Comparative Biochemistry and Physiology Part B 137 (2004) 159–168
Esterase and lipase in camel tick Hyalomma dromedarii (Acari: Ixodidae) during embryogenesis Afaf S. Fahmy, Somia S. Abdel-Gany, Tarek M. Mohamed, Saleh A. Mohamed* Molecular Biology Department, National Research Centre, Cairo, Egypt Received 8 January 2003; received in revised form 20 October 2003; accepted 27 October 2003
Abstract Esterase and lipase activity showed significant changes during embryogenesis of camel tick Hyalomma dromedarii. From the elution profile of chromatography on DEAE-cellulose, six forms of H. dromedarii esterase (El to EVI) can be distinguished. Esterase EIII was purified to homogeneity after chromatography on Sepharose 6B. The molecular mass of esterase EIII was 45 kDa for the native enzyme and represented a monomer of 45 kDa by SDS-PAGE. Esterase EIII had an acidic pI at 5.3. Lipase activity was detected in the same DEAE-cellulose peaks (LI to LVI) of H. dromedarii esterases. The highest lipase activity was exhibited by lipase LIII. Esterase EIII and lipase LIII were compared with respect to Michaelis constant, substrate specificity, temperature optimum, heat stability, pH optimum, effect of metal ions and inhibitors. This study suggests that H. dromedarii lipolytic enzymes may play a central role in the interconversion of lipovitellins during embryogenesis. 䊚 2003 Elsevier Inc. All rights reserved. Keywords: Camel; Tick; Hyalomma dromedarii; Embryogenesis; Esterase; Lipase; Purification; Characterization
1. Introduction Esterases (EC 3.1.1.1) and lipases (EC 3.1.1.3) attack the fatty acyl linkage of water-insoluble triacylglycerols. In insects, esterases are involved in important physiological processes, including the catabolism of juvenile hormone (Zera et al., 1992, 2002), pesticide resistance (Whyard et al., 1995; Rosario-Cruz et al., 1997), digestion (Kerlin and Hughes, 1992; Argentine and James, 1995) and reproduction (Richmond and Senior, 1991; Karotam and Oakeshott, 1993). Lipid in insects is released as diacylglycerol (DAG) by lipase and transported to the flight muscles by lipophorin. In the flight muscles, DAG is hydrolyzed and liber*Corresponding author. Tel.: q202-3669980; fax: q2023370931. E-mail address:
[email protected] (S.A. Mohamed).
ated fatty acids are taken up and oxidized to provide energy (Ogoyi et al., 1998; Ryan and Van der Horst, 2000). Lipases are also involved in digestion (Brahimi-Horn et al., 1989; Biesiot and Capuzzo, 1990). Vitellins represent the bulk of insect yolk proteins. They are believed to be a nutritive source of amino acids, lipids, carbohydrates and phosphorus during embryogenesis (Engelmann, 1979; Hagedorn and Kunkel, 1979). Two lipovitellins A and B (lipoglycohaemoproteins) were isolated and purified from Dermacentor andersonii eggs. The purified fractions were immunologically identical to female haemolymph proteins but not to host haemoglobin (Boctor and Kamel, 1976). The disappearance of haemoproteins A and B within the first 8 days of embryogenesis and the appearance of new haemoproteins C and D suggested their
1096-4959/04/$ - see front matter 䊚 2003 Elsevier Inc. All rights reserved. doi:10.1016/j.cbpc.2003.10.017
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interconversion (Kamel et al., 1982). Two vitellins A and B, were isolated and purified to homogeneity from H. dromedarii and identified as a haemoglycolipoprotein (Hamed et al., 1990). Insect yolk proteins contain appreciable amounts of lipids ranging from 6.9 to 15.7% (Kunkel and Pan, 1976; Mintzas and Kambysellis, 1982). The total lipid contents of lipovitellins A and B in D. andersoni are 5.7% and 8.5%, respectively, (Boctor and Kamel, 1977). However, the lipid contents of H. dromedarii lipovitellins A and B are 12.2% and 20.8%, respectively (Hamed et al., 1990). Besides lipolytic enzymes as degradative enzymes involved in utilization of yolk materials during embryogenesis of H. dromedarii, no information had been reported on these enzymes in this organism, although these enzymes had been studied in other insects. The lack of this information has prompted the present study, which concentrates on understanding the physiology of degradation of lipids in H. dromedarii during embryogenesis. The purification and characterization of lipolytic enzymes are studied from the view point of comparative biochemistry. 