Some properties of the sucrase from the digestive gland of the green mussel Perna viridis L.

Comp. Biochem. Physiol.Vol. 96B, No. 1, pp. 47 51, 1990

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SOME PROPERTIES OF THE SUCRASE FROM THE DIGESTIVE G L A N D OF THE GREEN MUSSEL P E R N A VIRIDIS L. L. H. TEO,* ZAHARA LATEEF a n d Y. K. IP Department of Zoology, National University of Singapore, Kent Ridge, Republic of Singapore 0511

(Received 4 September 1989) Abstract--l. The optimal pH of sucrase was 5.8, which was slightly lower than the pH of the digestive gland. 2. The K M of sucrase was 0.184 M and the Vmaxwas 106.8/~mol/mg protein/hr. 3. The optimal temperature was 40°C and the activation energy was 10,600 cal/mol. 4. The most potent inhibitor of sucrase was mercury chloride, followed by copper sulphate, cadmium chloride, lead chloride and zinc chloride. 5. Starvation caused a decrease in the sucrase activity of the digestive gland.

INTRODUCTION

the enzyme extracts were heated in a boiling water bath for 10 rain to destroy the enzymes. The reaction mixtures were incubated at 35°C for 2 hr. Two drops of toluene were added to each tube, which was then sealed with parafilm. The amount of reducing sugar produced as a result of sucrase activity was determined with the dinitrosalicylate reagent (Sumner, 1925). Colorimetric determinations were made with a spectrophotometer at 530 nm. The protein contents of the enzyme extract were determined with the Bio-Rad standard assay procedure, with bovine serum albumen as standard. To study the effect of temperature (four samples for each temperature), on sucrase activity the reaction mixtures were incubated for 1.5 hr at temperatures varying from 20 to 65°C. The inhibition of sucrase activity by bivalent salts of heavy metals was also investigated, by adding various concentrations of different compounds to the enzyme extract, which had been subjected to dialysis against three changes of distilled water (1 1) at 5°C for 24 hr. Buffer and substrate solutions were then added in the same manner and volume as above. The reaction mixture was incubated at 35°C for 2 hr. The mussels were starved for 2-10 days. At the end of each starvation period the digestive gland of each animal was assayed for its sucrase activity, as described above. Feeding mussels fed with unicellular alga Isoehrysis galbana were used as a control.

The digestive gland o f bivalves is rich in carbohydrases (Owen, 1974). The carbohydrases include glycosidases t h a t act on sucrose, maltose, lactose, raffinose (Fox a n d Marks, 1936), fl-N-acetylglucosaminidase which hydrolyzes chitobiose ( M o z o et al., 1977), a n d amylase (Kamat, 1957). However, a p a r t from this, little is k n o w n a b o u t the properties of these enzymes. T h e present work was u n d e r t a k e n to study the kinetics of the sucrase from the digestive gland o f the green mussel. D e t e r m i n a t i o n of properties such as p H t e m p e r a t u r e optima, Michaelis' c o n s t a n t a n d m a x i m u m velocity allow one to m a k e meaningful c o m p a r i s o n s of the same enzyme from different species (Haldane, 1965). Y o n g e (1935) has stressed t h a t bivalves are c o n t i n u o u s filter feeders, with mainly intracellular digestion, a n d are therefore unlikely to show a periodic enzyme output, b u t Reid (1968) showed t h a t starvation in bivalves caused a reduction in enzyme production. Thus, we decided to look at the effect of starvation on the o u t p u t of the enzyme again. MATERIALS AND METHODS Specimens of the green mussel were brought back from the field and then kept in a refrigerator at 4°C for no more than three to four days. Animals of shell lengths 3-5 cm (average 3.5cm) were selected for all experiments. The digestive gland was homogenized in a homogenizer containing marine invertebrate saline (Morton, 1955). The homogenate was made in the ratio of one gland to 1.5 ml saline. The preparation was then centrifuged for 20 min at 6000 g to remove the larger particles. The supernatant was decanted off" and assayed for enzymic activity. The substrate used was a 5% (0.146 M) sucrose solution, except in the study of the effect of substrate concentration. The reaction mixture in each tube consisted of 1 ml enzyme extract, 2 ml substrate solution and 1 ml standard buffer solution [of Clark and Lubs (Oser, 1965)]. In the control

RESULTS The optimal p H of the sucrase was f o u n d to be 5.8 (Fig. 1), which is close to the p H of the digestive gland 6.19 + 0.02 (Teo a n d U m a , u n p u b lished observation). As the m o l a r c o n c e n t r a t i o n s of sucrose were increased from 0.005 to 0.03 M, there was a rapid increase in the rates o f sucrase hydrolysis (Fig. 2), b u t further increase in m o l a r c o n c e n t r a t i o n hardly b r o u g h t a b o u t any significant increase in the rates of enzymic activities. The W o o l f - A u g u s t i n s s o n - H o f s t e e plot (Fig. 3) was done to estimate the following kinetic constants: K m = 0.014 M, Vmax = 106.78/zmol glucose/mg protein/hr. The direct linear plot (Fig. 4) also gave the same estimate of Km.

