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Quantitative studies on trehalose in the oyster, Crassostrea virginica Gmelin
Comp. Biochem. Physiol., 1967, VoL 23, pp. 621 to 629. Pergamon Press. Printed in Great Britain
QUANTITATIVE STUDIES ON TREHALOSE IN THE OYSTER, CRASSOSTREA VIRGINICA GMELIN D A V I D G. BADMAN* Department of Zoology, University of Florida, Gainesville, Florida 32601, U.S.A.
(Received 20 May 1967) Abstract--1. Trehalose was identified in the oyster, Crassostrea virginica Gmelin, and found to be present throughout the period of one year in amounts ranging from 0.188 mg/g wet wt. to 3"300 mg/g wet wt. 2. Trehalose levels remained constant except for February and July, when they were significantly elevated. 3. The amount of trehalose relative to glycogen is normally very small except immediately after spawning, when the glycogen level is drastically reduced. INTRODUCTION TIu~H~a~OSE, l(~-v-glucopyranosyl) c~-v-glucopyranoside, long believed to be confined largely to fungi and yeast, has been shown to be a major sugar in insect hemolymph (Wyatt & Kalf, 1957). Falrbairn (1958) identified trehalose in many other invertebrate species. In general, the trehalose of arthropods appears to be a storage compound. It is a glucose source during flight in Musca domestica (Sacktor, 1955) and in Phorraia regina (Clegg & Evans, 1961). The dormant embryo of the brine shrimp, Artemia sally, contains a large trehalose reserve which is converted to glycogen at the end of the dormant period (Clegg, 1956). When the silkworm is subjected to exhaustion, hemolymph trehalose decreases rapidly (Duchateau & Florkin, 1959). Saito (1963) reported the presence of a mechanism to regulate the hemolymph sugar level in the silkworm, Bombyx ~ r i . As the original hemolymph trehalose is used, its concentration is kept constant by the biosynthesis of new trehalose from carbohydrate stored as glycogen in the fat body. In the cockroach, the balance between hemolymph trehalose and fat body glycogen appears to be regulated by hormones (Steele, 1963). It is evident that in insects there is a close relationship between trehalose and glycogen. It has long been known that molluscs contain large amounts of glycogen, and an annual cycle of this carbohydrate has been observed in oysters (Galtsoff, 1964). In view of this, a study of trehalose content of oysters throughout the period of 1 year was undertaken. * This paper is based on a thesis submitted to the Graduate School Of the University of Florida in partial fulfdlment of the requirements for the degree of Master of Science in the Department of Zoology. 621
622
DAVIDG. BADMAN
MATERIALS AND METHODS Oysters, Crassostrea virginica Gmelin, used in this study were collected from the dock at the University of Florida Marine Laboratory on Seahorse Key, near Cedar Keys, Florida. Collections were made in the middle of each month beginning April, 1965, and ending March, 1966. The shells of oysters from Apalachicola Bay reach 100 mm in height after one summer of growth (Ingle, 1950). Six oysters of this approximate size (75-110 ram) were selected each month for analysis. The oysters were placed in buckets and transported 50 miles to Gainesville where they were frozen as soon as possible, usually within 2-3 hr after collection.
Preparation of extracts The oysters were thawed and their shells measured. The shucked, drained tissue was weighed and then homogenized for 5 rain in 70% (v/v) ethanol in a Waring blender. The homogenate was centrifuged in a Servall centrifuge (Type SS-1 head) at 50 V (ca. 10,300 g) for 10 min. The supernate was evaporated to dryness by warming on a hot plate. The water-soluble portion of the dried supernate was redissolved in distilled water at room temperature, and the volume adjusted so that 1 ml represented 100 mg of the original wet wt of the oyster. The adjusted extracts were desalted in a Torbal-BLT Chromatographic Desalting Apparatus (Model CD-1) using a Permutit C-20 cation resin membrane and A-20 anion resin membrane (Blainey and Yardley, 1956). Desalted extracts were then frozen for later use.
