Characteristics of glycogen synthase activity in the oyster, Crassostrea virginica Gmelin

Comp. Biochem. Physiol. Vol. 90B, No. 2, pp. 361-365, 1988 Printed in Great Britain

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CHARACTERISTICS OF G L Y C O G E N SYNTHASE ACTIVITY IN THE OYSTER, C R A S S O S T R E A VIRGINICA G M E L I N MARY L. SWIFT,* THOMAS P. THOMASand CHERYL L. HUMPHREY Department of Biochemistry, College of Medicine Howard University, Washington, DC 20059, USA

(Received 10 July 1987) Abstract--1. Glycogen synthase I and D activities were found in homogenates of oyster adductor muscle and digestive diverticula. 2. Upon starvation the ratio of GSI/GS(I + D) increased in both tissues in summer animals. 3. Kinetic characteristics of the purified enzyme from the adductor muscle were found to be: KmtuDP~)= 0.86 raM, Kacc~p)= 0.15 mM for the D form of the enzyme; Km(uD~) = 7.09 mM for the I form of the enzyme. 4. Addition of glucose to muscle homogenates resulted in increases in the GSI/GS(I + D) ratio within 10 rain.

INTRODUCTION Seasonal variations in tissue glycogen content have been observed in several animal phyla (Pessacq and Gagliardino, 1975). Perhaps the most marked variation in glycogen content occurs in marine bivalve molluscs where extremes of 2.5--40% dry wt have been measured (Galtsoff, 1964; Pieters et al., 1978). In the higher animals, one may speculate that the increase in glycogen content aids thermoregulation during the cold months. However, the seasonal variation in glycogen content of the marine bivalves is almost certainly related to their reproductive cycle (Galtsoff, 1964; Thompson, 1977; Pieters et al., 1978). In addition, these large glycogen reserves permit the marine bivalves to survive prolonged periods of starvation (Riley, 1976) and anoxia (Zandee et al., 1980). Despite this central role of glycogen in the metabolic strategy of these animals, only a few reports have appeared in which glycogen metabolism of bivalve molluscs has been examined (L.-Fando et al., 1972; Cook and Gabbott, 1978; VazquezBaanante and Rosell-Perez, 1979; Ebberink and Salimans, 1982; Gabbott and Whittle, 1986; Whittle and Gabbott, 1986). The primary site of control of glycogenesis is the regulation of UDP-glucose: ~t1, 4-glucan ,,-4 glucosyltransferase (E.C.2.4.1.11), glycogen synthase (GS) (Danforth, 1 9 6 5 ; Van de Werve and Jeanrenaud, 1987). This enzyme has been extensively studied in mammalian tissues, especially liver, skeletal and heart muscle (Stalmans and Hers, 1973; Hers, 1976). However, GS from molluscan sources has been systematically investigated only from the mantle of Mytilus edulis (Cook and Gabbott, 1978; Gabbott and Whittle, 1986; Whittle and Gabbott, 1986) and from the cephalopedal region of Biomphalaria glabrata (Schwartz and Carter, 1982). This paper reports on aspects of GS activity in the American oyster, Crassostrea virginica Gmelin. We *To whom correspondence should be addressed. C,B.P. 90/2B--H

