Vol. 127, No. 1 Printed in U.S.A.
JOURNAL OF BACTERIOLOGY, JUlY 1976, p. 84-90 Copyright © 1976 American Society for Microbiology
Localization of Glycogen Synthetase During Differentiation of Presumptive Cell Types in Dictyostelium discoideum JAMES F. 'HARRIS AND CHARLES L. RUTHERFORD* Department of Biology, Virginia Polytechnic Institute and State University, Blacksburg, Virginia 24061
Received for publication 6 April 1976
Ultramicrochemical techniques were utilized to assay glycogen synthetase (EC 2.4.1.11) activity in cell samples of Dictyostelium discoideum as small as 0.01 jig (dry weight) in reaction volumes of 0.1 ,ul. The activity was assayed by an amplification procedure employing the enzymatic cycling of pyridine nucleotides. These techniques were used to determine the extent of localization of glycogen synthetase in the two cell types during differentiation of D. discoideum. Localization studies in developing spore cells revealed decreasing enzyme activity to the culmination stage. During this phase of development, the enzyme required the presence of soluble glycogen for activity. From culmination to sorocarp stage, enzyme activity increased and was independent of the soluble glycogen. In developing stalk cells, synthetase showed a decreasing gradient of activity. In sorocarps, the cells in the stalk apex showed synthetase activity similar to that of the spores. The cells at the bottom of the stalk had no detectable activity.
The cellular slime mold Dictyostelium discoideum offers several advantages as a model system for studying the spatial, temporal, and biochemical relationships during differentiation. Some of the characteristics of the life cycle include the following. (i) The differentiation process represents a closed system, being induced by deficient nutritional conditions and proceeding in the absence of growth. (ii) Two major cell types are involved. (iii) Differentiation is accompanied by the synthesis of distinct carbohydrate end products synthesized from a common glycogen precursor. (iv) Energy metabolism is separated from end product synthesis in that protein is utilized as the primary energy source and carbohydrate is preserved. The enzymes regulating glycogen levels in D. discoideum have been studied frequently since glycogen degradation supplies glucose units for the end products of differentiation. The activity of the degradative enzyme glycogen phosphorylase (EC 2.4.1.1) is tightly coupled to the differentiation cycle (1, 4). The enzyme is synthesized during development, with a peak of activity occurring at the culmination stage. The peak activity of the degradative enzyme coincides precisely with the initiation of net glycogen degradation. Glycogen synthetase (EC 2.4.1.11), on the other hand, shows no apparent relationship to the time period of development. When the enzyme is assayed in the presence of glycogen, which in most systems is required as a primer, the activity is nearly the
same at all stages of development (3, 15). This lack of correlation between the activity of glycogen synthetase and the stages of differentiation is only superficial, however. Studies on primer dependency of glycogen synthetase over the time period of development showed a sharp loss in requirement for glycogen primer at the culmination stage (16). Thus, the loss in primer requirement coincides precisely with the initiation of glycogen degradation. Ward and Wright (11) investigated a cell wall-associated glycogen synthetase from cell husk extracts prepared at the culmination stage of development. The glycogen product of the enzyme activity was found to be associated with the cell wall material. Wright et al. (17) showed that soluble glycogen synthetase obtained from myxamoebae will adhere to cell wall material prepared from a later stage of development. The enzyme could be eluted from the cell wall material with glycogen. Subsequently, the cell wall material was used as a source of both enzyme and primer to synthesize insoluble cell wall-associated glycogen from uridine 5'-diphosphate glucose (UDPG). In this report we utilize ultramicrochemical techniques to investigate the extent to which glycogen synthetase is localized in two cell types during development of the organism. MATERIALS AND METHODS Chemicals. All reagents were purchased from Sigma Chemical Co., St. Louis, Mo. The enzymes 84