Chromatography and quantitative analysis Thin layer chromatography was used to separate trehalose from other carbohydrates present in the desalted extracts, using prespread Eastman Chromagram Sheets (Type K301R2) and Developing Apparatus. Chromatograms were activated by heating for 30 min at 100°C, and were then spotted with 100/zg authentic D(+) trehalose dihydrate from Mann Research Laboratories, Inc. and with the desalted extracts. To prepare an extract for spotting, 0"5 ml of the desalted extract was evaporated to dryness by warming in a sand bath, and then redissolved in 50/~1 of water. Of this, 20/zl were spotted on each chromatogram. N-butanol: ethanol : water (4 : 1 : 1.9) was the solvent used to separate the components of the extracts. When the solvent front had moved 10 cm, the chromatograms were removed, dried at room temperature and cut into strips corresponding to knowns (authentic trehalose), unknowns (oyster extracts) and blanks (distilled water). In order to visualize the spots, the dried chromatogram strips were dipped in AgNO3 reagent (prepared by diluting 0.1 ml of saturated aqueous AgNO3 to 20 ml with acetone and then dissolving the precipitated AgNO3 by dropwise addition of water). The chromatogram strips were then dipped in 0"5 N NaOH in aqueous ethanol (made by diluting saturated NaOH to 0.5 N with absolute ethanol). Reducing sugars form black spots at room temperature, while trehalose forms
~TgEHALOSBIN CRASSOSTREA VIRGINICA
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light brown spots. This procedure detected 0.5/~g trehalose in a solution containing 50/~g per ml. Trehalose was located on one known strip and one unknown strip to ascertain its position on the remaining three strips representing each oyster. From the unstained strips and from the blank, a rectangular area corresponding to the standard trehalose position was cut out and eluted with 1.0 ml of water. Quantitative elution with gravity was accomplished in 15-20 rain with the strip fed continuously by a wick inserted into a container of distilled water. Eluates of the unknown strips were negative for glucose when analyzed with the glucose oxidase system, Glucostat, obtained from Worthington Biochemical Corporation, showing that there had been adequate separation of trehalose from glucose. The eluates were analyzed for trehalose with the anthrone procedure of Carroll, Longley & Roe (1956).
Criteria of trehalose presence T h e presence of trehalose in the extract was proven by several criteria used by Clegg & Filosa (1961). (1) Strips were sprayed with 0"5% (w/v) sodium metaperiodate and allowed to dry. After drying, the strips were stained with alkaline silver nitrate as before. Treatment with sodium metaperiodate oxidizes the ~,~ bond of trehalose to form glucose, which then gives the characteristic reaction of a reducing sugar with silver nitrate. T h e results, shown in Table 1, indicate that the unknown sugar in the "trehalose" position (i.e. with R/the same as that of authentic trehalose) is non-reducing. TABLE 1----COLORREACTIONWITH ALKALIHESILVERNITRATE Chromatogramed sample
Not treated with Na m-periodate
Treated with Na m-periodate
Authentic glucose
Black spot
Black spot
Authentic trehalose
immediately Brown spot after 30 sec
immediately Black spot immediately
Black spot immediately
Black spot immediately
Brown spot after 30 sec
Black spot immediately
Unknown: "glucose" position "trehalose" position
(2) The unknown eluate reacts with anthrone. (3) T h e spot corresponding to trehalose on the unknown strip was eluated and rechromatogramed in four different solvent systems: (a) n-butanol:ethanol : acetone : water (5 : 4 : 3 : 20), (b) n-propanol : ethyl acetate : water (7 : 1 : 2), (c) ethyl
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acetate : pyridine : water ( 8 : 2 : 1), and (d) n-butanol : aceticacid : water(4 : 1 : 5). In each of these systems, the unknown eluted spot had the same R t as the authentic trehalose, and in each no other spot appeared, showing that the eluate was pure. (4) The unknown eluate was hydrolyzed quantitatively to glucose by trehalas¢. Trehalase was prepared by a procedure modified from Kalf & Rieder (1958). Approximately 1000 three-day-old house flies, Musca domestica, weighing a total of 27.82 g were used. The trehalase obtained was dissolved in 10"1 ml of 0.10 M phosphate buffer, pH 6.8, and stored at 2°C. It was found to be free of maltase activity, and hydrolyzed authentic trehalose to glucose with an average yield of 95 per cent. The assay mixture contained 0.1 ml of 0"10 trehalose or other substrates (distilled water blank, maltose or unknown), 0.35 ml of 0.10 M acetate buffer (pH 5.5), and 0.05 ml of the enzyme. After a 15 rain incubation at 30°C, the reaction was stopped by immersing the tubes in a boiling water bath for 3 rain. The tubes were cooled in an ice bath, the precipitate removed by centrifugation, and a sample removed for determination of glucose. A zero-time control was always run to correct for any glucose in the trehalose or enzyme preparation. Unknown eluates were analyzed for trehalose by this procedure and by the anthrone procedure. In all cases the results of the two methods were identical.
