Abstract: Two lipases designated as Lip-1 and Lip-2 were purified from dorsal part of grey mullet (Liza parsia) to homogeneity by 85% (NH4)2SO4 fractionation followed by simultaneous desalting and concentration by ultrafiltration and Sephadex G-50 and DEAE-cellulose chromotography and CM-cellulose chromotography. The molecular weight of two lipases was determined by SDS-PAGE and gel filtration about 46.5 and 41.2 KDa, respectively. Both the enzymes were dimer in nature remained unchanged the presence and absence of reducing agent under SDA-PAGE. The Lip-1 and Lip-2 lipases were active within the pH range of 7-8.5, with an optimum pH of 8 and 8.5.0 and were stable from 2.0-10.5. The enzyme was active within the temperature range of 30-60°C and maximum activities were observed around 33 and 35°C, respectively and beyond which it lost activity progressively. The hydrolytic activity was enhanced by Ca+ and EDTA (concentration 0.001-0.003 M) but strongly inhibited by heavy metal s Cd++, Zn++ and Hg++. The presence of Zn++ and Hg++ potently inhibited lipolytic activities of the lipases from grey mullet, while activities were slightly inhibited in the presence of Cu++ salts.
INTRODUCTION
Lipases are hydrolytic enzymes (EC 3.1.13) act at fat-water interfaces to catalyze the hydrolysis of tryglycerides to fatty acid and glycerol (Sarda and Desnuelles, 1958, Desnuelle et al., 1960; Norin et al., 1993). Lipases are very important because of the role they play in the postmortem quality deterioration of seafood and other foodstuffs during handling, chilled frozen storage and widely used for biotechnological applications in dairy industry, oil processing, production of surfactants and preparation of enantiomercally pure pharmaceuticals. Compared with other hydrolytic enzymes (e.g., proteases and carbohydrases), lipases are relatively less well studied and in this regard, lipases from aquatic animals are even less well known versus their counterparts from mammalian, plant and microbial sources (L`opez-Amaya et al., 2001). The aquatic world contains a wide variety of living species and, hence represents great potential for discovering new enzymes (Cherif et al., 2007). the presence of a lipase activity has been described for some aquatic organisms such as lobster (Brockerhoff and Stewart, 1967), crab (Vonk, 1960) and Homarus americanus (Brockerhoff et al., 1970) and few lipases that have been studied from fish and other aquatic animals include lipases from the leopard shark (Patton et al., 1977) rainbow trout (Tocher and Sargent, 1984), Atlantic cod (Lie and Lambersten, 1985; Gjellesvik et al., 1992), dog fish (Raso and Hultin, 1988), sardine (Mukundan et al., 1985), anchovy, striped bass and salmon (Leger et al., 1977), as well as red sea bream (Iijima et al., 1998). Other investigators have been reported lipases and lipolytic activities in the digestive glands, liver and adipose tissues of different fish species (Izquierdo and Socorro, 2000; Liang et al., 2002).
From few years ago, considerable attention has been devoted to lipases of botanical origin and microorganisms (Aquino, 1994; Desnuelle, 1961). Several lipases of animal origin have been purified and their chemical and enzymatic properties were elucidated in details (Verger, 1984; Mc Crae, 1983). Lipase can hydrolyze lipids and produce undesirable rancid flavor in milk products, meat, fish and other food products containing fat. For instance lipases have been used extensively in the dairy industry for house hold detergent in the oleochemical industry and to produce structural triglycerides (Verger et al., 1982). Lipase also used in the synthesis of polymers, agrochemical leather textile, baking pharmaceutical and paper industry.
Most of the purification procedures for lipases reported involve series of non-specific techniques, such as ammonium sulphate precipitation, gel filtration and ion-exchange chromatography. In recent years, affinity chromatographic techniques have become frequently used, decreasing the number of steps necessary for lipase purification.
Most recently, reversed-micellar and aqueous two phase systems, ultra filtration membranes and immune purification have also been applied to purify some lipases, mainly of microbial origin. In the present study purification of grey mullet lipases was carried out mainly by ion-exchange chromatography followed by gel filtration.