*Author to whom correspondence should be addressed. 47

48

L.H. TEO et al. As the temperature was increased from 20°C to 35°C, there was almost a linear increase in the activity of sucrase. After the maximum activity was reached at 40°C, there was only a gradual decrease instead of a rapid drop in enzymic activity (Fig. 5). Even at 65°C the activity of sucrase still remained at about 78% of the activity at 40°C. This indicates that the sucrase of the green mussel is not very sensitive to high temperature. The Arrhenius plot (Fig. 6) shows that the activation energy for the green mussel sucrase was 10,600 cal/mol. The effects of different bivalent salts on the activities of sucrase are shown in Fig. 7. The most potent inhibitor was mercury chloride, followed by copper sulphate, cadmium chloride, lead chloride and zinc chloride, respectively. The data obtained for all replicates (n = 4) of a given concentration was pooled and analyzed by the probit method (Finney, 1971).

x: 1"2¸ ~.o

~ 0-8 ib ~

0.6

-~ 0.4 0.2

4.0

s'-o

6:o

7~0

810

pH

Fig. 1. Effect of pH on the activity of sucrase.

100 .c_. ®

f

90

o,.

~

80

~

~0

o

~

60

o E

50

~

40

o ~-

30

:a,

o.~2

o:o4 Molar

o.~6

o:o8

1.'oo

Concentration

Fig. 2. Effect of substrate concentration on the activity of sucrase.

100

80

P~ 7

60

:~ 40

°

20

0

1.0

20

3.0 v

50

6"0

x lO3

S

Fig. 3. The W o o l f - A u g u s t i n s s o n - H o f s t e e p l o t o f the r e l a t i o n s h i p between substrate c o n c e n t r a t i o n a n d sucrase activity.

Perna sucrase

49

=~ 2 . 0 Ih A C

C

g ~ E

1.0

~ O.5

_,e =L

2~

4'0

6'o

Temperature (Oc) Fig. 5. Effect of temperature on the activity of sucrase. o lo

Km(0.014)

o.o8 oo6 0 . 0 4 0.02 Concentration (M)

Fig. 4. The direct linear plot between substrate concentration and sucrase activity.

0.4 0-3 0.2

~

0 ~..,~.>9~X,~

0.1 0'0 = o --4

-0'1 EA= 10,600 cal Imole

-0,2

O

|

310

2.9

3'1

31"2

3:3

3'4

T Fig. 6. The Arrhenius plot of the effect of temperature on sucrase activity.

80

a-a mercury chloride H copper Sulphate O--Ocadmlum chloride e--e lead chloride

,

t

°

4

6

v

~

~

~ 6o

~ C

2O

2

Concentration (,AIM)

8

10

Fig. 7. Relationship between percentage inhibition of sucrase a n d / t M concentration of the inhibitor.

50

L.H. TEa et al.

4~

-¢o

-6'.0

-51o

-4'.o

Log Inh'lbitorconcentration(2uM)

Fig. 8. Relationship between probit of inhibition of sucrase and log pM concentration of the inhibitor. This method allowed us to estimate the concentrations of inhibitors required to cause 50% inactivation of the sucrase activity. Probit of inhibition was plotted against log p M concentration, and the regression of each line was then computed (Fig. 8). For mercury chloride the concentration for 50% inactivation of sucrase was 2.239/~mol; copper sulphate 2.818pmol; cadmium chloride 5.623/~mol; lead chloride 7.943 #mol; and zinc chloride 13.331 #mol. Figure 9 shows that when the green mussels were deprived of food the activity of sucrase in the digestive gland started to fall, and the trend continued for 10 days. The activity of sucrase in the digestive gland of those starved for eight days was only half of that in the feeding mussels.

c IO0~"X

~ 80

~N ¥ =110-7,13X o

\ <

~g 2o

Starvation Period (Days) Fig. 9. Effect of starvation on sucrase activity.

