresis gel,

formed by b-naphthol-Fast Blue-coloured complex

revealed activity of lipases after 10–15 min. There-

after, gel was washed with distilled water prior to

protein staining for 2 h with 0.1% Coomassie Blue

(BBC R-250) in a methanol–acetic acid–water solu-

tion (40:10:50, v/v). Blue bands in the electrophoresis

gel, formed by protein-Coomassie Blue-coloured

complex, were revealed. Destaining was carried out

over night in a methanol–acetic acid–water solution

(40:10:50, v/v).

Rf and molecular mass calculation

A medium-range molecular mass marker (MWM GE

170446.01, GE Healthcare Home, UK LTD, Eng-

land) was applied to each 4–30% gradient PAGE at 5

lL per well. The MWM kit contained the following

protein markers: phosphorylase b (97 KDa), bovine

serum albumin (66 KDa), egg albumin (45 KDa),

carbonic anhydrase (30 KDa), soybean trypsin inhib-

itor (20.1 KDa), and a-lactoalbumin (14.4 KDa). The

relative electromobility (Rf) was calculated for all

bands in the zymogram, and the molecular mass of

each band in the gradient PAGE zymogram was

calculated by a linearly adjusted model between the

Rf and the decimal logarithm of molecular mass

proteins in the molecular marker using the software

program EXCEL 2007 (Microsoft, USA).

Hydrolysis of natural substrates

The hydrolysis of olive oil by S. aurata lipase was

assayed by the pH-drop method carried out within the

more suitable pH range for enzyme stability. The

emulsion was prepared as indicated by Nolasco

(2008), being the pH registered every 300 s, during

60-min incubation at 25�C, and 300 rpm (GLP 21,

Crison, Barcelona, Spain). Oil hydrolysis rate was

calculated as the slope of pH drop in comparison with

that obtained in a control experiment performed using

the same amount of a heat-inactivated lipase extract.

Results

Lipase activity in extracts measured at neutral pH

was 7.790 ± 0.003 U/mL, its specific activity being

1.08 U/mg. The activity of S. aurata lipase at

different pH is shown in Fig. 1. The differences in

the profile resulting from making or not a pH

adjustment prior to the addition of Fast Blue in the

reaction mixture were evident in the acidic zone.

Stability of the enzyme is shown in Fig. 2; a higher

0

20

40

60

80

100

3 4 5 6 7 8 9 10 11

pH

Rel

ativ

e ac

tivi

ty (

%)

NA

Fig. 1 Optimal pH for lipase activity. Effect of pH was

determined using universal buffer ranging from 4 to 11, instead

Tris–HCl buffer, according to the indicated method. Dash lineindicated that reaction mixture was adjusted to pH between 8

and 10 before colour development with FB reagent. Maximum

activity at pH 9 was considered as 100%

Fish Physiol Biochem

123

stability at alkaline pH (more stable at pH 8) was

evidenced. The effect of temperature on the activity

and stability of the enzyme is shown in Figs. 3 and 4,

respectively. Although maximum activity was

detected at 50�C, the stability of the enzyme was

lower at temperatures above 40�C (60, 20, and 10%

at 40, 50, and 60�C after 2 h treatment, respectively).

Results obtained after evaluation of the activity

within a physiological range of both pH and temper-

ature are shown in Fig. 5. Extreme pH values (6 and

9.5), and temperatures (10, and 40�C), negatively

affect the activity.

Optimal salinity for Sparus aurata lipase activity

is shown in Fig. 6. Lipase showed a low requirement

for Na?, since maximum activity was achieved at

50 mM of salt concentration in the reaction mixture.

The increase in salt concentration resulted in a

progressive decrease in the activity, with values

accounting for 83, 65, and 57% of total activity

measured at 0.5, 0.75, and 1 M, respectively. On the

other hand, different responses to the increased

concentration of divalent ions in the reaction mixture

were found (Fig. 7). Some ions like Ca2? or Mg2?

produced a non-significant effect on the activity until

reaching quite high concentrations (10 mM). In

contrast, other ions like Pb2?, Fe2?, Zn2?, or Hg2?

produced a negative effect, decreasing the activity

below 20% at concentrations of 5 mM or even 1 mM.