RESULTS
Thin layer chromatography proved to be a satisfactory technique for isolating trehalose from oyster extracts. In a warm room (above 20°C), the plates could be developed in 2-3 hr. Separation of monosaccharides from disaccharides was good. The R/ of glucose averaged 0.35, that of trehalose, 0.22. Recovery of authentic trehalose by elution from the chromatogram by gravity and analysis with anthrone averaged 95 per cent. Trehalose was found in all extracts. The amounts of trehalose found ranged from 0.188 mg/g wet wt. to 3.300 mg/g wet wt. Although oysters having reasonably uniform shell sizes were selected for analysis, the wet wt. of the tissue varied from 1.8 g to 151.1 g. There was no significant correlation between wet weight of tissues and amount of trehalose present (r = 0.0787, p >0"1). Figure 1 gives a summary of the results. From August until January., a t i m e of high glycogen synthesis and storage (Galtsoff, 1964), the trehalose concentration is nearly constant, averaging 0"600 mg trehalose/g wet wt. From January to March, the trehalose concentration is at a high level, reaching 1.700 mg/g wet wt. in February. In April and May, the concentration of trehalose drops to a low of 0-500 mg/g wet wt., but rises again in June and July to another high of about 1-900 mg/g wet wt. The two high levels of trehalose, in February and July, are significantly different from the levels found in other months (p < 0-05) but do not differ significantly from each other. There is no significant difference in the trehalose concentrations for the months of August through January, and April and May.
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FIG. 1. Amount of trehalose in oysters from April, 1965, to March, 1966. DISCUSSION The results show that the anthrone-positive substance in the disaccharide position on the chromatogram is trehalose. It is non-reducing, its R/is similar to that of authentic trehalose in all solvents used, and it is hydrolyzed quantitatively to glucose by Musca domestica trehalase. Although it is not possible to state definitely the function of trehalose in the oyster from the data obtained in this study, several possibilities can be examined. The biosyntheses of glycogen and trehalose are closely related as shown by descriptions of the process in yeast (Cabib & Leloir, 1958; Candy & Kilby, 1961; and Aligranati & Cabib, 1962):
Trehalose
UDP-glucose + glucose-6-phosphate
trehalose-phosphate
synthctasc
UDP + trchalose-phosphatc trehalose-phosphate + HIO trehalose + P~
trehalosc-6-phosphatasc
626
D a v m G. BADMAN
Glycogen glycogen synthetase UDP-glucose + (glycogen)n - UDP + (glycogen)m+x Murphy & Wyatt (1965) proposed the following scheme for regulation of glycogen and trehalose levels in the silk moth, Hyalophora cecropia. Trehalosephosphate synthetase has greater affinity for UDP-glucose than does glycogen synthetase, allowing preferential synthesis of trehalose when UDP levels are low. When trehalose accumulates, inhibiting trehalose-phosphate synthetase, the level of UDP-glucose increases and permits increased glycogen synthesis. This system provides for rapid production of both trehalose and glycogen when glucose is taken into the hemolymph, followed by readjustment to the initial hemolymph trehalose level. If oyster trehalose is an important storage compound, as it seems to be in insects, its concentration should compare favourably with that of glycogen, which has been considered the main molluscan storage carbohydrate. The amount of trehalose obtained from the oyster is less than that reported for insects, however, and amounts to only a small percentage of reported glycogen values in oysters (Galtsoff, 1964). In Fig. 2 the yearly cycle of glycogen (Marshall, 1966) is compared with that of trehalose from the present study. The glycogen data are from oysters collected at the same time as those in the present paper, and taken also from the dock on Seahorse Key. At the time of spawning, oysters rapidly use up their reserve supply of glycogen, and by the end of the spawning season the amount of glycogen is at a minimum. After a short period, the oysters again begin to accumulate glycogen (Galtsoff, 1964). With the onset of spawning in late March (Churchill, 1920), the glycogen level declines until a minimum is reached in June. While glycogen is diminishing, the trehalose level is very low, but in June and July the trehalose concentration increases to nearly the same as that of glycogen. With the rise of glycogen levels in August, trehalose drops until by October the amount of trehalose is 1/50 that of glycogen. Since the method of analysis involved the extraction of trehalose from the whole body, it is not known whether trehalose is spread throughout the body of the oyster or is restricted to the blood. It is possible that if trehalose is restricted to the blood and separated from the glycogen reserves it could perform a storage function. However, the quantity of trehalose found is so low relative to glycogen, that it is likely that it has a function other than storage. The action of trehalose in aiding glucose absorption through the intestine is another possibility. In the 1ocnst, Schistocercagregaria, the hemolymph trehalose level increases during glucose absorption (Treherne, 1958a). The rate at which glucose is absorbed across the gut wall is directly related to the extent of its conversion to trehalose (Treherne, 1958b). Therefore, trehalose formation in the
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TREHALOSE I N CRASSOSTREA VIRGINICA
oyster may be a means of keeping the blood glucose concentration low, establishing a steep glucose gradient, and allowing glucose to diffuse into the blood regardless of the concentration of sugars in the intestine.