DISCUSSION The optimal pH of the sucrase from the digestive gland of Perna viridis is very close to that of the slug Arian ater (pH 5.9) (Evans and Jones, 1962) and the snail Achatinafuliea (pH 5.6) (Tea, 1983). At the pH of the digestive gland (pH 6.19) the sucrase would still be very active, > 80?/0 of the maximum activity at optimal pH. The lowest optimal pH of 4.0 for ct-glucosidase was recorded in the Atlantic deep-sea scallop Placopecten magellanicus (Wojtowicz, 1972). The Km of the green mussel sucrase was found to be 0.0140 M, which is slightly higher than that of the slug Arian ater (0.0128 M), but slightly lower than that of the snail Achatina fulica (0.0160 M) (Evans and Jones, 1962; Tea, 1983). It is interesting that although the maximum temperature for the sucrase of the green mussel was only 40°C, it could still retain about 78% of its maximum rate at 65°C. This indicates that the sucrase is rather thermostable. In Achatina fulica the maximum temperature of the sucrase was 45°C, but the rate at 55°C was still close to the maximum (Tea, 1983). Both are tropical animals and so their sucrases have similar thermostabilities. The activation energy of the sucrase from the digestive gland of the green mussel, in the temperature range 20-40°C, was high, 10,600 cal/mol, in the slug it was 4500 cal/mol (Evans and Jones, 1962) and in the tropical snail, only 4100 cal/mol (Tea, 1983). The inhibitory effects of different bivalent salts on the activity of suerase varied quite a lot, but comparable data could not be found for sucrase of other bivalves. The sucrase of the house cricket was more resistant to the inhibitory effect of mercury chloride than the sucrase of the green mussel, the 50% inhibition concentration for the former was 59 ~mol (Tea and Woodring, 1989 (cf. 2.239 pmol). The trehalase of the tobacco hornworm larvae, and that of the grasshopper thoracic muscle, were also much more resistant to the inhibitory effects of mercury chloride, cadmium chloride, zinc chloride and copper sulphate than the sucrase of the green mussel (Dahlman, 1971; Tea and Heng, 1987). Our data confirmed the observation of Reid (1968) that starvation resulted in the reduction of enzyme

Perna sucrase production, and that feeding activity served as a stimulus for enzyme production. Thus, the continuous filter-feeders show the same response to starvation as the periodic feeders. It is certainly against the welfare of the animal to continue producing large quantities of enzymes after feeding stops because the proteins needed for the production of enzymes can be used to meet the needs for other more immediate requirements~ REFERENCES

Dahlman D. L. (1971) Purification and properties of trehalase from tobacco hornworm larvae. J. Insect Physiol. 17, 1677-1687. Evans W. A. L. and Jones E. G. (1962) Carbohydrases in the alimentary tract of the slug Arion ater L. Comp. Biochem. Physiol. 5, 149-160. Finney D. J. (1971) Probit Analysis (3rd edn), pp. 1-327. Cambridge University Press, Cambridge. Fox D. L. and Marks G. W. (1936) Digestive enzymes of Mytilus. Bull. Scripps. Inst. Oceanog. 4, 29-47. Haldane J. B. S. (1965) Enzymes, pp. 1-15. Longrnan, Green and Co., London. Kamat D. N. (1957) Carbohydrate digestion in the mussel: a brief study of amylase. J. Univ. Bombay. 24, 112-116. Morton R. K. (1955) Methods of extraction of enzymes from animal tissues. Meth. Enzym. 1, 25-51.

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Mozo P. S., Rama, M. F., Pernas, R. V. and Amil M. R. (1977) Purification and properties of two enzymatic forms of 3-N-acetylglucosaminidase from Mytilus edulis. Comp. Biochem. Physiol. 58, 29-34. Oser (1965) Hawk's Physiological Chemistry (14th edn), pp. 40-43. McGraw Hill, New York. Owen G. (1974) Feeding and digestion in Bivalvia. Adv. Comp. Physiol. Biochem. 5, 1-35. Reid R. G. B. (1968) The distribution of digestive tract enzymes in lamellibranchiate bivalves. Comp. Biochem. Physiol. 24, 727-744. Sumner J. B. (1925) A more specific method for the determination of glucose. J. biol. Chem. 65, 393-395. Teo L. H. (1983) The liver sucrase of the African giant snail, Achatina fulica Bowdich. J. Singapore natn. Acad. Sci. 10-12, 38~,1. Teo L. H. and Heng S. K. (1987) The trehalase of the grasshopper Valanga nigricornis. Comp. Biochem. Physiol. 87B, 373-378. Teo L. H. and Woodring J. P. (1989) The invertase of the house cricket Acheta domesticus L. (Orthoptera: Gryllidae). Comp. Biochem. Physiol. 93, 643446. Yonge C. M. (1935) On some aspects of digestion in ciliary feeding animals. J. mar. biol. Ass. UK 20, 341-346. Wojtowicz M. B. (1972) Carbohydrases of the digestive gland and the crystalline style of the Atlantic deepsea scallop, Placopecten magellanicus. Comp. Biochem. Physiol. 43, 131-141.