The effect of Mn2? or Co2? was intermediate

between the two described responses.

Optimal concentration of taurocholate for the

activity of S. aurata lipase is shown in Fig. 8. A

two-peak profile was obtained, showing a clear

activation at low concentration (0.1 mM) but also a

non-significative slight and secondary increase at a

higher concentration (20–30 mM).

Fig. 2 pH effect on lipase stability. The effect of pH on

stability was determined by preincubation of lipase extracts at

pH ranging from 4 to 12 for 180 min, and sampled at 0, 30, 60,

120 and 180 min, respectively, to assay for residual lipase

activity at pH 8, following the indicated method. Average of

initial activity at pH 7, 8, and 9 was considered as 100%

0

5

10

15

20

25

30

0 10 20 30 40 50 60 70 80

Temperature ( οC)

U/m

L

Fig. 3 Optimal temperature for lipase activity. Optimal

temperature for lipase activity was determined incubating at

temperatures ranging from 10 to 80�C, following the indicated

method. Initial activity at 30�C was considered as 100%

0

20

40

60

80

100

120

140

0 20 40 60 80 100 120 140 160 180

Exposure time (min)

Res

idu

al a

ctiv

ity

(%)

30405060

Fig. 4 Temperature effect on lipase stability. The effect of

temperature on stability was determined by preincubation of

lipase extracts at different temperatures ranging from 30 to

60�C for 180 min, and sampled at 0, 30, 60, 120 and 180 min,

respectively, to assay for residual lipase activity at 25�C,

following the indicated method

Fish Physiol Biochem

123

The visualization of lipase activity by the hydro-

lysis of b-naphtyl caprylate in zymogram is shown in

Fig. 9a. At least four isoforms, with molecular

masses of 34, 50, 68, and 84 KDa were revealed by

the PAGE (Fig. 9b) at the established conditions.

The ability of S. aurata lipase to hydrolyse a

natural substrate, like triglycerides present in olive

oil, was evidenced by results obtained in the pH-drop

assay (Fig. 10).

Discussion

The pyloric-duodenal extract of Sparus aurata pre-

sented lipase activity (1.08 U/mg), this being in

agreement with different authors who reported that

pyloric caecae and upper intestine are the regions

with the higher activity of digestive alkaline enzymes

6 6,5 7 7,5 8 8,5 9 9,5 1010

25

350

5

10

15

20

25

U/m

L

pH

T(o

C)

20-25

15-20

10-15

5-10

0-5

a

b

Fig. 5 Dual effect of pH and temperature on lipase activity. The

dual effect of pH (a) and temperature (b), at pH ranging from 6 to

9,5; and temperature ranging from 10 to 35�C, on the activity of

the lipases was evaluated, following the indicated method

40

60

80

100

0 0,5 1 1,5

NaCl (M)

Rea

ltiv

e ac

tivi

ty (

%)

Fig. 6 Optimal sodium chloride concentration for lipase

activity. Optimal salinity for lipase activity was determined

using NaCl final concentration, ranging from 0 to 1.5 M,

following the indicated method. Activity without additional

NaCl was considered as 100%

Fig. 7 Effect of divalent cations on lipase activity. The effect

of divalent ions in the activity (chloride salts of Ca2?, Mg2?,

Mn2?, Fe2?, Co2?, Cu2?, Hg2?, Pb2? and Zn2?) was evaluated

in the range from 0 to 20 mM (final concentration in reaction

mixture), with the exception of Fe2? and Pb2? (range from 0 to

5 mM), following the indicated method. Activity without

additional divalent ions was considered as 100%

Fish Physiol Biochem

123

(proteases, amylases, and lipases) in fish (Deguara

et al. 2003; Jun-Sheng et al. 2006; Matus de la Parra

et al. 2007; Xiong et al. 2010). This contrasts to

results obtained in other species like the turbot

(Scophtalmus maximus), on which the highest lipo-

lytic activity was measured at hindgut and rectum

(Koven et al. 1997). Such differences may be related

to the presence in this latter species of only a

rudimentary pair of pyloric caeca which are unlikely

to play a major role in lipid digestion, which in

contrast is carried out to a greater extent by the

intestinal microflora.