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used in the cycling reaction mixtures were Norite treated before use to remove any nucleotide contamination. The enzymes were suspended in 2% Norite in 100 mM phosphate buffer (pH 7.4) and incubated in an ice bath for 10 min. After the Norite was removed by low-speed centrifugation, ethylenediaminetetraacetate (EDTA) and bovine serum albumin were added to the supernatant to give final concentrations of 5 mM and 0.1%, respectively. Both glucose-6-phosphate dehydrogenase (G6PDH) and glutamic dehydrogenase (GDH) were found to contribute significantly to the cycling blank without prior Norite treatment. Agar, yeast extract, and peptone were obtained from Difco Laboratories, Detroit, Mich. Growth and harvesting conditions. The amoebae of D. discoideum NC-4 were grown with Escherichia coli on nutrient agar surfaces, as previously described by Liddel and Wright (5). The amoebae were spread onto 12.5-cm disks of Whatmal no. 50 filter paper, placed on non-nutrient agar in petri plates. The non-nutrient agar contained 20 g of agar and 1.0 mM EDTA in 1.0 liter of 0.01 M potassium phosphate buffer, pH 6.5. The petri plates were covered and maintained at 23 C. The filter paper was removed from the agar and placed on dry ice at hourly intervals during the life cycle. The filter paper was placed in a freeze-dry apparatus (model 10-800, Virtis Research Equipment, Gardiner, N.Y.) for 36 at -40 C. After lyophilization, the filter papers were cut into portions (2 by 4 cm). Each sample was stored in a lyophilization flask (screw-cap vial no. 10-159-10, Virtis Co., Gardiner, N.Y.) under vacuum at -30 C. Tissue stored in this manner has shown no loss of activity over a 2year period. On the day an assay was made, the vacuum flask was allowed to reach room temperature, the vacuum was released, and the tissue was removed. Microtechniques. When specific cell types were assayed, the freeze-dried individual organism was dissected. Cutting was done free-hand under a dissecting microscope. A microscapel was constructed from a small section of a razor blade edge anchored to a dowel rod with a short piece of copper wire. The two cell types were easily separated at all stages of development. Specific enzyme activity and substrate or product level were expressed per unit of dry weight as measured by a quartz-fiber balance (6). The balances were calibrated as described by Lowry and Passonneau (7). Dry weights ranging from 0.01 ,ug (sections) to 10 ,ug (whole organisms) were assayed. When sections of an individual organism were assayed, the initial reactions were carried out under a mineral oil surface (7). The reaction vessel was a 4mm well drilled into Teflon block (20 by 120 by 5 mm). Glycogen synthetase assay. When sections of an organism were assayed, the following protocol was used. To the bottom of a well was added a 0.1-,,l reaction mixture containing 50 mM glycylglycine buffer (pH 8.0), 10 mM UDPG, 5 mM EDTA, 18 mM glycogen, and 0.016% (vol/vol) Triton X-100. The section of organisms was added to the reaction mix-
85
ture at the bottom of the well. The well was filled with a hexadecane-mineral oil (3:7, vol/vol) mixture and incubated in a 37 C water bath for 90 min. The reaction was stopped by heating for 10 min at 90 C. The uridine-5'-diphosphate (UDP) produced in the glycogen synthetase reaction was assayed by adding 2.5 p.l of a UDP reaction mixture to the reagents in the well. The UDP reaction mixture contained 50 mM glycylglycine buffer (pH 8.0), 0.1 mM phosphoenol pyruvate, 65 mM KCI, 15 mM MgCl., 0.04 mM reduced nicotinamide adenine dinucleotide (NADH), 15 U of pyruvate kinase per ml, and 5 U of lactate dehydrogenase per ml. The well was incubated at 37 C for 40 min. The reactions were stopped, and excess NADH was destroyed by the addition of 2.5 ,ul of 0.35 N HCI. The low level of NADI produced was amplified by enzymatic cycling. The entire reaction volume was removed from the well and was added to a 3-ml test tube containing 50 jl of cycling reaction mixture. The cycling reaction mixture contained 100 mM phosphate buffer (pH 7.4), 0.02% albumin, 8 mM sodium acetate, 3.5 mM ammonium acetate, 0.2 mM adenosine 5'-diphosphate, 2 mM a-ketoglutarate, 2.5 mM mercaptoethanol, 2 