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FIo. 2. Amount of trehalose found in the present work from April, 1965, to March, 1966, compared with glycogen content of oysters from March, 1965, to February, 1966 (Marshall, 1966). • -• trehalose, O - - O glycogen. The present data correspond weU to this idea. The level of trehalose remains at a very low, constant level during time of high glycogen synthesis, suggesting a rapid turnover of trehalose. With the approach of spring, the rate of glycogen synthesis is reduced. At a time of probable high feeding activity in February and March, glucose intake is also high. The trehalose level could be expected to increase. Then as enough trehalose accumulates to inhibit trehalose-phosphate synthetase, trehalose synthesis would drop. In March, with rapid proliferation of sex cells, most of the glycogen store would be depleted, and concomitantly, the slight trehalose reserves.
628
DAwvG. BADMAN
In May and June little glycogen is produced and trehalose accumulates. At the end of the summer, glycogen synthesis increases, and trehalose is again depleted, dropping to a constant low level in August. SUMMARY ; 1. Trehaiose, the major blood sugar of insects, was measured i n Crassostrea virginica Gmelin collected from the G u l f Coast of Florida. 2. Trehalose was obtained from monthly oyster collections by preparative thin layer chromatography, and assayed with anthrone reagent. 3. Amounts of trehalose ranging from 0.188 mg/g wet wt. to 3.300 mg/g wet wt. were obtained. 4. T h e average amount of trehalose in the oyster is constant throughout the year except for an elevation in February and July. 5. T h e amount of trehalose in the oyster was found to be very low in comparison with reported amounts of glycogen, averaging only about 1/50 of the peak glycogen level. 6. A possible role of trehalose in aiding in the uptake of glucose is discussed. Acknowledgements--The author is grateful to Dr. R. M. DeWitt of the Department of Zoology, University of Florida, for his guidance and valuable criticism. REFERENCES ALIGRANATI I. D. & CABIB E. (1962) Uridine diphosphate D-glucose-glycogen glucosyltransferase from yeast. •. biol, Chem. 237, 1007-1013. BL~XN~ J. D. & Y ~ v H. J. (1956) Electrolytic desalting with ion exchange membranes. Nature, Lond. 117, 83. C~am E. & L~Lom L. F. (1958) The biosynthesis of trehalose phosphate..7, b/ol. Chem. 231, 259-275. CANDY D. J. & KILBY B. A. (1961) The biosynthesis of trehalose in the locust fat body. Biochem. jT. 78, 531-536. CXaaOLL N. V., LONGLm"R. W. & RoE J. H. (1956) The determination of glycogen in liver and muscle by use of anthrone reagent. J. biol. Chem. 220, 583-593. CHm~CmLL E. P. JR. (1920) The oyster and the oyster industry of the Atlantic and Gulf coasts~ Bureau of Fisheries Document No. 890. Appendix VIII to the report of the U.S. Commissioner of Fisheries for 1919. CI~C,o J. S. (1965) Origin and significance of trehalose in formation of encysted dormant embryos of Artemis salina. Comp. Biochera. Physiol. 14, 135-143. CLEOO J. S. & EVANSD. R. (1961) Blood trehalose and flight metabolism in the blowfly. Sdence 134 (3471), 54-55. CLEOO J. S. & FILOSa M. F. (1961) Trehalose in the cellular slime mould Dictyosteliura mucoroides. Nature, Lond. 192, 1077-1078. Due~u G. & FLORKrS M. (1959) Sur la tr6halos6rnie des insectes et sa signification. Archs int. Physiol. 67, 306-314. FAXmaAIRND. (1958) Trehalose and glucose in Helminths and other invertebrates. Can..7. Zool. 36, 787-795. G ~ o F F P. S. (1964) The American Oyster. Fishery Bulletin of the Fish and Wildlife Ser~e, Vol. 64. K ~ v G. E. & RmDm~ S. V. (1958) The purification and properties of trehalase. ~t. b/ol. Chem. 230, 691-698.
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MARSHALLH. L. (1966) Personal communication. MURPHY T. A. & WYATTG. R. (1965) The enzymes of glycogen and trehalose synthesis in silkrnoth fat body. y. biol. Chem. 240 (4), 1500-1508. SACKTORB. (1955) Cell structure and the metabolism of insect flight muscle. ~7. biophys. biochem. Cytol. 1, 29-46. SAITO S. (1963) Trehalose in the body fluid of the silkworm, Bombyx mot/ L. ~. Insect. Physiol. 9, 509-519. ST~w.~ J. E. (1963) The site of action of the insect hyperglycemic hormone. Gen. Comp. Endocrinol. 3, 46-52. T I ~ H ] ~ J. E. (1958a) The absorption of glucose from the alimentary canal of the locust Schistocerca gregaria (Forsk)..7. exp. Biol. 35, 279-306. TRm-IeRNE J. E. (1958b) The absorption and metabolism of some sugars in the locust Schistocerca gregaria (Forsk). J. exp. Biol. 35, 611-625. WYATr G. R. & KALF G. F. (1957) The chemistry of insect hemolymph II. Trehalose and other carbohydrates, jT. gen. Physiol. 40, 833-847.