2. Materials and methods 2.1. Ticks Engorged Hyalomma dromedarii (Acari: Ixodidae) females were collected from camels in the market near Cairo and held at 28 8C and 85% relative humidity. Eggs were collected daily from fertilized ovipositing female ticks, incubated under the same conditions until the appropriate age and then frozen (y40 8C) at intervals of 3 days. Eggs frozen immediately were designated 0 day. 2.2. Chemicals p-Nitrophenyl derivatives of acetate, valerate, palmitate and stearate; tributyrin; tricaproin; tricaprin; trilaurin; trimyristin; tripalmitin; tristearin; triarachidin; tribehenin; molecular weight markers for gel filtration and for SDS polyacrylamide gel electrophoresis; isoelectric focusing markers 3.6– 8.8; phenylthiourea; diethylaminoethyl (DEAE)cellulose for chromatography; p-hydroxymercuribenzoate (p-HMB); N-ethylmaleimide; 5,59 dithiobis (2-nitrobenzoicacid) (DTNB); phenylmethysulfonyl fluoride (PMSF); iodoacetamide and all resins and reagents for electrophoresis were
obtained from Sigma Chemical Co. (St Louis, MO, USA). 2-Mercaptoethanol and tributyrin were products of Fluka (Switzerland). Arabic gum and olive oil were purchased from a local market. Sepharose 6B was obtained from Amersham BioScience (Sweden). Other chemicals were of analytical grade. 2.3. Esterase assay Esterase activity was measured by the procedure of Galliard and Dennis (1974). The 1 ml assay reaction mixture contained 2 mM p-nitrophenyl acetate as substrate, 50 mM Tris–HCl buffer, pH 7.0 and suitable amount of enzyme preparation. The reaction mixture was incubated for 1 h at 37 8C and then the enzyme activity was determined spectrophotometrically by reading the p-nitrophenol released at 405 nm. One unit of esterase activity was defined as the amount of enzyme liberating 1 mmol p-nitrophenol per hour under standard assay conditions. The rate of p-nitrophenol release was linearly increased with increasing the H. dromedarii enzyme concentration, and incubation time up to 0.2 units and 1 h incubationyassay, respectively. 2.4. Lipase assay Lipase activity was measured according to the method of Tietz and Fiereck (1966, 1972). Olive oil and tributyrin emulsions were prepared by mixing 100 ml water, 0.2 g sodium benzoate and 7 g gum acacia in a mechanical blender at slow speed until dissolved. 100 ml of olive oil or tribuytrin was added slowly and then mixed for 10 min at a maximum speed, cooling at intervals. The 3 ml assay reaction mixture contained 2 ml of substrate emulsion (0.78 g of olive oil or 1.3 g of tribuytrin), 20 mM Tris–HCl, pH 8.0, 5 mM CaCl2 and suitable amount of enzyme preparation. The reaction mixture was incubated for 1 h at 37 8C. The reaction was stopped by addition of 0.6 ml 96% ethanol. This was then titrated with 5 mM sodium hydroxide in the presence of an indicator (5 drops of phenolphthalein). One unit of lipase activity was defined as the amount of enzyme liberating 1 mmol fatty acid per hour under standard assay conditions. The amounts of oleic and isobutyric acids increased linearly with increasing H. dromedarii enzyme concentration, and incuba-
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tion time up to 15 and 21 units and 1 h incubationy assay, respectively. 2.5. Purification of esterase from H. dromedarii 24-day-old eggs Unless otherwise stated all steps were performed at 4–7 8C using 50 mM Tris–HCl buffer, pH 7.0. Step 1: Preparation of crude extract. Crude extract was prepared by homogenizing 2 g of 24day old eggs in 10 ml of 50 mM Tris–HCl buffer, pH 7.0, containing 1 mM EDTA, 0.01% phenylthiourea and 1 mM 2-mercaptoethanol using a Teflon pestle homogenizer. The homogenate was centrifuged at 13 200=g for 20 min to remove insoluble debris and the supernatant was designated as the crude extract and dialyzed overnight against the same buffer. The dialysate was then centrifuged at 16 300=g for 20 min to remove precipitated protein. Step 2: Ammonium sulfate precipitation. Crude extract was precipitated by 80% ammonium sulfate at 4 8C with continuous stirring. After 20 min, the precipitate was collected by centrifugation at 16 330=g for 15 min and dissolved in 50 mM Tris–HCl buffer, pH 7.0 and dialyzed overnight against the same buffer. Step 3: DEAE-cellulose chromatography. The dialyzed fraction was applied directly to a DEAEcellulose column (30=2.6 cm i.d.) equilibrated with the same buffer. The exchanged material was eluted with a stepwise gradient ranging from 0.0 to 1.0 M sodium chloride prepared in the same buffer at a flow rate of 60 mlyh and 5 ml fractions. Protein fractions exhibiting esterase activity were pooled in six peaks (El to EVI) according to their elution order. Step 4: Sepharose 6B chromatography. Esterase III was applied to a Sepharose 6B column (90=1.6 cm i.d.) equilibrated with the same buffer and developed at a flow rate of 20 mlyh and 3 ml fractions. The esterase III was eluted with the same buffer.