have found that two forms of GS, GSI and GSD, are present in both adductor muscle and digestive diverticula. The ratio of GSI to total GS activity in these tissues increases upon prolonged starvation. GS from the adductor muscle was partially purified and characterized. Also presented is evidence for /n vitro regulation of GS by glucose. MATERIALS AND METHODS Oysters (Crassostrea virginica Gmelin), 4-5 cm in height were obtained on the day of harvest from a commercial supplier (Chesapeake Bay Oyster Culture Co., Shady Side, MD). Animals not killed immediatelywere scrubbed with a nylon brush and placed in aquaria under 12 ppt artificial sea-water (Instant Ocean, Aquarium Systems, Eastlake, OH) at 15°C (Swift et al., 1975). No attempt was made to feed the oysters. Aquaria were cleansed and the saline was changed every three days. Levels of GS activity were measured in digestive diverticula and adductor muscle homogenates prepared from freshly harvested oysters (fed) and from animals maintained unfed in the laboratory for 25 days. The excised tissues were homogenized at 4°C in 0.25 M sucrose, containing 0.15 M NaF and 5 mM EDTA at pH 7.4, using an Omni-Mixer (DuPont Company, Sorvall Instruments Division, Newton, CT) at a setting of 7 for 2 rain. Adductor muscle homogenares were prepared at a 2:1 (w/v) ratio whereas digestive diverticula homogenates were at a 1:1 (w/v) ratio. Prior to enzymatic activity assay samples were diluted with a volume of 0.5 M sucrose containing 0.3 M NaF, 5 mM EDTA and 0.1 M Tris-HCl, pH 8.15 (Witters and Avruch, 1978). The radiometric method of Thomas et al. (1968) was employed to measure GSI (glucose (,-phosphate omitted) and total GS (with l0 mM glucose 6-phosphate) activities. Glycogen synthesized during the enzymatic reaction was precipitated onto filter paper and washed. These filter papers were placed in l0 ml of a water miscible cocktail (Hydromix, Yorktown Research, South Hackensack, NJ) and monitored in a liquid scintillation counter (Beckman LS9000, Beckman Instruments, Irvine, CA) using an automatic quench correction program. Data were analyzed for significance using the Student's t-test. Glycogen synthase was prepared from adductor muscle using the procedure outlined by Cook and Gabbott (1978)

361

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except that prior to chromatography on ConA-Sepharose the homogenate was centrifuged at 105,000g for 90 min at 4°C (Rosell-Perez and Lamer, 1964). The pellet was resuspended in 50 mM Tris-HC1, pH 7.5 containing 25 mM KF and 1 mM dithiothreitol (DTT) and applied to the CortA-Sepharose column. Enzymatic activity was estimated by the method of Thomas et al. (1968). Protein was monitored according to Lowry et al. (1951). Kinetic data were fitted to the best straight line using linear regression analysis. The effect of glucose on levels of GSI activity in homogehates of adductor muscle from freshly harvested oysters was examined. Tissue was homogenized at a 1:3 (w/v) ratio in 50 mM Tris-HCl, pH 7.5 containing 12.5 mM DTT. After centrifugation at 50008 for 10min at 4°C, samples of the supernatant liquid were incubated with glucose at final concentrations of 0, 10 and 20 mM. At various time intervals, aliquots were withdrawn, and immediately mixed with 4 volumes of 50 mM Tris-HCl, pH 7.5 containing 20 mM EDTA, 25mM KF, 10mM DTT at 0°C. These were assayed for GS activity as outlined above. UDP-UJ4C-D-glucose was obtained from ICN Radiochemicals (Irvine, CA). Except as specified, all other reagents and chemicals were purchased from Sigma Chemical Co. (St. Louis, MO).

RESULTS AND DISCUSSION G l y c o g e n synthase I a n d D activities were f o u n d in h o m o g e n a t e s o f the a d d u c t o r muscle a n d digestive diverticula o f the A m e r i c a n oyster. T h e percentage o f G S expressed as G S I in freshly collected s u m m e r animals was o n e - h a l f to one-third o f t h a t f o u n d in fall or spring ( a d d u c t o r muscle only) oysters. U p o n s t a r v a t i o n in the l a b o r a t o r y the percentage o f G S I activity in the s u m m e r groups, i.e. those oysters with

lOW initial ratios o f G S activity, increased significantly to a b o u t the levels expressed in the fall or spring animals. This t r e n d in G S activity ratios is consistent with t h a t observed in the liver from rats subjected to longer periods of fasts. In this case, as liver glycogen levels declined G S I increased ( S h i k a m a et al., 1980). Overall, the increases in the G S I / G S (I + D) ratio were m o r e evident in the a d d u c t o r muscle, averaging 376 vs 150% for the digestive diverticula (Table 1). These d a t a suggest t h a t there is a physiologic limit o n the in vivo ratio of G S I to total G S activity. C h a n g e s in G S activity ratios were due to higher G S I activity (Table 2) rather t h a n decreases in total G S activity (Table 3). Protein c o n t e n t of the tissues did n o t change significantly (data n o t shown). Similarities to the seasonal p a t t e r n o f M . edulis G S activity ( G a b b o t t a n d Whittle, 1986) were seen in oysters. In b o t h bivalves the percentage of G S activity present as G S I is low in summer. In the spring, just p r i o r to spawning, this ratio increases. In fall the ratio o f G S I to total G S activity was elevated. This m i g h t be expected as d u r i n g the early fall the oyster is replenishing its glycogen reserve after spawning. As the increases in the G S I to G S (I + D) ratio were m o r e m a r k e d in a d d u c t o r muscle homogenates, enzyme from this tissue was subjected to further study. Typically 60--70-fold purification o f G S was achieved. T h e p r e p a r a t i o n s were stable for 2-3 days at 4°C. Purified enzyme could be o b t a i n e d as 7 0 - 7 5 % I form w h e n all solutions were devoid of fluoride. P r e p a r a t i o n s used to determine kinetic c o n s t a n t s a n d p H o p t i m a expressed activity which was at least 7 0 % o f the I or D form (Table 4). As has been observed for G S f r o m M . edulis ( C o o k a n d G a b b o t t , 1978) a n d