QUANTITATIVE STUDIES ON TREHALOSE IN THE OYSTER, CRASSOSTREA VIRGINICA GMELIN D A V I D G. BADMAN* Department of Zoology, University of Florida, Gainesville, Florida 32601, U.S.A.
(Received 20 May 1967) Abstract--1. Trehalose was identified in the oyster, Crassostrea virginica Gmelin, and found to be present throughout the period of one year in amounts ranging from 0.188 mg/g wet wt. to 3"300 mg/g wet wt. 2. Trehalose levels remained constant except for February and July, when they were significantly elevated. 3. The amount of trehalose relative to glycogen is normally very small except immediately after spawning, when the glycogen level is drastically reduced. INTRODUCTION TIu~H~a~OSE, l(~-v-glucopyranosyl) c~-v-glucopyranoside, long believed to be confined largely to fungi and yeast, has been shown to be a major sugar in insect hemolymph (Wyatt & Kalf, 1957). Falrbairn (1958) identified trehalose in many other invertebrate species. In general, the trehalose of arthropods appears to be a storage compound. It is a glucose source during flight in Musca domestica (Sacktor, 1955) and in Phorraia regina (Clegg & Evans, 1961). The dormant embryo of the brine shrimp, Artemia sally, contains a large trehalose reserve which is converted to glycogen at the end of the dormant period (Clegg, 1956). When the silkworm is subjected to exhaustion, hemolymph trehalose decreases rapidly (Duchateau & Florkin, 1959). Saito (1963) reported the presence of a mechanism to regulate the hemolymph sugar level in the silkworm, Bombyx ~ r i . As the original hemolymph trehalose is used, its concentration is kept constant by the biosynthesis of new trehalose from carbohydrate stored as glycogen in the fat body. In the cockroach, the balance between hemolymph trehalose and fat body glycogen appears to be regulated by hormones (Steele, 1963). It is evident that in insects there is a close relationship between trehalose and glycogen. It has long been known that molluscs contain large amounts of glycogen, and an annual cycle of this carbohydrate has been observed in oysters (Galtsoff, 1964). In view of this, a study of trehalose content of oysters throughout the period of 1 year was undertaken. * This paper is based on a thesis submitted to the Graduate School Of the University of Florida in partial fulfdlment of the requirements for the degree of Master of Science in the Department of Zoology. 621
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MATERIALS AND METHODS Oysters, Crassostrea virginica Gmelin, used in this study were collected from the dock at the University of Florida Marine Laboratory on Seahorse Key, near Cedar Keys, Florida. Collections were made in the middle of each month beginning April, 1965, and ending March, 1966. The shells of oysters from Apalachicola Bay reach 100 mm in height after one summer of growth (Ingle, 1950). Six oysters of this approximate size (75-110 ram) were selected each month for analysis. The oysters were placed in buckets and transported 50 miles to Gainesville where they were frozen as soon as possible, usually within 2-3 hr after collection.
Preparation of extracts The oysters were thawed and their shells measured. The shucked, drained tissue was weighed and then homogenized for 5 rain in 70% (v/v) ethanol in a Waring blender. The homogenate was centrifuged in a Servall centrifuge (Type SS-1 head) at 50 V (ca. 10,300 g) for 10 min. The supernate was evaporated to dryness by warming on a hot plate. The water-soluble portion of the dried supernate was redissolved in distilled water at room temperature, and the volume adjusted so that 1 ml represented 100 mg of the original wet wt of the oyster. The adjusted extracts were desalted in a Torbal-BLT Chromatographic Desalting Apparatus (Model CD-1) using a Permutit C-20 cation resin membrane and A-20 anion resin membrane (Blainey and Yardley, 1956). Desalted extracts were then frozen for later use.