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2.7. Polyacrylamide gel electrophoresis (PAGE) Electrophoresis under non-denaturing conditions was performed in 10% (wyv) acrylamide slab gels (Davis, 1964) using a Tris–glycine buffer, pH 8.3, in a Bio-Rad MINI PROTEAN apparatus. Protein bands were located by staining with Coomassie Brilliant Blue R-250. 2.8. Molecular weight determination Molecular weight was determined by gel filtration using Sepharose 6B. The column (90=1.6 cm i.d.) was calibrated with cytochrome C (12 400), carbonic anhydrase (29 000), bovine albumin (67 000), alcohol dehydrogenase (150 000) and b-amylase (200 000). Dextran blue (2 000 000) was used to determine the void volume (V0). Subunit molecular weight was estimated by SDS-polyacrylamide gel electrophoresis (Laemmli, 1970). SDS-denatured bovine serum albumin (66 000), ovalbumin (45 000), glyceraldehyde-3-phosphate dehydrogenase (36 000), carbonic anhydrase (29 000), trypsinogen (24 000), trypsin inhibitor (20 100) and a-lactalbumin (14,200) were used for calibration. 2.9. Isoelectric focusing The focused protein was separated by analytical isoelectric focusing using a high voltage vertical slab polyacrylamide gel system according to Giulian et al. (1984). A calibration curve was constructed by plotting the isoelectric point (pI) of the marker proteins (amyloglucosidase, pI 3.6; trypsin inhibitor, pI 4.6; b-lactoglobulin A, pI 5.1;carbonic anhydrase II, pI 5.9; carbonic anhydrase I, pI 6.6; myoglobin, pI’s 6.8; 7.2; lentil lectin pI’s 8.2; 8.6; 8.8) against their distance from the anode. For pI determination, the distance from anode of the unknown was measured and its pI was interpreted from the calibration curve. 3. Results
2.6. Protein determination and buffers 3.1. Developmental changes Protein was determined either by measuring the absorbance at 280 or by the method of Bradford (1976) using bovine serum albumin as a standard. Buffers were prepared according to Gomori (1955), and the final pH was checked with an ElL pH meter Type 7020.
The developmental profiles of lipolytic enzymes showed significant changes during embryogenesis (Fig. 1). Esterase activity exhibited its lowest level on day 6 (0.376"0.023 units mgyl protein), then significantly increased (P-0.01) on day 9
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(P-0.01) to a maximum level on day 24 (0.41"0.018 units mgyl protein). With tributyrin as substrate lipase activity was 0.07"0.002 units mgyl protein on day 6 and 0.194"0.009 units mgyl protein (P)0.01) on day 24. The results indicated that the activity of lipase with olive oil was 2.11-fold higher than with tributyrin. 3.2. Purification of esterase from H. dromedarii 24-day-old eggs The purification of esterase is summarized in Table 1. Six forms of esterase EI to EVI were demonstrated according to their elution order in DEAE-cellulose chromatography (Fig. 2). Their specific activities ranged from 2.84 to 12.2 units mgyl protein. Further purification was restricted to esterase EIII. A Sepharose 6B column (Fig. 3) was used to obtain esterase EIII with a specific activity of 22.5 units mgyl protein. Fig. 1. Changes in esterase and lipase activities levels during embryogenesis of H. dromedarii using p-nitrophenyl acetate and olive oil, tributyrin as substrates, respectively. Each point represented the mean of 3 runs for each developing stage"S.E.
(0.64"0.025 units mgyl protein). The activity was fairly constant until day 18, when it was significantly increased (P-0.01) to reach its maximum level on day 24 (1.19"0.07 units mgyl protein). Lipase activity using olive oil as substrate exhibited its lowest level on day 9 (0.079"0.002 units mgyl protein) and significantly increased
3.3. Homogeneity The electrophoretic behaviour of different purification steps was examined. Only one band was visualized on polyacrylamide gel after Sepharose 6B step, which indicated the homogeneity of the final preparation of esterase EIII (Fig. 4). 3.4. Molecular weight and isoelectric point The molecular weight of H. dromedarii esterase EIII was estimated to be 45 000 using gel filtration
Table 1 Purification scheme for H. dromedarii esterase Step
Total protein (mg)
Total activity (units)*
S.A. (unitsymg protein)
Fold purification
Crude extract Ammonium sulphate (80%) DEAE-cellulose 0.0 M NaCl (EI) 0.1 M NaCl (EII) 0.2 M NaCl (EIII) 0.3 M NaCl (EIV) 0.5 M NaCl (EV) 1 M NaCl (EVI) Gel filtration of EIII on Sepharose 6B
483 107.79
571 636
1.18 5.9
– 5
12.6 7.3 10.4 5.06 14.85 10.1 4
105 85 127 30 41 60 90
8.3 11.6 12.2 5.9 2.84 5.9 22.5
7 9.83 10.3 5 2.33 5 19
Recovery % 100 111.3
18.4 14.9 22.2 5.23 7.1 10.5 15.8
* One unit of esterase activity was defined as that amount of enzyme liberating 1 mmol p-nitrophenol per hour under standard assay conditions.