mM glyceraldehyde-3-phosphate (GAP), 27 U of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) per ml, and 31 U of GDH per ml. The test tubes were capped and incubated for 60 min at 37 C. Glutamate and 3-P-glycerate accumulated as a result of the cycling reactions. After the cycling reactions were completed, 6.5 pA of 3% hydrogen peroxide was added to destroy excess a-ketogluterate (8). The enzymes were denatured by boiling for 3 min. The glutamate produced in the cycling reaction was assayed by adding 1.0 ml of the glutamate reaction mixture, containing 50 mM hydrazine-HCl (pH 9.3), 0.02% albumin, 0.3 mM adenosine-5'-diphosphate, 0.4 mM NADI, and 5 U of GDH per ml, to the reagents in the test tube. Incubation was for 30 min at room temperature. The fluorescence of the NADH produced was read on a fluorometer (model A-4, Farrand Optical Co., New York, N.Y.). The fluorometer was equipped with a Corning no. 5840 filter for the incident light and filter no. 3387 plus no. 4308 for the emitted light. UDP standards, in triplicate, were carried through all steps of the assay. The concentration of stock UDP solutions was determined in the UDP reaction mixture. At least 90% recovery of the standards was obtained. A complete range of standards was included each time the assay was done. Enzyme specific activities based on UDP standards are reported as millimoles per hour per kilogram of
dry weight. Glycogen was assayed by the method of Passonneau et al. (9). A sample of the glycogen synthetase reaction mixture was added to an equal volume of a glycogen reaction mixture containing 100 mM imidazole buffer (pH 7.0), 2 mM magnesium acetate, 3.5 mM EDTA, 0.25 mM NADP+, 0.34 mM adenosine5'-monophosphate, 10 mM K.,HPO4, 0.07% albumin, 1.8 mM dithiothreitol 1.7 ,uM glucose-1,6-diphosphate, 10 U of phosphoglucomutase per ml, 4 U of G6PDH per ml, and 7.7 U of phosphorylase a per ml. Incubation was for 60 min at 37 C. The reactions
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stopped and excess NADP+ was destroyed by adding 100 ,l of 6 N NaOH and incubating for 30 min at 60 C. A 100-IAI portion was added to 500 Al of 6 N NaOH containing 0.03% H2O., in a 3-ml test tube. The mixture was incubated for 10 min at 60 C. The final volume was attained by adding 500 ,lp of distilled water, and the fluorescence was measured. were
RESULTS
Glycogen synthetase assay. Initially, satisfactory conditions for each reaction in the assay were accomplished by using 10-- M quantities of standards. The proportions of reagents were then scaled down to 10-" M quantities and eventually to 10-'2 M levels where fluorometric techniques were used to measure reaction rates or product levels. The glycogen synthetase assay system was dependent upon the complete reaction mixture as described in Materials and Methods (Table 1). The low activity found when soluble glycogen was omitted from the reaction mixture was probably due to the presence of endogenous glycogen. This cellular glycogen may be of sufficient level to act as primer for the reaction catalyzed by the enzyme. The reaction was linear for at least 90 min at 37 C. UDP and glycogen were both used as standards for the glycogen synthetase assay. The reaction for UDP and glycogen was linear to at least 80 pmol. The product levels produced by the glycogen synthetase in sections of tissue fell within this range. Both UDP and glycogen were produced in these extracts, in a near 1:1 stoichiometry. Over this range of concentrations, 500-fold amplification by the cycling reagent was needed. The addition to the glycogen reaction mixture of NaOH to final concentrations ranging from 0.08 to 0.10 N resulted in maximum destruction of residual NADP+. The TABLE 1. Assay conditions ofglycogen synthetase activity Components present in glycogen synthetase reaction mixture
NADH oxidized (10-7 M) 30 mine"
Complete"
6.2 0 1.5 0 0
60 mim
12.3
Without cell extract 0 Without glycogen 2.3 Without UDPG 0.8 Without glycogen and UDPG 0 a Incubation time at 37 C. b Complete reaction mixture contained 50 mM glycylglycine buffer (pH 8.0), 10 mM UDPG, 5 mM EDTA, and 18 mM glycogen (standardized as glucose units). Late-aggregation cell extract (75 ,ug, wet weight) was added to 50 ,lp of the reaction mixture. Assayed in 1 ml of UDP reaction mixture as described in the text.