Chromatography and quantitative analysis Thin layer chromatography was used to separate trehalose from other carbohydrates present in the desalted extracts, using prespread Eastman Chromagram Sheets (Type K301R2) and Developing Apparatus. Chromatograms were activated by heating for 30 min at 100°C, and were then spotted with 100/zg authentic D(+) trehalose dihydrate from Mann Research Laboratories, Inc. and with the desalted extracts. To prepare an extract for spotting, 0"5 ml of the desalted extract was evaporated to dryness by warming in a sand bath, and then redissolved in 50/~1 of water. Of this, 20/zl were spotted on each chromatogram. N-butanol: ethanol : water (4 : 1 : 1.9) was the solvent used to separate the components of the extracts. When the solvent front had moved 10 cm, the chromatograms were removed, dried at room temperature and cut into strips corresponding to knowns (authentic trehalose), unknowns (oyster extracts) and blanks (distilled water). In order to visualize the spots, the dried chromatogram strips were dipped in AgNO3 reagent (prepared by diluting 0.1 ml of saturated aqueous AgNO3 to 20 ml with acetone and then dissolving the precipitated AgNO3 by dropwise addition of water). The chromatogram strips were then dipped in 0"5 N NaOH in aqueous ethanol (made by diluting saturated NaOH to 0.5 N with absolute ethanol). Reducing sugars form black spots at room temperature, while trehalose forms
~TgEHALOSBIN CRASSOSTREA VIRGINICA
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light brown spots. This procedure detected 0.5/~g trehalose in a solution containing 50/~g per ml. Trehalose was located on one known strip and one unknown strip to ascertain its position on the remaining three strips representing each oyster. From the unstained strips and from the blank, a rectangular area corresponding to the standard trehalose position was cut out and eluted with 1.0 ml of water. Quantitative elution with gravity was accomplished in 15-20 rain with the strip fed continuously by a wick inserted into a container of distilled water. Eluates of the unknown strips were negative for glucose when analyzed with the glucose oxidase system, Glucostat, obtained from Worthington Biochemical Corporation, showing that there had been adequate separation of trehalose from glucose. The eluates were analyzed for trehalose with the anthrone procedure of Carroll, Longley & Roe (1956).
Criteria of trehalose presence T h e presence of trehalose in the extract was proven by several criteria used by Clegg & Filosa (1961). (1) Strips were sprayed with 0"5% (w/v) sodium metaperiodate and allowed to dry. After drying, the strips were stained with alkaline silver nitrate as before. Treatment with sodium metaperiodate oxidizes the ~,~ bond of trehalose to form glucose, which then gives the characteristic reaction of a reducing sugar with silver nitrate. T h e results, shown in Table 1, indicate that the unknown sugar in the "trehalose" position (i.e. with R/the same as that of authentic trehalose) is non-reducing. TABLE 1----COLORREACTIONWITH ALKALIHESILVERNITRATE Chromatogramed sample
Not treated with Na m-periodate
Treated with Na m-periodate
Authentic glucose
Black spot
Black spot
Authentic trehalose
immediately Brown spot after 30 sec
immediately Black spot immediately
Black spot immediately
Black spot immediately
Brown spot after 30 sec
Black spot immediately
Unknown: "glucose" position "trehalose" position
(2) The unknown eluate reacts with anthrone. (3) T h e spot corresponding to trehalose on the unknown strip was eluated and rechromatogramed in four different solvent systems: (a) n-butanol:ethanol : acetone : water (5 : 4 : 3 : 20), (b) n-propanol : ethyl acetate : water (7 : 1 : 2), (c) ethyl
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acetate : pyridine : water ( 8 : 2 : 1), and (d) n-butanol : aceticacid : water(4 : 1 : 5). In each of these systems, the unknown eluted spot had the same R t as the authentic trehalose, and in each no other spot appeared, showing that the eluate was pure. (4) The unknown eluate was hydrolyzed quantitatively to glucose by trehalas¢. Trehalase was prepared by a procedure modified from Kalf & Rieder (1958). Approximately 1000 three-day-old house flies, Musca domestica, weighing a total of 27.82 g were used. The trehalase obtained was dissolved in 10"1 ml of 0.10 M phosphate buffer, pH 6.8, and stored at 2°C. It was found to be free of maltase activity, and hydrolyzed authentic trehalose to glucose with an average yield of 95 per cent. The assay mixture contained 0.1 ml of 0"10 trehalose or other substrates (distilled water blank, maltose or unknown), 0.35 ml of 0.10 M acetate buffer (pH 5.5), and 0.05 ml of the enzyme. After a 15 rain incubation at 30°C, the reaction was stopped by immersing the tubes in a boiling water bath for 3 rain. The tubes were cooled in an ice bath, the precipitate removed by centrifugation, and a sample removed for determination of glucose. A zero-time control was always run to correct for any glucose in the trehalose or enzyme preparation. Unknown eluates were analyzed for trehalose by this procedure and by the anthrone procedure. In all cases the results of the two methods were identical.
RESULTS
Thin layer chromatography proved to be a satisfactory technique for isolating trehalose from oyster extracts. In a warm room (above 20°C), the plates could be developed in 2-3 hr. Separation of monosaccharides from disaccharides was good. The R/ of glucose averaged 0.35, that of trehalose, 0.22. Recovery of authentic trehalose by elution from the chromatogram by gravity and analysis with anthrone averaged 95 per cent. Trehalose was found in all extracts. The amounts of trehalose found ranged from 0.188 mg/g wet wt. to 3.300 mg/g wet wt. Although oysters having reasonably uniform shell sizes were selected for analysis, the wet wt. of the tissue varied from 1.8 g to 151.1 g. There was no significant correlation between wet weight of tissues and amount of trehalose present (r = 0.0787, p >0"1). Figure 1 gives a summary of the results. From August until January., a t i m e of high glycogen synthesis and storage (Galtsoff, 1964), the trehalose concentration is nearly constant, averaging 0"600 mg trehalose/g wet wt. From January to March, the trehalose concentration is at a high level, reaching 1.700 mg/g wet wt. in February. In April and May, the concentration of trehalose drops to a low of 0-500 mg/g wet wt., but rises again in June and July to another high of about 1-900 mg/g wet wt. The two high levels of trehalose, in February and July, are significantly different from the levels found in other months (p < 0-05) but do not differ significantly from each other. There is no significant difference in the trehalose concentrations for the months of August through January, and April and May.