Table 1, Changes in the glycogen synthase I/(I + D) activity ratio in unfed oysters GSI/GS (I + D) Tissue Collection Day 1 Day 25 % Change (% _ SD) Adductor muscle Summer 7.98 +_6.5 25.4 _+5.2* 218 2.80 + 2.1" 47.8 --!-_28* 1607 Fall 25.5 _ 3.6 25.0 +_16 2 Spring 20.5 + 6.7 31.7 +_ 13 55 34.2+ 12 33.9+ 15 -1 Digestive diverticula Summer 4.41 + 1.8t 15.1 + 2.2t 242 5.66 _+5.3t 34.2 ± 17t 504 Fall 16,1 + 2.1 12.5 _ 4.5 -22 Spring 17.9 _ 8.6 23.4 ± 12 31 8.60 + 2.0 8.20 __.1.5 -5 Number of animals in each group was 4. *Means are significantly different at the P ~<0.05 level. tMeans are significantly different at the P ~<0.01 level. Table 2. Changes in glycogen synthase I activity in unfed oysters Glycogen synthase I Tissue Collection Day I Day 25 0tmol/min per mg protein + SD) Adductor muscle Summer 0.82 + 0.29* 2.74 _+0.85* 1.62 + 0.46* 14,1 + 0.82* Fall 1.20 ±_0.16 0.88 + 0.30 Spring 2.24 + 1.17 1.90 + 0.78 1.92 +_0.57 2.24 ± 0.57 Digestive diverticula Summer 0.84 + 0.38* 2.23 _ 0.33* 0.65 + 0.26* 6.90 +_0.84* Fall 1.25 + 0.11 0.72 + 0.31 Spring 2.45 + 1.05 2.28 __.0.25 0.76 + 0.18 1.02 _+0.18 Number of animals in each group was 4. *Means are significantly different at the P ~<0.01 level.

% Change 235 770 - 27 - 15 17 166 960 --43 -7 35

Glycogen synthase from oyster

363

Table 3. Changes in total glycogen synthase (I + D) activity in unfed oysters Tissue

Glycogen synthase (I + D) Day I Day 25

% Change

(/~mol/min per mg protein + SD) 15.5 -I- 11 10.7 ± 2.6 37.1 ± 12 30.7 ± 7.3 4.77 ± 0.64 4.10 ± 1.3 5.33 + 0.48* 6.71 + 0.21" 9.42 ± 0.35 9.55 ± 2.2

-31 - 17 - 14 26 1

Collection

Adductor muscle

Summer Fall Spring

Digestive diverticula

Summer Fall Spring

19.2 ± 3.6 13.6 ± 2.1"[" 8.19 ± 0.66 12.2 ± 2.7 9.18 ± 3.0

15.2 ± 4.1 26.9 ± 8.1t 6.97 ± 1.7 8.33 ± 1.8 13.0 ±4.6

- 21 98 - 15 -6 -32

Number of animals in each group are 4. *Means are significantly different at the P ~<0.01 level. tMeans are significantly different at the P ~<0.02 level.