MATERIALS AND METHODS
Biological Materials
Adult grey mullet (1-1.5 kg) were purchased from local fish landing
centre and the dorsal part was removed with knife and stored in crushed
ice in plastic bags and transported directly to the laboratory where were
stored frozen at -20°C until used for experimental purpose.
Chemicals and Reagents
Sephadex G-50 and deae-cellulose and cm-cellulose were purchased from
pharmacia Biotech Inc. Sweden. Diisopropylfluorophosphate (DIP) was from
ICN and phenylmethyl sulphonylfluoride (PMSF) was from Wako pure chemical
Industry Ltd, Japan, Sodium cholate (NaC), sodium taurocholate (NaTC),
sodium deoxycholate (NaDC), dioxane, arabic gum, glycerol, benzamidine
hydrochloride, calcium chloride (CaCl2), Bovine Serum Albumin
(BSA), sodium hydroxide (NaOH), ammonium sulphate ((NH4)2SO4),
ethylenediamine tetraacetic acid (EDTA), tris(hydroxymethyl) aminomethane
(Tris), mercaptoethanol, Sodium Dodecyl Sulfate (SDS) and mercury chloride
(HgCl2) were purchased from Fisher Scientific (Fair Lawn, NJ,
USA). Acetone, chloroform, butanol, methanol, diethyl ether, iso-propanol,
hydrochloric acid, glacial acetic acid, benzene, toluene, hexane, n-heptane,
iso-octane, dimethyl sulfoxide, dimethyl formamide (HPLC/analytical grade)
were also purchased from Fisher Scientific (Fair Lawn, NJ, USA).
Sample Preparation for Enzyme Extraction
The frozen dorsal part of fish were thawed at 4°C and cleaned
by flushing with distilled water followed by thorough rinsing with ice-cold
0.85% NaCl solution to remove adhered blood. The dorsal part of fish were
chopped into small pieces and rapidly frozen in liquid nitrogen and then
comminuted into fine powder in a Waring blender. The powder was defatted
using a modified version of the procedure by Verger et al. (1982)
and Andrews (1965), with successive changes of cold acetone, chloroform:
n-butanol (9:1, v/v), chloroform: n-butanol (8:2, v/v), acetone and diethyl
ether, all at -20°C with intermittent stirring then filtration. The
ratio of tissue to solvent was 1:10 (w/v) and after each solvent treatment,
the homogenate was filtered via vacuum suction with a B¨uchner funnel.
The defatted material was air dried at room temperature (23°C) in
a fume hood then stored at 20°C until needed.
Enzyme Extract Preparation
The dried defatted powder was homogenized in 25 mM Tris-HCl buffer,
pH 7.8, containing 5 mM benzamidine-HCl, 1mM EDTA and 10% (w/v) glycerol
(TBEG buffer). The defatted powder to TBEG buffer ratio was (1:10, w/v);
the homogenate was gently stirred at 4°C for 1 h and then centrifuged
at 8000xg for 20 min at 4°C. The precipitate was discarded and the
supernatant was filtered through several layers of cheesecloth (to remove
the floating fatty material) and then fractionated with solid (NH4) 2SO4
up to 85% saturation. The mixture was gently stirred for 3 h at 4°C
and re-centrifuged at 8000xg for 20 min at 4°C. The resulting precipitate
was collected and redissolved in TBEG buffer (pH 7.8) to form the ammonium
sulfate fraction. The fraction was dialyzed overnight against three changes
of 4 lTBEG buffer (pH 7.8) in a cellulose membrane dialysis tubing (12
kDa MW. CO, Sigma Chemical Co., St. Louis, MO, USA). The dialyzed fraction
was centrifuged again at 8000xg for 20 min at 4°C and the resulting
dialysate was simultaneously desalted and concentrated using a Millipore
Amicon® Ultra centrifugal filter device (30 kDa MWCO, Amicon Co. Ltd.,
Bedford, MA, USA). The concentrated and desalted extract, ultrafiltrate
(UF fraction) was stored frozen at 20°C until needed.