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FIG. 1. Amount of trehalose in oysters from April, 1965, to March, 1966. DISCUSSION The results show that the anthrone-positive substance in the disaccharide position on the chromatogram is trehalose. It is non-reducing, its R/is similar to that of authentic trehalose in all solvents used, and it is hydrolyzed quantitatively to glucose by Musca domestica trehalase. Although it is not possible to state definitely the function of trehalose in the oyster from the data obtained in this study, several possibilities can be examined. The biosyntheses of glycogen and trehalose are closely related as shown by descriptions of the process in yeast (Cabib & Leloir, 1958; Candy & Kilby, 1961; and Aligranati & Cabib, 1962):
Trehalose
UDP-glucose + glucose-6-phosphate
trehalose-phosphate
synthctasc
UDP + trchalose-phosphatc trehalose-phosphate + HIO trehalose + P~
trehalosc-6-phosphatasc
626
D a v m G. BADMAN
Glycogen glycogen synthetase UDP-glucose + (glycogen)n - UDP + (glycogen)m+x Murphy & Wyatt (1965) proposed the following scheme for regulation of glycogen and trehalose levels in the silk moth, Hyalophora cecropia. Trehalosephosphate synthetase has greater affinity for UDP-glucose than does glycogen synthetase, allowing preferential synthesis of trehalose when UDP levels are low. When trehalose accumulates, inhibiting trehalose-phosphate synthetase, the level of UDP-glucose increases and permits increased glycogen synthesis. This system provides for rapid production of both trehalose and glycogen when glucose is taken into the hemolymph, followed by readjustment to the initial hemolymph trehalose level. If oyster trehalose is an important storage compound, as it seems to be in insects, its concentration should compare favourably with that of glycogen, which has been considered the main molluscan storage carbohydrate. The amount of trehalose obtained from the oyster is less than that reported for insects, however, and amounts to only a small percentage of reported glycogen values in oysters (Galtsoff, 1964). In Fig. 2 the yearly cycle of glycogen (Marshall, 1966) is compared with that of trehalose from the present study. The glycogen data are from oysters collected at the same time as those in the present paper, and taken also from the dock on Seahorse Key. At the time of spawning, oysters rapidly use up their reserve supply of glycogen, and by the end of the spawning season the amount of glycogen is at a minimum. After a short period, the oysters again begin to accumulate glycogen (Galtsoff, 1964). With the onset of spawning in late March (Churchill, 1920), the glycogen level declines until a minimum is reached in June. While glycogen is diminishing, the trehalose level is very low, but in June and July the trehalose concentration increases to nearly the same as that of glycogen. With the rise of glycogen levels in August, trehalose drops until by October the amount of trehalose is 1/50 that of glycogen. Since the method of analysis involved the extraction of trehalose from the whole body, it is not known whether trehalose is spread throughout the body of the oyster or is restricted to the blood. It is possible that if trehalose is restricted to the blood and separated from the glycogen reserves it could perform a storage function. However, the quantity of trehalose found is so low relative to glycogen, that it is likely that it has a function other than storage. The action of trehalose in aiding glucose absorption through the intestine is another possibility. In the 1ocnst, Schistocercagregaria, the hemolymph trehalose level increases during glucose absorption (Treherne, 1958a). The rate at which glucose is absorbed across the gut wall is directly related to the extent of its conversion to trehalose (Treherne, 1958b). Therefore, trehalose formation in the
627
TREHALOSE I N CRASSOSTREA VIRGINICA
oyster may be a means of keeping the blood glucose concentration low, establishing a steep glucose gradient, and allowing glucose to diffuse into the blood regardless of the concentration of sugars in the intestine.
I$.0
4.0
3.0
2.G
1.0
M 1965
A
M
d
d
A Monlh
S
0
N
D
d
F
M
1966
FIo. 2. Amount of trehalose found in the present work from April, 1965, to March, 1966, compared with glycogen content of oysters from March, 1965, to February, 1966 (Marshall, 1966). • -• trehalose, O - - O glycogen. The present data correspond weU to this idea. The level of trehalose remains at a very low, constant level during time of high glycogen synthesis, suggesting a rapid turnover of trehalose. With the approach of spring, the rate of glycogen synthesis is reduced. At a time of probable high feeding activity in February and March, glucose intake is also high. The trehalose level could be expected to increase. Then as enough trehalose accumulates to inhibit trehalose-phosphate synthetase, trehalose synthesis would drop. In March, with rapid proliferation of sex cells, most of the glycogen store would be depleted, and concomitantly, the slight trehalose reserves.