rabbit muscle (Rosell-Perez and Larner, 1964) the Km(uDm)of oyster GSD is one tenth of that found for GSI. The effect of nucleotides, phosphate and sulfate on GS activity was examined (Table 5). As found for GS from rat muscle (Piras et al., 1968), the D form of GS activity was markedly reduced in the presence of nucleotides. These results are similar to those found for GS from mammalian liver or M. edulis mantle. In these tissues the expression of the D form of enzyme activity is more subject to inhibition by various nucleotides and phosphate (DeWulf et al., 1968; Stalmans and Hers, 1973; Cook and Gabbott, 1978). In rabbit muscle, phosphate and sulfate have little effect on GS activity (Rosell-Perez and Lamer, 1964). Oyster adductor muscle GSD activity was stimulated by phosphate and sulfate (Table 5). Exposure of crude homogenates of adductor muscle to glucose resulted in an increase in GSI activity (Fig. 1). Over the time interval of the experiment there was no change in total GS activity expressed. The increase in GSI activity observed appears to be related to the concentration of glucose added to the homogenate. Treatment of mammalian Table 4. Characteristics of oyster adductor muscle glycogen synthase Glycogen synthase D I

KmCvDm~(raM) Ko<~p)(mM) Vm~ (nmol/min per mg protein) pH optimum

0.86 0.15 25.5 7.5-7.8

7.09 -16.4 7.0-7.3

liver preparations with exogenous glucose causes an increase in GSI activity. This effect occurs rapidly, beginning within 5 rain of the treatment and lasting 40-60rain (Stalmans et al., 1974; Witters and Avruch, 1978). As reported herein, the response of GS in oyster muscle preparations is within this time frame, however the duration of the response is as yet unknown. GS activity in M. edulis mantle homogenares remained elevated for up to 3 hr (Whittle and Gabbott, 1986). A glucose effect on GS from muscle tissue has not been previously reported. Indeed evidence from mammalian studies would suggest that skeletal muscle is incapable of such a response. GS activation in liver due to glucose is mediated through phosphorylase a which in the presence of glucose binds synthase phosphatase. In skeletal muscle, synthase phosphatase is unaffected by phosphorylase a. Hence, mammalian muscle GS would not be activated by increases in glucose concentration (Mvumbi et al., 1983; Alemany and Cohen, 1986).

6°I

50

4O 3o

Table 5. Effect of nucleotides and selected ions on oyster adductor muscle glycogen synthase activity Effector CTP UDP UTP AMP ADP ATP cAMP Pi

SO~-2 Mg + 2 Mg+2/ATP

Concentration (mM) 10 10 l0 10 10 5 5

Glycogen synthase -G6P 10mM G6P 51 1 3 79 54 22 79

(% activity)* 84 16 54 118 100 I 13 103

5

86

120

5 20 20/10

106 66 70

120

113 128

*Activity observed in the absence of effectors was set at 100%.

-IO

o

J

I

t

IO

20

30

Time (rain)

Fig. 1. Changes in glycogen synthase l activity upon incubation of adductor muscle homogenates with various concentrations of glucose (O, 0.0 m M glucose, 0 , 10 mM glucose and I'q, 20 mM glucose). At the intervals indicated samples were withdrawn from the incubation mixtures and assayed for enzymatic activity. All values expressed relative to GSI activity found at time zero.

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MARY L. Swwr et al.