0

2

4

6

8

10

12

14

0 10 20 30 40

Sodium tauracholate (mM)

U/m

L

0

5

10

15

0 0,1 0,2 0,3 0,4 0,5 0,6 0,7 0,8 0,9 1

Sodium tauracholate (mM)

U/m

L

Fig. 8 Optimal sodium taurocholate concentration for lipase

activity. Optimal bile salt concentration for lipase activity was

determined using sodium taurocholate ranging from 0 to

40 mM final concentration in the reaction mixture, following

the indicated method. Activity without sodium taurocholate

was considered as 100%

Fig. 9 a Electrophoregram of pyloric-duodenal extract con-

taining lipases. Row 1: MWM, row 2: 10 lL of lipase extract,

row 3: 20 lL of lipase extract, row 4: 10 lL of lipase extract

diluted 1:10, row 5: 20 lL of lipase extract diluted 1:10.

b Zymogram, using b-naphtyl caprylate as substrate, of

pyloric-duodenal lipases in gradient native PAGE. Row 1:

MWM, row 2: 10 lL of lipase extract, row 3: 20 lL of lipase

extract, row 4: 10 lL of lipase extract diluted 1:10, row 5: 20

lL of lipase extract diluted 1:10, row 6: 10 lL of lipase extract

diluted 1:20, row 7: 20 lL of lipase extract diluted 1:20, and

row 8: 10 lL lipase extract diluted 1:50

y = -0,0031xR2 = 0,9986

y = -0,0062xR2 = 0,9952

-0,16

-0,14

-0,12

-0,10

-0,08

-0,06

-0,04

-0,02

0,00

0,02

0 10 20

Incubation time (min)

pH

dro

p (

un

its)

Fig. 10 Olive oil hydrolysis by Sparus aurata lipase. L lipase

treatment, C control treatment. Oil hydrolysis is expressed as

the rate of pH change

Fish Physiol Biochem

123

Optimal pH for the lipase activity in S. aurata was

found to be close to 9, being in agreement with the

optimal pH reported for other intestinal enzymes in

this species (Munilla-Moran and Saborido-Rey

1996a, b). Similar values have been described for

lipases in other fish species like the rainbow trout

(Oncorhynchus mykiss), cod (Gadus morhua), red sea

bream (Pagrus major), Pacific blue tuna (Thunnus

orientalis), grey mullet (Liza parsia), Chinook

salmon (Oncorhynchus tshawytscha) and hoki (Ma-

cruronus novaezelandiae), or carnivorous teleost fish

of Tibet (Glyptosternum maculatum) (Metin and

Akpinar 2000; Tocher and Sargent 1984; Gjellesvik

et al. 1989; Iijima et al. 1998; Matus de la Parra et al.

2007; Islam et al. 2008; Kurtovic et al. 2010; Xiong

et al. 2010, respectively), while maximum activity at

a more neutral pH has been reported in species like

Cyprinion macrostomus or the tilapia (Taniguchi

et al. 2001; Degerli and Akpinar 2002; Jun-Sheng

et al. 2006). The lipase activity was highly stable at

pH 7–8, this being in agreement with the pH range

found for many fish intestinal digestive enzymes

(Iijima et al. 1997; Ugolev and Kuz0mina 1993, in

Kuzmina and Ushakova 2007), as well to the normal

pH values measured in the intestine of this species

(Deguara et al. 2003). Nevertheless, while a marked

reduction in activity at acidic pH has been described

for lipases in other species like the oil sardine

(Sardinella longiceps) or the grey mullet (Liza

parsia) (Mukundan et al. 1985; Islam et al. 2008),

sea bream lipase showed a great stability at a more

acid pH, retaining an important amount of activity

after 3-h incubation at pH 6. This could be related to

the presence of a highly functional stomach in this

species, since under the physiological conditions

existing in the live fish, and in spite of the secretion

of bicarbonate into the duodenum, the continuous

flow of acid digesta coming from the stomach to the

proximal intestine determines that optimal pH for the

in vitro activity of the enzyme is rarely reached.