remaining NADPH was amplified by adding 6 N NaOH containing 0.03% H.,O., to a final NaOH concentration of 1.0 N, or by enzymatic cycling. Glycogen synthetase activity was measured in specific cell types by enzymatic cycling involving the coupled reduction-oxidation of NAD+. The level of NAD+ produced by 0.1 ,ug (dry weight) of cells ranged from 10-12 to 10-'5 mol. Both cycling enzymes, GAPDH and GDH, contained NAD+ as a contaminant, which was removed by treatment with Norite. The maximum velocities, in micromoles per minute per milligram of protein, for GDH and GAPDH before and after Norite treatment were 5.83 and 13.1 and 5.00 and 16.8, respectively. Glutamate accumulation in the cycling reaction was determined by the levels of GDH and GAPDH and by the amount of NAD+ added to the cycling reaction mixture. An optimum rate of cycling was achieved with a GAPDH/GDH unit ratio of approximately 70:30. The steady-state level of NADH occurring during cycling was observed directly on the fluorometer in a preliminary test of the cycling reaction mixture (7). The ratio of the reduced to oxidized nucleotide was found to increase with the ratio of GAPDH to GDH. Maximum cycling coincided with a nucleotide ratio of 40:60, reduced to oxidized form. Figure 1 shows that the cycling reaction was linear with increasing amounts of NAD+. UDP levels produced during the glycogen synthetase reaction fell within this concentration range. Localization of total enzyme activity at the midculmination (20-h) stage of development. Figure 2 shows the localization of total glycogen synthetase specific activity in prespore cells and along the vertical axis of the developing stalk from apex to base. This is the first stage in the developmental cycle in which stalk and spore cells are sharply separated and in which cell specific enzyme activity could be detected. For the individual depicted in Fig. 2, the stalks were dissected into five equal sections. The prespore cells and the apical stalk cells showed similar specific activities of 105 + 24 and 115 mmol/h per kg, respectively. There was a decreasing gradient of specific activity from apex to the bottom section of the stalk, which had a specific activity of approximately 30 mmol/h per kg. Localization of total enzyme activity at the late-culmination (23-h) stage of development. The distribution of total glycogen synthetase specific activity between prespore cells and stalk cells at 23 h is shown in Fig. 3. At the late culmination stage of development, the spore mass had lifted off the substratum. The
LOCALIZATION OF GLYCOGEN SYNTHETASE.
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Glycogen synthetase dependence on soluble glycogen primer during differentiation of spores. Since changes in primer dependence of glycogen synthetase have been shown during the differentiation of D. discoideum (12, 16), assays were made to determine whether this also occurred in spore cells free from stalk materials. Soluble glycogen primer dependence of enzyme activity was found in prespore cells of preculmination, but not in the mature, spore cells (Table 2). This primer dependency was demonstrated by an eightfold increase in specific activity when prespore tissue was incubated in the presence of soluble glycogen. There was little difference, however, in the specific activities of sorocarp spores incubated with or without soluble glycogen. The preculmination prespore cells showed an increase in specific activity from 19 to 147 mmol/h per kg when soluble glycogen was added to the incubation mixture. The spore cells from sorocarp showed no change with specific activity remaining at about 200 mmol/h per kg with or without soluble glycogen in the incubation mixture.
10 8 6 NAD+ LEVEL (pmoles) 2
4
FIG. 1. Linearity of cycling with increasing amount of NAD+. NAD+ was added to 50 ,d of cycling mixture.
stalk was nearly completed and visibly protruding from beneath the spores. When the dry individual was dissected at this stage, the spore mass neatly separated from the stalk, leaving the intact stalk with the apical cells attached. The prespore cells and the apical stalk cells showed similar mean specific activities of 162 and 150 mmol/h per kg, respectively. As we found at midculmination, the stalk showed a decreasing gradient of activity with the bottom section having the lowest activity. Localization of total glycogen synthetase activity at sorocarp (24 h). Figure 4 shows the localization of total glycogen synthetase specific activity in the spore cells and along the vertical axis of the completed stalk from apex to base. As was the case with the mid- and late-culmination stages, the spore cells and apex of the stalk had similar specific activities of approximately 200 mmol/h per kg. There was a decreasing gradient of activity from the apex to the first section of stalk located immediately below the sorus. At this point in the stalk, the specific activity was approximately 30 mmol/h per kg. Beyond this section of stalk and continuing to the base of the stalk, there was little or no measurable glycogen synthetase activity remaining.