The rapidity of the glucose effect on GS in oyster and M. edulis preparations (Whittle and Gabbott, 1986) is to be contrasted with the effect of insulin on glycogen synthesis in invertebrates. When molluscs, which were previously found to contain insulin-like proteins, were challenged by insulin administration there was a response only after about 4 hr as measured by changes in glycogen synthesis or hemolymph glucose levels. For example, in order to observe an insulin effect in the gastropod, Strophocheilus oblongus, the following conditions were employed. A glucose load was administered 4 hr after insulin injection (25 IU/kg body wt). Hemolymph glucose levels were monitored and found to be about 30% lower than the controls for 6--12 hr after the initial insulin treatment. This regime also led to increased muscle and hepatopancreas glycogen content (Marques and Falkmer, 1976). Similar delays in metabolic response have been observed in the bivalves Anodonta cygnea, Unio pictorum, Mytilus galloprovinciallus and Chlamys glaba ponticus (Plisetskaya et al., 1978, 1979). In the latter studies, GSI activity increased 7 hr after insulin injection. These results are to be contrasted with those obtained when M. edulis was subjected to insulin treatments. The intact animal was treated with insulin or anti-insulin serum whereas mantle tissue slice preparations were treated only with insulin. In these studies, no change in GSI activity was expressed (Gabbott and Whittle, 1986; Whittle and Gabbott, 1986). Furthermore, insulin apparently is not absolutely necessary for a response of GS in mammals. In rat liver the increase of GSI noted in the postprandial state is not affected by injecting the animal with anti-insulin serum (Shikama et aL, 1980). In view of these very slow responses to insulin contrasted with the rapid response of oyster GS upon exposure to glucose, one may postulate that the regulation of glycogen metabolism in marine invertebrates has a significant intracellular component. This regulation would arise from alterations in the activity of GS as affected by fluctuations of the key metabolites such as glucose 6-phosphate and the adenine nucleotides. The mechanism of this regulation might be similar to that suggested for mammalian liver glycogen metabolism (Van de Werve and Jeanrenaud, 1987). The "energy-charge" of the cell (the ratio of concentrations of ATP to AMP and ADP) could serve to aid in control of the rate of glycogenesis. Hence there might be at least two mechanisms for regulation of GS activity. Increasing glucose concentration could mediate a shift in amounts of GSD and GSI whereas the intracellular concentration of the nucleotides would serve in fine regulation of the pre-existing enzymatic forms (Piras et al., 1968). Any inhibitory effect of these nucleotides could be surmounted by intracellular increases in glucose 6-phosphate concentration (Table 5 and Fig. 1). This general strategy may be particularly attractive for the marine bivalve molluscs. Their open circulatory system requires that the concentration of intracellular metabolites be reflected in the hemolymph (Gabbott and Whittle, 1986). Thus the diurnal shifts in the metabolic machinery may be determined by nutritional status. Hormonal responses then might be reserved for the more profound and longer lasting

metabolic changes required in moving from somatogenesis to gametogenesis and vitellogenesis. Acknowledgements--This work was funded in part by a grant from the National Science Foundation (PCM8118227). Thanks are due to Dr R. H. Pointer for his many suggestions and technical advice during the conduct of this project. REFERENCES

Alemany S. and Cohen P. (1986) Phosphorylase a is an allosteric inhibitor of the glycogenand microsomal forms of rat hepatic protein phosphatase-1. FEBS Letts 198, 194-202. Cook P. A. and Gabbott P. A. (1978) Glycogen synthase in the sea mussel Mytilus edulis L.--I. Purification, interconversion and kinetic properties of the I and D forms. Comp. Biochem. Physiol. 60B, 419--421. Danforth W. H. (1965) Glycogen synthetase in skeletal muscle, interconversion of two forms and control of glycogen synthesis. J. biol. Chem. 240, 588-593. De Wulf H., Stalmans W. and Hers H. G. (1968) The influenceof inorganic phosphate, adenosine triphosphate, and glucose 6-phosphate on the activity of liver glycogen synthase. Eur. J. Biochem. 6, 545-551. Ebberink R. H. M. and Salimans M. (1982) Control of glycogenphosphorylase activity in the posterior adductor muscle of the sea mussel Mytilus edulis. J. Comp. Physiol. 148, 27-33. Gabbott P. A. and Whittle M. A. (1986) Glycogen synthetase in the sea mussel Mytilus edulis L.--II. Seasonal changes in glycogen content and glycogen synthetase activity in the mantle tissue. Comp. Biochem. Physiol. $3B, 197-207. Galtsoff P. S. (1964) The American Oyster, Crassostrea virginica Gmelin. Fishery Bulletin of the Fish and Wildlife Service, Vol. 64, p. 386. U.S. Dept. of Interior. Hers H. G. (1976) The control of glycogen metabolism in the liver. Ann. Rev. Biochem. 45, 167-189. L.-Fando J. J., Garcia-Fernandez M. C. and R.-Candela J. L. (1972) Glycogen metabolism in Ostrea edulis (L.)---Factors affecting glycogen synthesis. Comp. Biochem. PhysioL 43B, 807-814. Lowry O. H., Rosebrough N. J., Fair A. L. and Randall R. J. (1951) Protein measurement with the Folin phenol reagent. J. biol. Chem. 193, 265-275. Marques M. and Falkmer S. (1976) Effects of mammalian insulin on blood glucose level, glucose tolerance and glycogencontent of musculature and hepatopancreas in a gastropod mollusk, Strophocheilus oblongus. Gen. Comp. Endocr. 29, 522-530. Mvumbi L., Dopere F. and Stalmans W. (1983) The inhibitory effect of phosphorylase a on the activation of glycogen synthase depends on the type of synthase phosphatase. Biochem. J. 212, 407-416. Pessacq M. T. and Gagliardino J. J. (1975) Glycogen metabolism in muscle: Its circadian and seasonal variations. Metabolism 24, 737-743. Pieters H., Kluytmans J. H., Zurburg W. and Zandee D. I. (1978) Proc. Eur. Mar. Biol. 13th Syrup., pp. 285-292. Pergamon Press, Oxford. Piras R., Rothman L. B. and Cabib E. (1968) Regulation of muscle glycogen synthetase metabolites. Differential effects on the I&D forms. Biochemistry 7, 5(~66. Plisetskaya E., Kazakov V. K., Soltitskaya L. and Leibson L. G. (1978) Insulin-producing cells in the gut of freshwater bivalve molluscs Anodonta cygnea and Unio pictorum and the role of insulin in the regulation of their carbohydrate metabolism. Gen. Comp. Endocr. 35, 133-145. Plisetskaya E. M., Soltitskaya L. P. and Leibson L. G. (1979) Role of insulin in the regulation of metabolism in