Hence, a low sensitivity to acid pH could be

interpreted as a functional adaptation to perform

lipid hydrolysis even under non-optimal conditions.

An equivalent adaptation has been described by

Borlongan (1990), who described the presence of

intestinal and pancreatic lipases in the milkfish

Chanos chanos, showing maximum activities at pH

6.8 and 8.0, respectively. According to this author,

the detection of two well-defined pHs for both the

intestinal and pancreatic lipases suggests a physio-

logical versatility for lipid digestion in this species.

Maximum activity of lipase was measured at

50�C; similarly to what reported for Thunnus orien-

talis lipase (Matus de la Parra et al. 2007), but higher

than the range of 25–40�C reported for hybrid

juvenile tilapia (Oreochromis niloticus 9 Oreochr-

omis aureus) by Jun-Sheng et al. (2006). Neverthe-

less, the observed stability at this temperature was

low. Although a great number of studies oriented to

the functional characterization of digestive enzymes

present similar results (Munilla-Moran and Saborido-

Rey 1996a; Alarcon et al. 1998; Klomklao et al.

2006), it must be concluded that such data offer very

limited information about functionality of the

enzymes under physiological conditions. In this

sense, the measurement of a given activity within a

physiological range of both pH and temperature (as

detailed in Fig. 5) provides a more valuable insight

into potential differences in activity in the live fish.

These data are also needed for a more suitable

simulation of the digestion under in vitro conditions.

Tested extreme pH values (6 and 9.5), and temper-

atures (10, and 40�C), negatively affect lipase

activity, however, at physiological values of pH

(6–8, according with Deguara et al. 2003) and

temperatures (10–30�C) lipase is considerably active.

As Ugolev and Kuz0mina reported in 1993 (Kuzmina

and Ushakova 2007), within the range of tempera-

tures typical for fish life, the strongest effects on

digestive enzymes (protease activity) were found

when temperature dropped down towards 0�C, also

pH increase leads to increase intestinal proteinase

activity in the studied fish, similarly as found in

Sparus aurata lipase.

No information about the effect of salinity on fish

lipases has been found; however, most of the methods

used for determination of lipase activity in fishes

include about 100 mM or lower NaCl concentration

(Iijima et al. 1997; Uchiyama et al. 2002; Murray

et al. 2003; Perez-Casanova et al. 2004; Albalat et al.

2007; Matus de la Parra et al. 2007; Chatzifotis et al.

2008). None of those authors justify the routine use of

NaCl in the lipase assay, although it could be

explained as a way to simulate the high concentration

of salt in the sea water which is continuously ingested

by marine fish. Evaluation of the optimal NaCl

concentration for lipase activity is neither reported in

such studies, this being an important feature since, as

Fish Physiol Biochem

123

it was found in the present study for S. aurata lipase,

a great sensitivity to concentrations over 50 mM

could affect activity determinations.

The lipase of S. aurata showed not to be activated

by Ca2?, in contrast to what described for bovine or

porcine pancreatic lipases (Khan et al. 1975; Alvarez

and Stella 1989), human lipoprotein lipase (Zhang

et al. 2005), or lipases of Pagrus major (Iijima et al.

1997) Chinook salmon (Oncorhynchus tshawytscha)

and hoki (Macruronus novaezelandiae) (Kurtovic

et al. 2010). In relation to the effect of different metal

ions, in spite of their clear negative effect on the S.

aurata lipase, this seems to be more resistant than

other similar enzymes, like the phospholipase of the

red sea bream Pagrus major which retained only 38,

39, and 0.5% of activity after exposure to 5 mM

chloride salts of Mg2?, Cu2?, and Zn2?, or the lipases

of Oreochromis niloticus, and grey mullet which

were highly inhibited by heavy metals like Cd2?,

Zn2?, and Hg2? (Taniguchi et al. 2001; Islam et al.

2008). Those latter authors suggest that the presence

of SH groups close to the catalytic site of the enzyme

may be affected by such divalent ions, as it was also

demonstrated for dogfish lipase (Raso and Hultin

1988). Hg2? has been shown to inhibit the rat lipase

(Fredrikson et al. 1981). From an applied point of

view, the negative effect of metals on digestive

enzymes must be a factor to be taken under consid-

eration when selecting suitable zones to place aqua-

culture facilities, since pollution produced by such

ions may reduce digestive efficiency of the cultured

fish.

