Camp. Biockm. Physiol. Vol. 103C, No. 1, pp. 221-225, 1992

0306-4492/92$5.00+ 0.00 0 1992Pergamon Press Ltd

Printed in Great Britain

ESTERASE HETEROGENEITY IN MUSSEL MYTILUS GALLOPROVINCIALZS: EFFECTS OF ORGANOPHOSPHATE AND CARBAMATE PESTICIDES IN VITRO B. OZRETIC and M. KRAJNOVIC-OZRETIC Institute “Rudjer BoSkoviC”, Center for Marine Research, 52210 Rovinj, Croatia (Received 20 December

Abstract-l. In mussel Mytihs gahprovincialis molecular forms of esterases were observed.

1991)

tissue-specific electrophoretic

patterns

of multiple

2. Carboxylesterases and acetylesteraseswere identified following the application of pertinent substrates and inhibitors. 3. Hepatopancreas was the richest source of esterase activity followed by gills, ovaria, adductor muscle and mantle. 4. In vitro studies indicated that organophosphate and carbamate insecticides have distinct effects on different esterase isozymes. Esterases classified as carboxylesterase were more sensitive to parathion and particularly to paraoxon.

INTRODUCI’ION

Organophosphate and carbamate insecticides can contaminate surface waters through direct application for aquatic insects control, or indirectly through watershed drainage or accidental spillage. Aquatic invertebrates living in surface waters can thus be exposed to insecticides which range from acutely lethal to sublethal levels. The inhibition of acetylcholinesterase (AChE) activity has been successfully used to diagnose organophosphate and carbamate poisoning in birds and fish (Busby et al., 1983; Coppage, 1972; Hill and Fleming, 1982; Holland et al., 1967; Jarvinen et al., 1983; Lochart et al., 1985; Zinkl et al., 1987). However, few studies have examined the effects of organophosphate insecticides on AChE in aquatic invertebrates (Bocquiene et al., 1990; Day and Scott, 1990; Srivinasulu Reddy and Ramana Rao, 1988). Certain organophosphates and carbamates at low concentration are also very efficient to inhibit other serum esterases, as carboxyl or aliesterases (“B”) and, due to their more rapid and higher inhibition rate, carboxylesterases may be preferable indicators of poisoning by pesticides than brain AChE (Thompson et al., 1988; Chambers et al., 1989). Since AChE and carboxylesterases exist in various isozyme forms (Booth and Metcalf, 1970; Bunyan et al., 1968; KrajnoviC-OzretiC and de Ligny, 1978; Tripathi and O’Brien, 1973; 1975) it may be possible to recognize distinct inhibition patterns peculiar to different insecticides and can therefore provide an additional early warning system to perceive detrimental effects on population and communities levels. Before mussels or any other marine organisms could be utilized as indicators of environmental contamination, more data must be acquired on the nature of their enzymes. Although the literature on mammalian esterase and to lesser extent on avian esterase is quite comprehensive, very little is known

about shellfish esterases. The present paper contributes to the identification and characterization of mussel esterases according to their tissue distribution, some biochemical properties and isozyme forms. The potential use of mussel esterases for monitoring purposes was investigated on the basis of their inhibition by various organophosphate and carbamate pesticides in vitro. MATERIALSAND METHODS Mussels (Mytilus galloprovincialis Lam.) used in this study were furnished from the Limski Kanal mariculture facilities (Center for Mariculture, Mirna Rovinj). Mussels were about two years old and their size varied between 65-75mm. Tissues were separately excised and immediately homogenized in a Polytron homogenizer with 4 parts of 0.1 M Tri-HCl b&I& pH 7.8. Homogenates were-centrifuged for 30min. at 27000~ in a Sorvall RCZ-B centrifuge. Suuernatant was imm&ately removed and used preferably fresh or stored at -20°C. The esterase activity and their characteristics were examined in gills, mantle, adductor muscle and ovaria but with particular concern in extracts from hepatopancreas. Esterase activity was estimated by the UV method of Mastropaolo and Yourno (1981). Samples of 0.05 ml tissue extracts were introduced into cuvettes containing 0.93 ml of 0.05 M Tris-HCl buffer, pH 7.8 and 0.02 ml of a 0.3 mM solution of cc-naphthyl acetate in ethylene glycol monomethyl ether. The formation of naphthol at 25°C was monitored with a spectr*photometer at 235nm. The optimal pH and substrate concentration were also determined. Protein concentration was determined according to Bradford (1976). Electrophoresis was carried throughout 5 to 6 h at 50 mA and 280 V on horizontal slabs of 12% hydrolysed starch in Tris-borate buffer at pH 8.6 for gel and pH 9.1 for electrodes. The enzymatic activity of separate estcrase bands on starch gels was revealed by histochemical methods. After completion of electrophoresis gels were sliced horizontally and incubated for 45 min in a solution of 250 ml 0.1 M Tri-HCl buffer pH 7.8 with 5 ml 1% substrate in acetone and 250mg of Fast Blue BB salt. After staining, gels