Isolation and Purification of Lipases
The enzyme extract was loaded into the DEAE-cellulose column previously
equilibrated with Tris-HCl buffer, pH 8.4 for 24 h and the proteins were
recovered from the column by step wise elution with the same buffer containing
different concentration of NaCl at 4°C. The fraction showing the lipase
activity was pooled and dialyzed against 10 mM Tris-HCl buffer, pH 8.4
for overnight with four changes of buffer at 4°C. After centrifugation
the supernatant was applied into a CM-cellulose column previously equilibrium
with 5 mM PBS, pH 6.5 and the proteins were eluted from the column with
the same buffer. The fraction showing the lipase activity were pooled
and dialyzed again against 10 mM Tris-HCl buffer, pH 8.4 for overnight
with four changes of buffer at 4°C. After centrifugation the supernatant
was applied into gel filtration on Sephadex G-50 which was previously
equilibrium with 5 mM PBS, pH 7.6 at 4°C and the proteins were eluted
from the column with the same buffer. Lipase containing active fraction
was pooled and tested for homogeneity.
Electrophoresis
Polyacrylamide disc gel electrophoresis was conducted at room temperature,
pH 8.4 on 7.5% gel and amido black was used as staining reagent.
Determination of Molecular Weight (MW): MW of the purified native enzyme was determined by gel filtration on Sephadex-50 column (0.9x90 cm) as described by Laemmli (1970). The marker proteins used were lysozyme (14.6 KD), Egg albumin (4.5 KD), BSA (66 KDS), α galactosidase (116 KDs), β-amylase and tripsinogen (24 KD). The MW also determined by SDS-PAGE according to the method of Laemmli (1970) and Sugihara et al. (1990) using the marker proteins, Myosine (205 KD) α galactosidase (116 KDs), BSA (66 KDS), Carbonia anhydrase (29 KD), β lactoalbumin (18 KD) and Aprotinin (6.5 KD)
Enzyme Assay
Lipase activity was assayed as reported (Sugihara et al., 1990)
using olive oil as substrate. The lipase activity was measured by estimating
the release of free fatty acids. One unit of lipase activity is defined
as the amount that liberates one micromole of fatty acid under the specified
conditions. Specific activity of lipase is expressed as the enzyme unit
per mg of protein.
Estimation of Protein
Protein concentration was estimated by the method by Lowry et al.
(1951) using BSA as standard as well as by measurement of absorbance at
280 nm.
Effect of pH on the Activity and Stability of Lipases
The activity of two lipase denoted as Lip-1 and Lip-2 was investigated
within the pH range of 2.0-8.0 using the following buffer solutions: 0.2
M phosphate buffer, pH 6.0 and 8.0; 0.2 M Tris-HCl buffer pH 8.5 and 9.0;
0.2 M carbonate-bicarbonate buffer pH 9.5 and 10.5 with olive oil as substrate
as described above. The results were expressed as percentage of the activity
obtained at pH 5.5. Furthermore, the data obtained for the measurements
above pH 5.5 were excluded from the results presented here due to substrate
instability. The effect pH on lipase stability was determined by incubating
the lipase fraction in various buffer solutions ranging from 2.0 to 8.0
for 30 min at 25°C in a Haake circulating water bath. The compositions
of the buffer solutions used for the pH stability studies were as follows:
0.2 M acetate buffer, pH 4.0; 0.2 M AcONa-HCl buffer pH 3.10; 0.2 M citrate-phosphate
buffer, pH 6.0; 0.2M phosphate buffer, pH 8.0; and 0.2M bicarbonate-carbonate
buffer, pH 10.5. After the incubation period, 100 μL aliquots of
the buffered enzyme solutions were added to 900 μL of the olive oil
substrate and lipase activity was assayed spectrophotometrically at 410
nm as described previously.