Bile acids or their conjugated forms (bile salts) are

secreted by the gallbladder of vertebrates as a result

of the oxidation of cholesterol. They collaborate in

the action of lipases acting as emulsifiers of their

substrates, this role being particularly important when

the diet includes a high amount of long-chain fatty

acids. Although bile salts are recognized as factors

affecting the functionality of fish lipases (Iijima et al.

1998), very few papers have studied the composition

or their role in the digestion of cultured fish. From

such studies, it is deduced that teleosts have princi-

pally taurocholate and taurochenodeoxycholate (the

taurine derivative) in their bile salts (Une et al. 1991;

Alam et al. 2001). In fact, classification of fish lipases

recognized such a strong dependence by identifying

some activities reported in different species as ‘‘bile

salt-dependent lipases’’ (BSDL). The reported type

and relative concentration of bile salts affecting

lipase activity lies within a wide range. Cod lipase

requires 2–10 mM sodium taurocholate for activity,

presenting a higher bile salt requirement for hydro-

lysis of olive oil than for hydrolysis of tributyrin

(Gjellesvik et al. 1989). Pagrus major lipase requires

about 20 mM sodium taurocholate or cholate, but

was inhibited by sodium deoxycholate (Iijima et al.

1998). In our results, higher lipase activity was

recorded at a concentration of 0.1 mM sodium

taurocholate, being reduced by higher concentrations.

This in agreement with results reported in other

species like the Pacific bluefin tuna Thunnus orien-

talis, which lipase showed no requirement for bile

salts, even being 60% inhibited by 6 mM of sodium

taurocholate or by a mixture of sodium cholate–

deoxicholate or by natural bile salts extracted from

gallbladder (Matus de la Parra et al. 2007).

Our results, obtained under the conditions of a

native gradient electrophoresis, showed two main

lipase isoforms with 34 and 68 kDa and another two

with 50 and 84 KDa. Accordingly, to such molecular

masses, it is suggested that the lipase of S. aurata

may be constituted by only two peptides of 34 and 50

KDa, which can be combined in other different

structural quaternary associations (34 ? 34) and

(34 ? 50). Consistent with our results, Degerli and

Akpinar (2002) reported that the purified intestinal

lipase from Cyprinion macrostomus had a molecular

mass of 51 KDa. Red sea bream Pagrus major lipase

showed a mass of 64 KDa (Iijima et al. 1998); also

Iijima et al. (1997) reported a small phospholipase

with a molecular mass of 14 KDa from Pagrus major

pyloric caeca. Consistent as well are the lipase

molecular mass found by Degerli and Akpinar (2002)

in Cyprinion macrostomus (51 KDa), the two lipases

from dorsal part of grey mullet, purified by Islam

et al. (2008), with about 46.4 and 41.2 KDa,

respectively, salmon lipase (79.6 and 54.9 kDa),

and hoki lipase (44.6 kDa) (Kurtovic et al. 2010).

The hydrolysis of triglycerides of long-chain fatty

acids, like those present in olive oil, by fish lipases

has been assessed in different species like cod (Gadus

morhua) (Gjellesvik et al. 1989) and Cyprinion

macrostomus (Degerli and Akpinar 2002). Our results

demonstrate that S. aurata lipase is able to hydrolyze

this complex substrate. The general trend to increase

lipid content in fish diets, as a way to increase energy

content, and to substitute fish oil by vegetable oils,

Fish Physiol Biochem

123

requires additional studies that could benefit from in

vitro digestibility assays performed using lipases

from cultivated species. The results obtained in the

present study, describing some of the main opera-

tional parameters to develop such activity in S. aura-

ta, are a first step towards such direction.

Acknowledgments The technical assistance of Antonia

Barros, Mariam Hamdam, and Patricia Hinojosa is

acknowledged. HN thanks the Government of Mexico, through

CONACYT by for grant 000000000081074, and for financial of

the CONACYT research project No. 0000000000084652 related

to lipid digestibility, and to Universidad de Almeria for

acceptance for research stay.

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