221

B. OZRETICand M. KRAJNOVIC-OZRETIC

222

a- Naphtyl

acetate

(mM)

PH

Fig. 1. Mytilus gulloprooinciulis. Determination of the optimal substrate concentration (A) and pH (B) for the measurement of esterase activity (optical density change mitt-’ at 235 nm) in extracts from hepatopancreas.

were rinsed with distilled water and stored in a methanol/water/acetic acid mixture (45/45/10%). For the qualitative identification of esterases a-naphthyl acetate, j?-naphthyl acetate, a-naphthyl butyrate, anaphthyl propionate and acetylthiocholine iodide (Sigma, Co) were used as substrates. According to the staining intensity, the esterase fractions were scored from + + + (dark-high activity), + + (medium-moderate activity), + (weak-low activity) and-+0 staining-absence of activity). The inhibitory studies were performed by preincubation of sliced gels in solutions containing eserine (10e4M), diisopropylfluorophosphate (10e4 M DFP) and parachloromercurybenzoate (10m3M pCMB) for 30 min before the addition of appropriate substrates. The classification of esterase bands was determined according to their sensitivity to specific substrates and inhibitors (Augustinson, 1961; Holmes et al., 1968; Hart and Cook, 1976). Nine pesticides, representing several different chemical classes were used to study their inhibitory effects on the mussel esterases. Dichlotvos, bromphos, phosalone, malathion, guthion, parathion, carbaryl and baygon were applied at concentrations within 10m3to 10e6 M. Paraoxon, the P(0) analog of parathion was also included in this study. The purity of pesticides was about 95-98%. The inhibition rating was scored by comparing the staining intensity of the control zymograms (gels not treated with inhibitors) with the intensity of the bands in the incubated gels. The degree of inhibition was ranked as follows: + + + (complete inhibition-no evidence of bands) + + (moderate inhibition-bands faintly visible), + (slight inhibition-weak but distinct bands) and - (no inhibition-staining intensity of the bands as in control slices).

RESULTS For the best accurate measurement of esterase activity the substrate concentration and pH optima were determined. As displayed in Fig. lA, the optimum substrate concentration of 0.3 mM a-naphthyl acetate was adopted. The optimum pH range was within 7.0 to 8.5 and, for routine the pH 7.8 was adopted (Fig. 1B). The distribution of esterase activity in tissue extracts and in respective organs is illustrated in Table 1. Owing to the highest specific activity and to biomass, hepatopancreas was found to be the richest source of esterase activity followed by ovaria, gills, mantle and adductor muscle. Figure 2 gives a synthesized map of the isozyme electrophoretic patterns from tissue extracts prepared from nearly 300 specimens. Electrophoretic separation and histochemical staining displayed the existence of several esterase isozyme forms. The number of bands varied from seven in ovaria to fifteen in adductor muscle. According to electrophoretic mobility and sensitivity to various substrates and inhibitors, bands were grouped in five anodic zones (A 1-5) and one cathodic zone (C 1). Zone A 5 was not found in extracts of ovaria, while in the adductor muscle extracts, the C 1 zone was missing. Within zones A 1 and A 2 the isozyme bands were found singly or in pairs. The greatest variations were found in zone A 3, with two till six bands. Esterases from zone A 4 were usually restricted to only one band resembling to wide diffuse blots on the starch gel slices. The slowest anodic zone A 5 was also very variable. It was missing

Table 1. Mytilus galloprovincialis. Mean protein concentration and esterase specific activity in tissue extracts, and rough estimate of the whole enzyme activity in the organs

N = 35 Hepatopancreas Mantle Gills Ovaria Adductor muscle

Tissue extracts A B mg ml-’ U mg-’ 12.17 2.85 2.75 13.57 6.07

0.27 0.08 0.13 0.03 0.06

Whole tissues/organs E C D u 106 u mg-1 g 16.59 0.727 12.06 1.13 0.969 I .09 1.a2 1.079 1.96 1.69 1.984 3.35 1.67 0.597 1.oo

A-Protein concentration; kxtract specific activity (JI mole min-’ referred to 1 mg protein in the extract); C-tissue specific activity 01 mole min-’ referred to 1 mg tissue wet weight); D-mean weight of whole organs; E-estimated whole activity in the organs.