Effect of Temperature on the Activity and Stability of Lipases
The temperature dependence of Lip-1 and Lip-2 activity were measured
by equilibrating olive oil at 10, 15, 20, 25, 30, 32, 33, 34, 35, 40,
45, 50, 55, 60 and 70°C for 30 min. In each assay, 100 μL of
the enzyme extract was added to 900 μL of pre-equilibrated substrate.
The thermostability of the lipase fraction was studied by incubating the
enzyme extract at various temperatures for 10, 30, or 60 min. At the end
of the incubation period the enzyme extract was rapidly cooled and the
remaining lipase activity was assayed using olive oil.
Effect of Various Chemicals on Enzyme Activity and Stability
Aliquots of the enzyme were incubated with equal volumes of the various
chemicals and metal ions at various concentrations at 25°C in a Haake
circulating water bath (Haake D1-G, HAAKE Mess-Technik GmbH Co., Germany)
for 30 min. Appropriate blanks and control were performed for each chemical
tested. The relative activity of Lip-1 and Lip-2 after incubation period
was assayed with olive oil as substrate as previously described in duplicates
and the results were expressed as percentage of the activity obtained
without the chemical agent.
RESULTS AND DISCUSSION
Isolation and Purification of Lipases
Few scientists have been studied fish lipases and reported different
purification methods and yields (Gjellesvik et al., 1992; Iijima
et al., 1998). The crude enzyme solution after dialysis against
10 mM Tris-HCl buffer, pH 8.4 for 24 h, at 4°C was applied to firstly
ion exchange chromotography a DEAE for purification.
In Fig. 1, the proteins of crude enzyme solution were separated in two major peaks: F-1 and F-2 and one minor peak F-3. Of these fractions F-1 was eluted by buffer only while F-2 and F-3 were eluted by the buffer containing 0.1 M NaCl and 0.2 M NaCl, respectively. Further, it was found that only F-1 fraction contained lipase activity. So, the area as indicated by solid bar of this fraction was pooled together and used for further purification by CM-cellulose chromatography.
The enzyme active fraction F-1 was dialyzed against distilled water for
12 h and against the eluting buffer (5 mM sodium phosphate buffer, pH
6.5) for 12 h at 4°C. After centrifugation the
| Fig. 1: | Ion exchange chromatography of crude enzyme solution on DEAE-cellulose. The crude solution (20 mg) was applied to the column (2.1x25 cm) pre-equilibrated with 10 mM Tris-HCl buffer, pH 8.4 at 4°C and eluted by stepwise increases of NaCl concentration in same buffer. Flow rate: 25 mL h-1 |
| Fig. 2: | CM-cellulose chromatography of F-1a fraction obtained from ion exchange chromatography on DEAE-cellulose. Fraction F-1a (12 mg) was applied to the column (1.5x15 cm) pre-washed with 5 mM sodium phosphate buffer, pH 6.5 at 4o C and eluted by stepwise increases of NaCl in the same buffer. Flow rate was 20 mL h-1 |
clear supernatant was applied to the CM-cellulose column. As shown in Fig. 2, the components in F-1 was separated into two sharp peaks F-1a and F-1c and one small peak, F-1b. Of these fractions F-1a was eluted by the buffer only while F-1b and F-1c were eluted by the buffer containing 0.1M and 0.2 M NaCl, respectively. It was found that the fractions F-1a and F-1b contained lipase activities while F-1c` possessed no such activity
The area as indicated by solid bar of these fractions (F-1a, F-1b) were pooled separately and their purity was checked by SDS-PAGE.The phtographic representation of the elctrophoretic patterns of F-1a and F-1b are presented in Fig. 3. From the result it might be concluded that the fraction F-1a was not pure as it gave more that one band on the gel which the fraction F-1b must be contained pure protein as it gave single band on the gel. The fraction F-1a was then further purified by gel filtration, on Sephadex G-50.