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Fig. 2. A typical elution profile for the chromatography of H. dromedarii esterase on DEAE-cellulose column (30=5 cm i.d.) previously equilibrated with 50 mM Tris–HCl buffer, pH 7.0 at a flow rate of 60 mlyh and 5 ml fractions, absorbance at 280 nm (d –– d), esterase activity (x—x).
technique (Sepharose 6B). This value was confirmed by SDS–PAGE, where the molecular weight of esterase was estimated to be 45 000 (Fig. 5). The homogenous esterase EIII is seen on isoelectric focusing gel as one band with isoelectric point (pI) of 5.3 (Fig. 6). 3.5. Detection of lipase activity Lipase activity was detected in the same DEAEcellulose peaks of H. dromedarii esterase using
olive oil as substrate. The highest lipase activity was exhibited for lipase LIII (3.75 units mgyl protein). 3.6. Characterization of esterase EIII and lipase LIII Characterization was carried out for esterase EIII using p-nitrophenyl acetate and lipase LIII using olive oil as substrates.
Fig. 3. A typical elution profile for H. dromedarii esterase EIII DEAE-cellulose fraction on Sepharose 6B column (90=1.6 cm i.d.) previously equilibrated with 50 mM Tris–HCl buffer, pH 7.0 at a flow rate of 20 mlyh and 3 ml fractions, absorbance at 280 nm (d –– d), esterase activity (x—x).
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Fig. 4. Polyacrylamide gel electrophoresis of H. dromedarii esterase during purification steps: (1) Crude extract. (2) DEAE-cellulose (EIII). (3) Gel filtration (EIII).
The Km values of esterase EIII and lipase LIII were estimated to be 1.43 mM p-nitrophenyl acetate and 0.5 g olive oil, respectively. Affinity of esterase EIII decreased as increase in carbon atom number of the substrates (Table 2) in the order of p-nitrophenyl acetate) p-nitrophenyl valerate) pnitrophenyl palmitate) p-nitrophenyl stearate. The
Fig. 6. Identification of H. dromedarii esterase EIII on isoelectric focusing gel. (1) Esterase EIII. (2) Standard proteins.
lipase LIII had more affinity toward olive oil (100%), with the affinity decreasing in the order of triarachidin)tributyrin) tricaprin )trilaurin. However, tricaproin, trimyristin, tripalmitin and tristearin had the same affinity toward the enzyme with 25% relative activity. Table 2 Relative activity of H. dromedarii esterase EIII and lipase LIII toward a series of substrates. Each value represents the mean of three experiments "S.E. Substrate specificity
Fig. 5. Sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis for molecular weight determination of H. dromedarii esterase EIII. (1) Standard proteins. (2) Esterase EIII.
Esterase EIII p-nitrophenyl p-nitrophenyl p-nitrophenyl p-nitrophenyl Lipase LIII Tri butyrin Tricaproin Tricaprin Trilaurin Trimyristin Tripalmitin Tristearin Triarachidin Tribehenin Olive oil
Relative activity % acetate valerate palmitate stearate
100"7 91"6 50"3.4 35"2.6 62.5"4.8 25"1.9 50"3.2 37.5"2.8 25"2.1 25"1.9 25"1.8 75"5.3 25"1.8 100"7.2
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Table 3 Effect of metal ions on H. dromedarii esterase EIII and lipase LIII. Enzyme was pre-incubated for 15 min at 37 8C with 1 mM of listed metals as final concentration prior to substrate addition. Activity without metal ions was taken as 100% activity. Each value represents the mean of three experiments "S.E.
Fig. 7. Temperature optimum (a), temperature stability (b) and pH optimum (c) of H. dromedarii esterase EIII (d –– d) and lipase LIII (x—x). Each point represents the mean of three experiments "S.E.