Mussel esterases MM4TL.B

HBPATOPANCREAS

------

-

-

A5

----_Cl

_-

ADDUCTOR NDSCLS

---

-

----

--

-

---

---

223

------------------

--

-----

DILLS

_

_

_

-

-

----__ -

-

-

OVhRIA

---

0

Fig. 2. Mytilusgalloprovincialir. Esterase electrophoretic patterns from various tissue extracts. Five anodic

zones (A 1-5) and one catodic (Cl) were identified. in the ovarian extracts, gills contained only one band, one to three bands were found in hepatopancreas and mantle, while in extracts from the adductor muscle three to six bands were present. In the cathodic zone, mantle extracts contributed with the highest number of bands. In general the number of bands and their staining intensity from various tissues was very variable, but according to the highest specific activity, extracts from hepatopancreas displayed the best resolution and the highest staining intensity (Table 1 and Fig. 2). Hepatopancreas extracts were therefore used for further analyses concerning the classification of esterase bands and zones in response to various substrates, inhibitors and pesticides. The specific response of esterase zones to various substrates and inhibitors are summarized in Table 2. According to Augustinson (1961), Holmes et al. (1968) and Hart and Cook (1976) bands from the A 1, A 4 and C 1 zones were classified as acetylesterases since they were not inhibited by DFP (lo-‘M), eserine (lo-’ M) or pCMB (1O-3 M) and preferentially hydrolysed acetyl ester of naphthol. The activity was higher with a-naphthyl acetate, than with /3naphthyl acetate, while a-esters of propionate and butyrate were not hydrolysed. All isozyme bands from zones A 2, A 3 and A 5 were found to hydrolyse a - and B-esters of acetate as well as a -naphthyl esters

of propionate and butyrate (Table 2). However, the staining intensity of zone A 3 was lower with b-naphthy1 acetate. All isozymes were resistent to eserine (lo-‘M) and to pCMB (lo-‘M), while DFP (lo-‘M) totally or partially inhibited the isozymes from these zones. According to the described characteristics A 2, A 3 and A 5 bands were classified as carboxylesterases (Augustinson, 1961; Holmes et al., 1968; Hart and Cook, 1976). The inhibition of esterase activity on single bands and zones was investigate in vitro by post-incubation of starch gel slices with pesticides. It was obvious that, owing to specific properties of various pesticides and their concentrations, different groups of esterases responded with distinct inhibition patterns (Table 3). Esterases from hepatopancreas were considerably more sensitive to parathion and particularly to paraoxon, than to other tested pesticides. Paraoxon, at the highest concentration (lo-) M) inhibited all esterase bands but, at lower concentrations, both paraoxon and parathion inhibited only the esterases from zones A 2 and A 3, which were classified as carboxylesterases. Other pesticides exhibited different inhibition patterns, specific to various esterase fractions and dependent to pesticide concentration. Considerable differences were observed also among pesticides from the same group. Although malathion,

Table 2. Mytilus galloprouinciaIis. Estemsc activity and specific response of esterase zones from hepatopancreas to various substrates and inhibitors and their classification Esterase zones

Al

A2

A3

A4

Substrate specificity* a-naphthyl acetate /3-naphthyl acetate a-naphthyl butyrate a-naphthyl propionate acetylthiocholine iodide

++ + -

+++ +++ +++ +++ -

++-I++ +++ +++ -

+++ ++ + -

Inhibitor spe=cificityt Eserine sulfate (lo-‘M) DFP (lo-‘M) pCMB (1O-3 M)

-

+++

++

Classificationf

ACE

CE

CE

ACE

AS + + + + +++ _ CE

Cl +++ +++ + ACE

‘Estimation of esterase activity based on the staining inteosity of the electrophoretic bands: + + + high activity, + + moderate, + low activity, and - absence of activity. tThe inhibitory effects were scored in comparison to control aymogram as follows: + + + complete inhibition; + + moderate, + slight and - no inhibition. $AcE: acetylesterases; CE: carboxylesterases.