628
DAwvG. BADMAN
In May and June little glycogen is produced and trehalose accumulates. At the end of the summer, glycogen synthesis increases, and trehalose is again depleted, dropping to a constant low level in August. SUMMARY ; 1. Trehaiose, the major blood sugar of insects, was measured i n Crassostrea virginica Gmelin collected from the G u l f Coast of Florida. 2. Trehalose was obtained from monthly oyster collections by preparative thin layer chromatography, and assayed with anthrone reagent. 3. Amounts of trehalose ranging from 0.188 mg/g wet wt. to 3.300 mg/g wet wt. were obtained. 4. T h e average amount of trehalose in the oyster is constant throughout the year except for an elevation in February and July. 5. T h e amount of trehalose in the oyster was found to be very low in comparison with reported amounts of glycogen, averaging only about 1/50 of the peak glycogen level. 6. A possible role of trehalose in aiding in the uptake of glucose is discussed. Acknowledgements--The author is grateful to Dr. R. M. DeWitt of the Department of Zoology, University of Florida, for his guidance and valuable criticism. REFERENCES ALIGRANATI I. D. & CABIB E. (1962) Uridine diphosphate D-glucose-glycogen glucosyltransferase from yeast. •. biol, Chem. 237, 1007-1013. BL~XN~ J. D. & Y ~ v H. J. (1956) Electrolytic desalting with ion exchange membranes. Nature, Lond. 117, 83. C~am E. & L~Lom L. F. (1958) The biosynthesis of trehalose phosphate..7, b/ol. Chem. 231, 259-275. CANDY D. J. & KILBY B. A. (1961) The biosynthesis of trehalose in the locust fat body. Biochem. jT. 78, 531-536. CXaaOLL N. V., LONGLm"R. W. & RoE J. H. (1956) The determination of glycogen in liver and muscle by use of anthrone reagent. J. biol. Chem. 220, 583-593. CHm~CmLL E. P. JR. (1920) The oyster and the oyster industry of the Atlantic and Gulf coasts~ Bureau of Fisheries Document No. 890. Appendix VIII to the report of the U.S. Commissioner of Fisheries for 1919. CI~C,o J. S. (1965) Origin and significance of trehalose in formation of encysted dormant embryos of Artemis salina. Comp. Biochera. Physiol. 14, 135-143. CLEOO J. S. & EVANSD. R. (1961) Blood trehalose and flight metabolism in the blowfly. Sdence 134 (3471), 54-55. CLEOO J. S. & FILOSa M. F. (1961) Trehalose in the cellular slime mould Dictyosteliura mucoroides. Nature, Lond. 192, 1077-1078. Due~u G. & FLORKrS M. (1959) Sur la tr6halos6rnie des insectes et sa signification. Archs int. Physiol. 67, 306-314. FAXmaAIRND. (1958) Trehalose and glucose in Helminths and other invertebrates. Can..7. Zool. 36, 787-795. G ~ o F F P. S. (1964) The American Oyster. Fishery Bulletin of the Fish and Wildlife Ser~e, Vol. 64. K ~ v G. E. & RmDm~ S. V. (1958) The purification and properties of trehalase. ~t. b/ol. Chem. 230, 691-698.
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629
MARSHALLH. L. (1966) Personal communication. MURPHY T. A. & WYATTG. R. (1965) The enzymes of glycogen and trehalose synthesis in silkrnoth fat body. y. biol. Chem. 240 (4), 1500-1508. SACKTORB. (1955) Cell structure and the metabolism of insect flight muscle. ~7. biophys. biochem. Cytol. 1, 29-46. SAITO S. (1963) Trehalose in the body fluid of the silkworm, Bombyx mot/ L. ~. Insect. Physiol. 9, 509-519. ST~w.~ J. E. (1963) The site of action of the insect hyperglycemic hormone. Gen. Comp. Endocrinol. 3, 46-52. T I ~ H ] ~ J. E. (1958a) The absorption of glucose from the alimentary canal of the locust Schistocerca gregaria (Forsk)..7. exp. Biol. 35, 279-306. TRm-IeRNE J. E. (1958b) The absorption and metabolism of some sugars in the locust Schistocerca gregaria (Forsk). J. exp. Biol. 35, 611-625. WYATr G. R. & KALF G. F. (1957) The chemistry of insect hemolymph II. Trehalose and other carbohydrates, jT. gen. Physiol. 40, 833-847.