The fraction F-1a was dialyzed against distilled water for 12 h and then
against 10 mM Tris-HCl buffer, pH 8.4 for 12 h. After concentration, this
fraction was applied to a Sephadex G-50 column at 4°C, which was previously
equilibrated with the same buffer. In Fig. 4, the components
present in
| Fig. 3: | Polyacrylamide disc gel electrophoretic pattern of the protein at room temp. pH 8.3 on 7.5% Polyacrylamide gel. Staining reagent : 1% amido black. Protein concentration : 50 μg. a : F-1a fraction b: F-1b fraction |
| Fig. 4: | Gel filtration of F-1a fraction obtained from CM-cellulose chromatography on Sephadex G-50. F-1a fraction (6 mg) was applied to the column (3x120 cm) pre-washed with 10 mM Tris-HCl buffer, pH 8.4 at 4°C and eluted by the same buffer. Flow rate: 25 mL h-1 |
this fraction was eluted as one major peak, F-1a and one minor peak F-1b. It was found that only the fraction F-1a contained the lipase activity while F-1b contained no lipase activity. The active enzyme fraction, as indicated by solid bars was pooled and its homogeneity was checked by Polyacrylamide gel elctrophoresis.
In Fig-5, the fraction F-1a gave clear single band on polyacrylamide gel indicating that this kaj korchifraction contained pure lipase enzyme. Lipase activities in the course of purification steps are shown in the following table (Table 1).
Although the activities of the enzymes were decreased at each subsequent
purification steps but the purification fold of the enzymes were increased
(Helisto et al., 2001; Yu et al., 2007) at each step
| Fig. 5: | Disc electrophoretic pattern of F-1a´ protein on 7.5% polyacraylamide gel. Protein concentration: 60 μg |
| Table 1: | Lipase activities in the course steps of purification of phaisa fish lipase |
and the purifical fold of F-1a and F-b became about 38 and 19 folds, respectively The decrease in yield might be due to denaturation of the enzymes during lengthy purification procedure.
Determination of Molecular Weight (MW) of Lipases
The MW of the enzymes were determined by gel filtration on Sephadex
G-150 using β-galactosidase, bovine serum albumin, α-amylase,
ovalbumin, trypsin inhibitor and lysozyme as standard proteins. The MW
of purified lipases were estimated to be 46.5KDa for F-1a´ and 41.2KDa
for F-1b. Iwai et al., 1975 purified two lipases from Penicillium
cyclopium having MW of 27.KDa and 36KDa, while Yu et al., 2007;
Aloulou et al., 2007 reported the MW of the lipase Lip2 from Yarrowia
lipolitica a were 38 and 38.,48KDa, respectively..
The MW of these enzymes were also determined by SDS-PAGE using the same
marker proteins and the MW of F-1a` and F-1b were found to be 46.5KDa
and 41.2KDa, respectively. Abbas et al., 2002. reported the molecular
weight of lipase from Mucor sp strain isolated from palm fruit
was 42KDa (Choo et al., 1998) and Helisto et al., 2001 showed
two protein bands on SDS-PAGE with MW 35,000 and 46,000 Da. Further, the
MW of the enzymes were found to be unchanged in the presence or absence
of β-mercaptoethanol indicating that both the lipases contained only
one subunit. The lipases, F-1a and F-1b were denoted as Lip-1 and Lip-2,
respectively.
| Fig. 6a: | Effect of pH on the activity of Lip-1 and Lip-2L. The buffers used were phosphate, pH 6.0-8.0; Tris-HCl, pH 8.4-9.0; carbonate-bicarbonate, pH 9.4 and 10.0; all the buffers were 0.2 M. Data used to plot figure are average values of duplicate results for two experiments |
| Fig. 6b: | Effect of pH on the stability of Lip-1 and Lip-2L. The enzyme extract was incubation at 25 °C for 30 min in various buffers; HCl-KCl, pH 2.0; glycine-HCl, pH 3.0; acetate, pH 4.0 and 5.0; phosphate, pH 6.0-8.0; and carbonate-bicarbonate pH 10.0; all the buffers were 0.2 M. Data used to plot figure are average values of duplicate results for two experiments |
Effect of Ph on the Activity of Lipases
Lip-1 and Lip-2 hydrolyzed olive oil appreciably over a relatively
broad alkaline pH range (from pH 7.0 to 10.0). Beyond pH 9.0, the activity
of the enzymes decreased gradually to pH 10.0 (Fig. 6a).