B. OZRETIC and M. KRAJNOVIC-OZRETIC

224

Table 3. Myrilus galloprouincialis. Inhibitory effectst of various pesticides on esterases zones from hepatopancreas Control

M

Al ++

A2 +++

A3 +++

A4 +++

A5 +

++ ++ ++ ++ ++ ++ _

+ + _ _

+ + _ -

++ + + -

++ ++ + -

++ ++ + _

++ ++ + -

+++ +++ +++ ++ +++ +++ ++ + ++ + + -

++ ++ + + ++ + + -

++ ++ + + ++ ++ + -

++ ++ ++ + ++ ++ + -

+++ +++ +++ +++ +++ +++ +++ +++ ++ + + ++ ++ + +

+++ +++ +++ +++ +++ +++ +++

+++ ++ + + ++ + + ++ + + + ++ ++ ++ +

+++ +++ +++ +++ +++ +++ +++ ++ +++ +++ ++ ++ +++ +++ +++ +++ +++ +++ +++ ++ +++ + _ _

Dichlorvos

10-s 10-J 10-s 10-e

Bromphos

lo-’ 10-d 10-S 10-e

Phosalone

lo-’ 10-d 10-5 10-e

Malathion

10-S 10-d 10-S 10-s

Guthion

10-S 10-d 10-S 10-L

++ ++ ++ ++ +++ +++ ++ ++ ++ ++ -

Paraoxon

10-S 10-d 10-S 10-s

+++ +++ +++ _

Parathion

10-4 10-S 10-s

Carbaryl

10-S 10-d 10-5 10-s

++ ++ ++ +++ ++ ++ ++ ++ ++ ++ ++

Baygon

10-3 10-d 10-5 10-s

*,tSee footnotes of Table 2; M-Molar

-I-+ ++ + + fi++ +

+

++ + + +++ +++ +++ ++ +++ +++ +++ +++

Cl

+++ +++ ++ + + ++ + + + _ _ _ +++ +++ ++ + +++ ++ + + +++ + _ _ + + f + f +++ +++ ++ +

concentration.

guthion and parathion are all phosphorothionates, with basic chemical similarity, they showed considerable differences in toxicity. Esterases from zone A 1 were particularly sensitive to malathion, while other pesticides produced only slight inhibitory effects. Esterases from zone A 2 and A 3 were inhibited only with the highest concentrations of all tested pesticides. Zone A4 consists of esterases particularly sensitive to dichlorvos and to high concentrations of bromphos while other pesticides, yet at the highest provided only slight inhibitory concentrations, effects. The most sensitive esterases were found in the A 5 zone, and with the exception of parathion and paraoxon they were substantially affected by all other pesticides. Esterases from region C 1 were completely inhibited by higher concentration of dichlorvos, malathion and baygon, whereas other pesticides displayed only slight inhibitory effects. DISCUSSION

In a recent report Bocquene et al. (1990) argued about the existence of very low acetylcholinesterase activity in hepatopancreas, mantle, adductor muscle and gills of the blue mussel (M. edzdis). In zymograms from hepatopancreas of the mediterranean mussel (M. galloprovincialis) we have not found any acetylcholinesterase or arylesterase activity. However, we

supposed that they were not present or, due to their very low concentration, they did not reach the level necessary for their histochemical demonstration on starch gel slices. Although the primary mode of action of organophosphate pesticides is generally related to the inhibition of acethylcholinesterase, the inhibition of other enzymes, like carboxylesterase was also expected as confirmed by Thompson et al. (1988) and Chambers et al. (1989). According to electrophoretic mobility and histochemical staining and following the application of pertinent substrates and inhibitors, esterase bands from all examined tissues were grouped in five anodic and one cathodic zones. Among them, two primary groups were identified as carboxyl and acetyl esterases. Thus, the presence of carboxyl and acetyl esterase groups particularly in hepatopancreas gave the opportunity to check these enzyme systems as possible targets for monitoring poisoning of mussel with organophosphate and carbamate insecticides. The specific inhibitory effects of several pesticides and their selective inhibition of different esterases from hepatopancreas were easily and rapidly surveyed in vitro by starch gel electrophoresis. The obtained results demonstrated that esterases from hepatopancreas were substantially more sensitive to parathion and particularly to paraoxon. Other pesticides exhibited different inhibition patterns, specific

Mussel esterases

for various esterase zones. Since all bands from each esterase zone responded to single pesticides as a whole independent unit, these results further substantiated the structural similarity within each esterase zone and emphasized the difference among zones. We also found that esterases either sensitive or resistant to DFP, and classified as carboxyl or acetyl esterases, behave different to various classes of organophosphates. Although eserine did not, carbamate pesticides like carbaryl and bygon significantly suppressed the activity of all esterase groups. In general, our findings provided a valuable evidence about the response of mussel esterase isozymes to estimate the cpecific effects of organosphosphate and carbamate pesticides. Finally, for monitoring purposes it would be also very attractive to examine the selective response of the identified esterase phenotypes in vivo: in laboratory and in field studies.

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Jan&n A. W., Nording B. R. and Henry M. E. (1983) Chronic toxicity of Dursban (chlorpyrifos) to the fathead minnow (Pimephales promelas) and the resultant acetylcholinesterase inhibition. Ecotoxicol. Environ. Safety 7, 423-443.

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