The effect of temperature on the activity and stability of esterase EIII and lipase LIII was examined (Fig. 7a,b). Esterase EIII and lipase LIII had temperature optima at 50 8C and 40 8C, respectively. Temperature stability of esterase EIII showed that the enzyme lost 52% of its activity at 60 8C after incubation for 15 min prior to substrate addition. The enzyme stability was decreased gradually with increasing temperature till the activity reached 15% at 90 8C. For lipase LIII, 53% of its activity was lost at 70 8C and completely lost at 90 8C. Esterase EIII and lipase LIII had pH optima
Metal ions
Esterase EIII Relative activity %
Lipase LIII Relative activity %
Ca2q Liq Fe3q Hg2q Mg2q Mn2q Ni2q Zn2q
67"4.2 100"7.5 28"1.9 79"4.5 94"6.3 90"6.8 75"4.8 73"4.2
50"3.9 44"3.5 32"3.0 25"2.1 100"8 19"1.5 14"1.4 2"0.2
at pH 7.0 and pH 8.0 in Tris–HCl buffer, respectively (Fig. 7c). Studying the effect of metal ions on esterase EIII and lipase LIII indicated that no inhibitory effect could be detected by Liq and Mg2q, respectively (Table 3). The effectiveness of metal ions as inhibitors for esterase EIII was listed in the order of Fe3q)Ca2q)Zn2q)Ni2q)Hg2q) Mn2q)Mg2q. The effectiveness of metal ions as inhibitors for lipase LIII was Zn2q)Ni2q) Mn2q)Hg2q)Fe3q)Liq)Ca2. Table 4 shows the effect of different inhibitors on esterase EIII and lipase LIII. No inhibitory effects could be detected with 2-mercaptoethanol, p-HMB and DTNB, while PMSF caused a 53% inhibition for esterase EIII. PMSF was estimated to be a noncompetitive inhibitor with inhibition constant (Ki) of 8.3 mM. The effectiveness of these inhibitors on lipase LIII decreases in the order p-HMB) Table 4 Effect of different inhibitors on H. dromedarii esterase EIII and lipase LIII. Enzyme activity was measured in the presence of 5 mM of the inhibitors listed. Activity without inhibitors was taken as 100% activity. Each value represents the mean of three experiments "S.E. Inhibitors*
Esterase EIII Relative activity %
Lipase LIII Relative activity %
b-Mercaptoethanol lodoacetamide N-Ethylmaleimide p-HMB DTNB PMSF
101"8.5 98.5"7.8 98"8.1 117"10 116"9.5 47"3.6
100"8.2 105"9.5 110"10 40"2.5 70"4.5 95"8.5
* p-Hydroxymercuribenzoate, p-HMB; 5,59-dithiobis (2-nitro benzoic acid), DTNB; phenylmethylsulfonyl fluoride, PMSF.
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DTNB)PMSF. p-HMB and DTNB were estimated to be competitive inhibitors for lipase LIII with Ki’s of 0.2 and 0.25 mM, respectively. 4. Discussion The developmental profiles of esterase and lipase during embryogenesis of H. dromedarii were studied, where both enzymes exhibited their highest levels on day 24. Several investigations studied the levels of esterases during embryogenesis and development. For house cricket, Acheta domesticus, the duration of embryogenesis was 9.5 days and the level of a-naphthyl acetate esterase activity remained relatively constant throughout embryogenesis and was similar to that of eggs dissected from the oviducts. Juvenile hormone esterase (JHE) activity was highest in the dissected and day 1 eggs, declined to one-third of this peak activity by day 5, and then remained unchanged through hatching (Roe et al., 1987). JHE was also followed during metamorphosis for red flour beetle, Tribolium freemani (Coleoptera: Tenebrionidae) (Hirashima et al., 1995), and wasp Biosteres longicaudatus (Hymenoptera:Braconidae) (Lawrence et al., 1990). Lipase of the American lobster (Homarus americanus) had been demonstrated during larval development (Biesiot and Capuzzo, 1990). Using chromatography on DEAE-cellulose, six forms (El to EVI) of H. dromedarii 24-day-old eggs esterases were demonstrated, where EIII represented the highest enzyme. However, only a single esterase from tick (Boophilus microplus; Willadsen et al., 1987) and mosquito (Culex tarsalis; Whyard et al., 1995) was obtained after chromatography on Mono Q and DEAE-Sepharose, respectively, while three esterases from workers of the eastern subterranean termite (Reticulitermes flavipes) were obtained using sequential preparative isoelectric focusing (Valles et al., 2001). The final specific activity of H. dromedarii esterase EIII (12.2 units mgyl protein) was higher than that obtained from tick B. microplus (2.84 units mgyl protein) employing ammonium sulfate, DEAE-Sepharose, isoelectric focusing and Sephacryl S-200 (Willadsen et al., 1987). The native molecular weight of H. dromedarii EIII was 45 000. This value was confirmed by SDS-PAGE (Laemmli, 1970) and represented a monomer with a subunit of 45 000. This value was similar to the molecular weight of an organophosphate resistant