These trend were similar to findings reported for other fish lipases (Gjellesvik
et al., 1992; Raso and Hultin, 1988; Mukundan et al., 1985;
Iijima et al., 1998 and is also consistent with the pH optima reported
for lipases from other sources (Lima et al., 2004; Nthangeni et
al., 2001). The Lip-1 and Lip-2 both enzymes were stable between pH
7.0 and 10.0 during incubation at 25°C for 30 min. There was 55% residual
activity above pH 7.0. However, there was a marked loss of activity in
the acidic region below pH 4.0 with 55% decrease in activity, compared
to the activity at pH 8.0 (Fig. 6b).
| Fig. 7: | Effect of temperature on the activity of Lip-1 and Lip-2. The substrate (Olive oil) was incubation at the indicated temperature for 30 min. Data used to plot figure are average values of duplicate results for t experiments |
This observation is also similar to those found for other fish lipases (Mukundan et al., 1985) as well as lipases from microorganisms such as Pseudomonas sp. strain B11-1, Bacillus licheniformis and Aspergillus carneus (Lima et al., 2004; Saxena et al., 2003; Zouari et al., 2005), but different from scorpion lipase that was reported to be most stable at pH 11.0 (Borlongan, 1990) It is worth mentioning here that Lip-1 and Lip-2 was both also exhibited greater stability in phosphate buffers than in Tris-HCl buffer. For instance, Lip-1 and Lip-2 retained about 60, 50 and 44% and 6%, 48 and 40% of initial activity when incubated in phosphate, citrate-phosphate and Tris-HCl buffer, respectively, at pH 7.0 (data not shown). This observation was similar to the findings of Lima et al., 2004; Nthangeni et al., 2001.
Effect of Temperature on the Activity of Lipases
The effects of temperature on the activities of Lip-1 and Lip-2 were
examined in the range of 10 to 70°C. Lipolytic activities of grey
mullet lipases were found to be profoundly affected by temperature. In
Fig. 7 the activities of Lip-1 and Lip-2 were increased
gradually with rise in temperature and the maximum activities were observed
around 33 and 35°C, respectively. These observed temperature optimum
in this study was similar to the lipase from sardine and cod were 37 and
25°C with tributyrine and p-nitrophenyl myristate as substrate, respectively
(Gjellesvik et al., 1992; Raso and Hultin, 1988), For instance,
the reported optimum temperature of lipase from milkfish, 45-50 oC
with olive oil as substrate (Tan et al., 2004). and lipases from
the psychrotroph Pseudomonas sp. strain B11-1 and Penicillium
camembertii Thom PG-3 with temperature optima of 45 and 48°C with
p-nitrophenyl butyrate and olive oil as substrates (Lima et
al., 2004; Destain et al., 1997). With further rise in temperature
the activities were decreased abruptly and the enzymes lost almost 100%
of their activities at 60 and 70°C for Lip-1 and Lip-2, respectively.
Grey mullet fish lipases are found to be highly active in the temperature
range from 30 to 35° C. Lipases purified from other sources were found
to be highly active in the temperature range from 30 to 40°C. Destain
et al., 1997 reported that the optima pH and temperature of extracellular
lipase from Y. lipolytica was 7.0 and 37 8°C, while this lipases from
Yarrowia lipolitica was 40°C (Aloulou et al., 2007).