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FIG. 2. Total glycogen synthetase activity in prespore and stalk cells at the midculmination (20-h) stage of development. Measured as UDP production with enzymatic cycling for amplification. Prespore point represents mean + standard deviation of 11 replications. Stalk points represent one determination from indicated location in the stalk. These data are representative of two assays. Specific activity is expressed as millimoles per hour per kilogram ofdry
weight.
88
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HARRIS AND RUTHERFORD
J. BACTERIOL.
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FIG. 3. Total glycogen synthetase activity in prespore and stalk cells at the late culmination (23-h) stage ofdevelopment. Prespore point represents mean + standard deviation of five replications. Stalk points represent one determination from indicated location in the stalk. These data are representative of three assays. The variation irn values taken from similar areas ofprespore or prestalk cells from different individuals was always less than 20% for all stages of development. Specific activity is expressed as millimoles per hour per kilogram of dry weight.
In addition, further tests were made to determine whether the prespore enzyme could utilize the primer from sorocarp spore cells. In an experiment in which primer-independent activity was measured in prespore cells and sorocarp spore cells separately and mixed, no additive effect was found when the two cell types were mixed (Table 2). This result showed that under the experimental conditions the enzyme from the prespore cells could not utilize the primer employed by the enzyme in the sorocarp spores. Furthermore, prespore extract did not contain an inhibitor that would reduce the activity of the enzyme from sorocarp extract.
DISCUSSION Cell-specific events occurring during spore differentiation. Some of events known to be associated with the differentiation of prespore cells include: (i) the accumulation of inorganic phosphate, (ii) the degradation of soluble glycogen, (iii) the conversion of glycogen synthetase from a soluble to a particulate form, and (iv)
0 L-r1i
I
SPORE
STALK
FIG. 4. Total glycogen synthetase activity in spore and stalk cells at the sorocarp (24-h) stage of development. Spore point represents mean ± standard deviation offive replications. Stalk points represent one determination from indicated location in the stalk. These data are representative of two assays. Specific activity is expressed as millimoles per hour per kilogram of dry weight. TABLE 2. Dependence ofglycogen synthetase activity on soluble glycogen primer in prespore and spore cells Sp acP' (mmol/h per kg, dry wt) Stage of development
-Glycogen +Glycogenb Preculmination (18 19 + 4 (20Y 147 ± 5 (13) h') Sorocarp (24 h) 198 ± 16 (13) 221 ± 30 (6) Preculmination plus 191 ± 17 (7) sorocarpY' a Measured as UDP produced as described in the text. Enzymatic cycling was used for amplification. b Glycogen added to a final concentration of 18 mM (as glucose units). ¢ Hours after removing bacterial food source. d Mean + standard deviation. Number of replications in parentheses. " Preculmination prespore cells (0.13 + 0.01 ,ug, dry weight) added to sorocarp spore cells (0.15 + 0.03 gg, dry weight).
the accumulation of trehalose. In the development of spore cells, inorganic phosphate (Pi) accumulates to an in vivo concentration of 30 mM at the sorocarp stage of development (C.