strain of the cattle tick B. microplus (43 000) (Rosario-Cruz et al., 1997) and smaller than that of esterases from mosquito Culex tarsalis (59 000) (Whyard et al., 1995), eastern subterranean termite Reticulitermes flavipes (69 000, 64 000, 62 000) (Valles et al., 2001), and organophosphate resistant strain of the cattle tick B. microplus (125 000, 115 000, 77 000, 67 000) (Rosario-Cruz et al., 1997). The homogenous H. dromedarii esterase EIII is seen on isoelectric focusing gel as one major fine band with an acidic pI of 5.3. This value was close to the pI for esterases from Gryllus rubens (pIs 4.1, 5.2) (Zera et al., 1992), Lymantria dispar (pIs 5.1, 5.3) (Nussbaumer et al., 2000), R. flavipes (pIs 4.61, 4.70, 4.73) (Valles et al., 2001), and G. assimilis (pIs 4.7–4.9) (Zera et al., 2002). The Km value of H. dromedarii EIII (1.43 mM) using p-nitrophenyl acetate was higher than that of most esterases of other invertebrates. Different Km’s were observed for Culex tarsalis esterases using different substrates, malathion, a-naphthyl acetate and b-naphthyl acetate. The Km’s ranged from 2.4 to 76.4 mM substrate (Whyard et al., 1995). The apparent Km values of R. flavipes for a-naphthyl acetate were similar for esterases ME1 (52.1 mM) and ME2 (47.9 mM), but comparatively lower in ME3 (26.5 mM) (Valles et al., 2001). The Km of H. dromedarii lipase LIII was 0.5 g olive oil. The Km values of Schistocerca gregaria (Ogoyi et al., 1998) and locust Locusta migratoria (Hirayama and Chino, 1987) lipases were 65 mM and 56 mM using triolein as substrate, respectively. A series of fatty acid nitrophenylesters with different chain length were tested as substrates for H. dromedarii esterase EIII. The affinity of esterase EIII decreased with increase in carbon atom number in the order of p-nitrophenyl acetate (C8))pnitrophenyl valerate (C11))p-nitrophenyl palmitate (C22))p-nitrophenyl stearate (C24). As a result it can be concluded that H. dromedarii esterase EIII had a broad substrate specificity toward the number of carbon atoms of the pnitrophenyl esters up to 24 carbons. In contrast, tick B. microplus esterase hydrolyses a series of fatty acid nitrophenyl esters of acetate, propionate, butyrate, caproate or valerate with little specificity for the chain length (Willadsen et al., 1987). Valles et al. (2001) reported a severe decline in specific activity of each of the purified microsomal esterases from R. flavipes when the a-naphthyl acyl chain length increased from two to six carbons,
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and the absence of detectable activity with 8 to 14 carbons. For substrate specificity of H. dromedarii lipase LIII, a series of triglycerides with different chain lengths have been tried as substrates. Substrate specificity of H. dromedarii lipase LIII was not similar to esterase EIII, where it did not depend on the length of carbon chain. However, triarachidin with 20 carbon atoms caused high relative activity (75%), the tributyrin (C4) caused relative activity of 62.5%. A more compelling reason may lie in the substrate used to measure lipase activity. Triolein was used with lobster H. americanus (Biesiot and Capuzzo, 1990), whereas tributyrin was used by Brockerhoff et al. (1970). H. dromedarii esterase EIII had an optimum temperature at 50 8C. It was higher than that reported for esterases from G. flavipes ME1 and ME2 (35 8C) and ME3 (40 8C) (Valles et al., 2001). H. dromedarii lipase LIII activity shows a sharp peak at 40 8C and stability until 40 8C. Lobster H. americanus lipase activity shows increases from 20 to 45 8C, although the rate of increase diminishes somewhat between 30 and 45 8C and the reaction rate decreases between 45 and 50 8C (Biesiot and Capuzzo, 1990). The pH optimum of H. dromedarii esterase EIII was 7.0. This was similar to the pH optima for esterase from tick Boophilus microplus (pH optima at neutral and alkaline pH values) (Willadsen et al., 1987). Most lipases studied in detail show neutral to slightly alkaline pH optima (Jensen, 1983). H. dromedarii lipase LIII had a pH optimum at 8.0. The pH optimum of lipoprotein lipase from Manduca sexta appeared to have slightly basic pH optimum (pH 7.5) (Van Heusden, 1993), which is quite similar to that of the flight muscle lipoprotein lipase of the locust Locusta migratoria (pH 7.65) (Wheeler et al., 1984) and that of mammalian lipoprotein lipases (pH 8–8.6) (Nilsson-Ehle, 1987), but different from that of M. sexta egg yolk lipolytic activity (pH 9.1–9.5) (Van Antwerpen and Law, 1992). 