Data used to plot figure are average values of duplicate results for experiments.
| Table 2: | Effect of metallic salts on the activities of phaisa fish lipases |
Effect of Metal Ions on the Activity of Lipases
Understanding the role of lipase inhibitors may provide a better perceptive
of their mechanism of action (Marguet et al., 1994) and successful
identification of potent and specific inhibitors have resulted in their
application in certain treatments (Bray, 2000; Kotsovolou et al.,
2007). In this study, the enzyme was incubated with various compounds
and relative activities were measured after 30 min of incubation at 25°C
From the Table 2, it is evident that activities of grey
mullet fish lipases inhibited by heavy metals such as Cd++,
Zn++ and Hg++. The presence of Zn2+
and Hg+ potently inhibited lipolytic activities of the lipases
from grey mullet fish, while activities were slightly inhibited in the
presence of Cu2+ salts. This is likely due to the proximity
of the SH group to the catalytic and interfacial binding site but partially
remote from the catalytic site, this may have induced the marked loss
of activity (Raso and Hultin, 1988; Verger et al., 1971). The catalytic
triad of lipases has been recognized to consist of Ser, His and Glu or
Asp (Guidoni et al., 1981, Fredrikson et al., 1981; Jin
et al., 2002). thus the bulky Hg2+ group might cause steric interference
to the approach of the substrate to the active site.. Salts of Fe were
found to inhibit lipase from Aspergillus niger (180) and lipases
from fungi of the genus Geotrichum. Mercury is known to bind to
thiol groups of proteins forming stable complexes. Mercury has been shown
to inhibit the lipse of rat adipose tissue (Fredrikson et al.1981).
It has been shown conclusively that some heavy metals inhibit lipase activity,
However, treatment with Zn2+ at both 0.003 and 0.005M significantly inhibited
the activity of Lip-1 and Lip-2 with only 30 and 20% and 25 and 18 % relative
activity after 30 min incubation and this was similar to the findings
of Choo et al., 1998.
Calcium ions carry out distinct role in the lipase action. The activities of both the lipases were found to be increased slightly in presence of lower concentration of Ca2+ but at higher concentration of Ca2+, the activities were decreased slightly which is consistent with the results reported elsewhere (Shastry. and Raghavendra Rao, 1971). Like pancreatic lipase (Liu et al., 1973) grey mullet lipases were stimulated in the presence of calcium ions at certain concentrations. The primary role of Ca++ seems to be to remove the released fatty acid as its calcium salt. The activating effect of calcium on a lipase derived from Humicola lanuginosa was explained by the removal of free fatty acids from the interface. In a calcium free system, the lipase can not adsorb at the water-fat interface and consequently no lipolytic activity occurs. Possibly, the calcium ions compensate for the electrostatic repulsion created between the enzyme and the substrate. (Shahani, 1975).
EDTA, 0.001 activated the Lip-1 and Lip-2 by 65 and 75%, respectively, similar to the results of Lima et al., 2004. The decrease in activity may be due to the removal of metal ions located on or near the active site.
CONCLUSIONS
The purification of a lipase from grey mullet fish (Liza parsia), is described in this paper for the first time. The pure enzyme, tentatively named Lip-1 and Lip-2, is a glycosylated protein with molecular weight of 46.5KDa and 41.2KDa, respectively. The optimum pH and temperature of Lip-1 and Lip-2, 8 and 8.3, 33 and 35°C, respectively. Lip-1 and Lip-2 activity were stimulated by Ca++ and Cu++ and inhibited by Hg++ and Zn++. Further studies to determine its performances in esters synthesis and specificity in hydrolysis of unsaturated fish oil.
ACKNOWLEDGMENTS
We are grateful to Dr. H. Rahman (Laboratoire d`Enzymology and protein, University of Rajshahi, Bangladesh) Dr. M A Bari, Dr. P. Hasan (Laboratory, Institute of Biological sciences, University of Rajshahi, Bangladesh) for his fruitful discussions and kind help during the preparation of this work. We acknowledge Mr K A Salam� for his technical assistance. This work is part of a doctoral thesis. This work received financial support from “Ministry of Science and technology, Bangladesh. granted to the Laboratory of Enzymology and protein, department of Biochemistry and Molecular Biology, University of Rajshahi, Bangladesh