Glycogen synthase from oyster marine bivalve mollusks. Zh. EvoL Biokhim. Fiziol. 15, 288-94 (Chem. abst. 91: 102580k). Riley R. T, (1976) Changes in the total protein, lipid, carbohydrate and extracellular fluid free amino acids of the Pacific oyster, Crassostrea gigas, during starvation. Proc. ham. Shellfish. Assoc. 65, 84-90. Rosell-Perez M. and Lamer J. (1964). Studies on uridine diphosphogiucose-~-glucan transglucosylase--IV. Purification and characterization of two forms from rabbit muscle. Biochemistry 3, 75-81. Schwartz C, F. and Carter C. E. (1982) Properties of glycogen synthase and phosphorylase from Biomphalaria glabrata (mollusca). J. Parasit. 68, 228-235. Shikama H., Yajima M. and Ui M. (1980) Glycogen metabolism in rat liver during transition from the fed to fasted states. Biochim. biophys. Acta 631, 278-288. Stalmans W., DeWulf H., Hue L. and Hers H. G. (1974) The sequential inactivation of glycogen phosphorylase and activation of glycogen synthetase in liver after the administration of glucose to mice and rats. Eur. J. Biochem. 41, 127-134. Stalmans W. and Hers H. G. (1973) The Enzymes, 3rd edn, Vol IX, pp. 309-361. Academic Press, New York. Swift M. L., Conger K., Exler J. and Lakshmanan S. (1975) Uptake of glucose and orthophosphate by the American oyster. Life Sci. 17, 1679-1684.

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Thomas J. A., Schlender K. K. and Lamer J. (1968) A rapid filter paper assay for UDP glucose-glycogen glucosyltransferase, including an improved biosynthesis of UDP-14C-giucose. Analyt. Biochem. 25, 486-499. Thompson R. F. (1977) Blood chemistry, biochemical composition, and the annual reproductive cycle in the giant scallop, Placopecten magellanicus, from southeast Newfoundland. J. Fish. Res. Bd. Can. 34, 2104-2116. Whittle M. A. and Gabbott P. A. (1986) Glycogen synthetase in the sea mussel, Mytilus edulis L.--III. Regulation by glucose in a mantle tissue slice preparation. Comp. Biochem. Physiol. 83B, 209-214. Witters L. A. and Avruch J. (1978) Insulin regulation of hepatic glycogen synthase and phosphorylase. Biochemistry 17, 406-410. Van de Werve G. and Jeanrenaud B. (1987) Liver glycogen metabolism: an overview. Diabetes/Metabolism Rev. 3, 47-78. Vazquez-Baanante I. and Rosell-Perez M. (1979) In vitro studies of Pecten maximus glycogen phosphorylase and the interconversion of their forms. Comp. Biochem. Physiol. 62B, 381-387. Zandee D. I., Holwerda D. A. and deZwaan A. (1980) Animals and Environmental Fitness, pp. 185-206. Pergamon Press, New York.