2-Merrcaptoethanol, p-HMB, iodoacetamide and N-ethylmaleimide had no inhibitory effect on H. dromedarii esterase EIII, while PMSF caused 53% inhibition, indicating that H. dromedarii esterase EIII possessed a serine residue in the active site. The same inhibitory effect of PMSF was detected for R. flavipes esterases (Valles et al., 2001). Eserine sulfate had no inhibitory effect on esterases MCEI and II from Culex tarsalis (Whyard et al., 1995). The arylesterase inhibitor, p-CMB, inhibited both MCEs
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equally. The MCEs were most sensitive to the organophosphates, showing that they both behave as carboxyesterases. In contrast, H. dromedarii lipase LIII was not inhibited by PMSF as H. dromedarii esterase EIII and the other insect lipases (Van Heusden and Law, 1991; Van Antwerpen and Law, 1992). H. dromedarii lipase LIII was inhibited by thiol reagents DTNB and p-CMB, suggesting that H. dromedarii lipase LIII contains cysteine residues. Esterase and lipase appear to play a principal role in the interconversion of H. dromedilrii lipovitellins during embryogenesis. Lipovitellins are considered a nutritive source for lipids, which are important fuels for insects. References Argentine, J.A., James, A.A., 1995. Characterization of a salivary gland-specific esterase in the vector mosquito Aedes aegypti. Insect Biochem. Mol. Biol. 25, 621–630. Biesiot, P.M., Capuzzo, J.M., 1990. Digestive protease, lipase and amylase activities in stages larvae of the American lobster Homarus americanus. Comp. Biochem. Physiol. A 95, 47–54. Boctor, F.N., Kamel, M.Y., 1976. Purification and characterization of two lipovitellins from eggs of the tick Dermacentor andersoni. Insect Biochem. 6, 233–240. Boctor, F.N., Kamel, M.Y., 1977. Biochemical studies of tick embryogenesis. Free amino acid pools during embryogenesis of Dermacentor anderonsi. Comp. Biochem. Physiol. B 56, 169–173. Bradford, M.M., 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72, 248–254. Brahimi-Horn, M.C., Guglielmino, M.L., Sparrow, L.G., Logan, R.I., Moran, P.J., 1989. Lipolytic enzymes of the digestive organs of the crown-of-thorns starfish (Aconthaster planci) comparison of the stomach and pyloric caeca. Comp. Biochem. Physiol. B 92, 637–643. Brockerhoff, H., Hoyle, R.J., Hwang, P.C., 1970. Digestive enzymes of the American lobster (Homarus americanus). J. Fish Res. Bd. Can. 27, 1357–1370. Davis, B.J., 1964. Disc electrophoresis. Method and application to human serum proteins. Ann. NY Acad. Sci. 121, 404–427. Engelmann, F., 1979. Insect vitellogenin: Identification, biosynthesis and role in vitellogenesis. Adv. Insect Physiol. 14, 49–108. Galliard, T., Dennis, S., 1974. Isoenzymes of lipolytic acyl hydrolase and esterase in potato tuber. Phytochem. 13, 2463–2468. Giulian, G.G., Moss, R.L., Greaser, M., 1984. Analytical isoelectric focusing using a high vertical slab polyacrylamide gel system. Anal. Biochem. 74, 430–440. Gomori, G., 1955. Preparation of buffers for use in enzyme studies. Methods. Enzymol. I, 138–146. Hagedorn, H.H., Kunkel, J.G., 1979. Vitellogenin and vitellin in insect. Annu. Rev. Entomol. 24, 475–505.
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Hamed, R.R., Fahmy, A.S., Kamel, M.Y., 1990. Lipovitellins and amino acid pools during embryogenesis of the camel tick Hyalomma dromedarii (Acarina:Ixodidae). Entomol. Gener. 15, 1–10. Hirashima, A., Takeya, R., Taniguchi, E.J., Eto, M., 1995. Metamorphosis activity of juvenile hormone esterase and alteration of ecdysteroid titres: Effects of larval density and various stress on the red flour beetle Tribolium freemani hinton (Coleoptera: Tenebrionidae). J. Insect Physiol. 4, 383–388. Hirayama, Y., Chino, H., 1987. A new method for determining lipase activity in locust fat body. Insect Biochem. 17, 85–88. Jensen, R.G., 1983. Detection and determination of lipase (acylglycerol hydrolase) activity from various sources. Lipids 18, 650–657. Kamel, M.Y., Shalaby, F.Y., Ghazy, A.M., 1982. Biochemical studies of tick embryogenesis DNA, RNA, haemoprotein, guanosine and guanine in developing eggs of Hyalomma dromedarii. Insect Biochem. 22, 15–23. Karotam, J., Oakeshott, J.G., 1993. Regulatory aspects of esterase 6 activity variation in sibling Drosophila species. Heredity 71, 41–50. Kerlin, R.L., Hughes, S., 1992. Enzymes in saliva from four parasitic arthropods. Med. Vet. Entomol. 6, 121–126. Kunkel, J.G., Pan, M.L., 1976. Selectivity of yolk protein uptake: Comparison of vitellogenins of two insects. J. Insect Physiol. 22, 809–818. Laemmli, U.K., 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685. Lawrence, P.O., Baker, F.C., Tsai, L.W., Miller, C.A., Schooley, D.A., Geddes, L.G., 1990. JH III levels in larvae and pharate pupae of Anastrepha suspensa (Diptera: Tephridae) and in larvae of the parasitic wasp Biosteres longicaudatus (Hymenoptera:Bracanidae). Arch. Insect Biochem. Physiol. 13, 53–62. Mintzas, A.C., Kambysellis, M.P., 1982. The yolk proteins of Drosophila melanogaster. Isolation and characterization. Insect Biochem. 12, 25–33. Nilsson-Ehle, P., 1987. Measurements of lipoprotein lipase activity. In: Borentsztajn, J. (Ed.), Lipoprotein Lipase. Evener, Chicago, pp. 59–77. Nussbaumer, C.A., Hinton, A.C., Schopf, A., Stradner, A., Hammock, B.D., 2000. Isolation and characterization of juvenile hormone esterase from the haemolymph of Lymantria dispar by affinity-and by anion-exchange chromatography. Insect Biochem. Mol. Biol. 30, 307–314. Ogoyi, B.O., Osir, E.O., Olembo, N.K., 1998. Fat body triacylglycerol lipase in solitary and gregarious phases of Schistocerca gregaria (Forshal) (Orthoptera: Acrididae). Comp. Biochem. Physiol. B 119, 163–167.
Richmond, R.C., Senior, A., 1991. Esterase 6 of Drosophila melanogaster: Kinetics transfer to females, decay in females and males recovery. J. Insect Physiol. 27, 849–853. Roe, R.M., Crawford, C.L., Clifford, C.W., Woodring, J.P., Sparks, T.C., Hammock, B.D., 1987. Characterization of juvenile hormone esterases during embryogenesis of the house cricket, Acheta domesticus. Int. J. Invert. Rep. Dev. 12, 57–72. Rosario-Cruz, R., Miranda-Miranda, E., Garcia-Vasquez, Z., Ortiz-Estrada, M., 1997. Detection of esterase activity in susceptible and organophosphate resistant strains of the cattle tick Boophilus microplus (Acari: Ixodidae). Bull. Entomol. Res. 87, 197–202. Ryan, R.O., Van der Horst, D.J., 2000. Lipid transport biochemistry and its role in enzyme production. Annu. Rev. Entomol. 45, 233–260. Tietz, N.W., Fiereck, E.A., 1966. A specific method for serum lipase determination. Clin. Chim. Acta 13, 352. Tietz, N.W., Fiereck, E.A., 1972. In: Cooper, G. R. (Ed.), Standard Methods of Clinical Chemistry, Vol. 7. Academic Press, New York and London, pp. 19. Valles, S.M., Oi, F.M., Strong, C.A., 2001. Purification and charactetrization of transpermethrin metabolizing microsomal esterases from workers of the eastern subterranean termite Reticulitermes flavipes (Kollar). Insect Biochem. Mol. Biol. 31, 715–725. Van Antwerpen, R., Law, J. H., 1992. Lipophorin lipase from the yolk of Manduca sexta eggs identification and characterization. Arch. Biochem. Physiol. 20, 1–12. Van Heusden, M.C., 1993. Characterization and identification of a lipoprotein lipase from Manduca sexta flight muscle. Insect Biochem. Mol. Biol. 23, 785–792. Van Heusden, M.C., Law, J. H., 1991. Lipoprotein lipase from Manduca sexta flight muscle. Sericologia 31, 91. Wheeler, C.H., Van der Horst, D.J., Beenakkers, A.M.T., 1984. Lipolytic activity in the flight muscles of Locusta migratoria measured with haemolymph lipoproteins as substrates. Insect Biochem. 14, 261–266. Whyard, S., Down, A.E.R, Walker, V.K., 1995. Characterization of a novel esterase conferring insecticide resistance in the mosquito Culex tarsalis. Arch. Insect Biochem. Physiol. 29, 329–342. Willadsen, P., Mckenna, R.V., Riding, G.A., 1987. Purification and characterization of an esterase from the tick Boophilus microplus. Insect Biochem. 17, 659–664. Zera, A.J., Gu, X., Zeisset, M., 1992. Characterization of juvenile hormone esterase from genetically-determined wing morphs of the cricket Gryllus rubens. Insect Biochem. Mol. Biol. 22, 829–839. Zera, A.J., Sanger, T., Hanes, J., Harshman, L., 2002. Purification and characterization of hemolymph juvenile hormone esterase from the cricket Gryllus assimilis. Arch. Insect Biochem. Physiol. 49, 41–55.