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LOCALIZATION OF GLYCOGEN SYNTHETASE
Rutherford, J. Embryol. Exp. Morphol., in press). Pi is known to exert multiple effects on both glycogen phosphorylase and soluble glycogen synthetase activities. The glycogen phosphorylase activity is stimulated whereas the activity of the soluble glycogen synthetase is inhibited by Pi. The in vivo levels of Pi localized in the prespore cells at culmination are below Km values for glycogen phosphorylase. Thus, the accumulation during spore maturation would result in increased activitv of this enzyme. These changes in phosphorylase and synthetase activities would lead to an increased capacity to degrade glycogen and to a decreased capacity to synthesize glycogen. A corresponding loss of glycogen is actually observed in these prespore cells; as during sorocarp construction, the glycogen level in the developing spore cells decreased from 30 to 3 mM (Rutherford, in press). The conversion of spore cell glycogen synthetase from a soluble to an insoluble form of the enzyme results in a situation favorable for the synthesis of insoluble glycogen and a decrease in the synthesis of soluble glycogen. An important result of soluble glycogen degradation might be the release of glycogen synthetase from a glycogen-enzyme complex. It has been suggested that some of the soluble enzyme becomes entrapped in the cell wall matrix as it is constructed (17). Therefore, some of the released glycogen synthetase may become insoluble. This change from a soluble to an insoluble form results in a loss of the Pi inhibition. An in vivo Pi concentration of 20 mM is known to inhibit soluble glycogen synthetase by 73% (2), whereas insoluble glycogen synthetase is inhibited by only 28% (13). Therefore, it is likely that the association of a fraction of the soluble glycogen synthetase with the spore cell wall protects it from the inhibitory effects of Pi. As glycogen synthetase becomes insoluble, the dependence of the enzyme on G-6-P levels is overcome. Rosness et al. (10) have demonstrated that the soluble form of glycogen synthetase changes from an I form early in development to a D form at culmination. We have shown that in developing spore cells some insoluble glycogen synthetase is in the I form. G-6-P levels decrease in spore cells, presumably as a result of utilization by the cellulose and trehalose synthetic pathways. Since the soluble glycogen synthetase in spore cells is in the D form, the insoluble I form of the enzyme could successfully compete for UDPG even in the presence of low G-6-P levels. Considering the spore-specific characteristics of glycogen synthetase, glycogen phosphoryl-
89
ase, Pi, and glycogen, we propose a preliminary model of the events associated with glycogen degradation during prespore cell differentiation. As Wright (13) has pointed out, a kinetic situation favoring glycogen breakdown is created in spore cells as a result of both an increase in glycogen phosphorylase activity and the accumulation of Pi. The accumulation of Pi would further decrease glycogen synthesis in prespore cells by inhibition of glycogen synthetase. Upon degradation of glycogen, the synthetic enzyme may become associated with the developing spore wall. The spore wall, or a small portion of the soluble glycogen pool now bound to the spore wall, could act as a primer for the synthesis of "insoluble" glycogen. Both insoluble glycogen synthetase and insoluble glycogen may remain entrapped in the cell wall until germination. Upon germination, cellulases may degrade the cellulose wall, thus releasing the enzyme and glycogen from the cell wall matrix. Thus, the newly emerged myxamoeba would contain both soluble glycogen synthetase and the soluble glycogen primer required for further production of soluble glycogen later in development. Glycogen degradation during prestalk cell migration. Glycogen synthetase specific activity decreases from the apex to base of the stalk. Those cells at the base of the stalk were the first prestalk cells to enter the position of stalk construction. Therefore, the decreasing gradient of activity may be a reflection of the degree of maturation of stalk cells. We cannot determine from the present study the reason for the decrease in activity in mature stalk cells. Since protein degradation is thought to provide an energy source for differentiation in D. discoideum (14), the loss of activity could be the result of degradation of the enzyme as well as other proteins in stalk cells. Alternatively, the loss ofactivity could be due to a conversion from an active to a less active form of the enzyme. The enzyme activity obtained from microgram quantities of lyophilized tissue was compared to levels in tissue homogenates and found to be similar when expressed on a dry weight basis. The glycogen synthetase activity from extracts of freeze-dried tissue (both pseudoplasmodium and sorocarp) was also compared with that from extracts prepared with a French pressure cell or by sonic treatment. Results from all methods of extraction were not significantly different. Thus, the preparation of extracts by freezing followed by lyophilization was sufficient to release enzymes and substrates to the assay conditions. Furthermore trehalase, acid phosphate, and Pi levels increase during stalk
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HARRIS AND RUTHERFORD
cell maturation (Rutherford, in press). Since these enzymes can be assayed in the lyophilized tissue, the freeze-dry treatment apparently disrupts the stalk cells. Thus, the lack of glycogen synthetase activity in stalk cells is not due to an inaccessibility of the enzyme to the reaction mixture. Regardless of the mechanism for the loss of glycogen synthetase activity in stalk cells, the result is a cell-specific kinetic situation favorable for glycogen degradation. Interestingly, glycogen phosphorylase is active in stalk cells (Rutherford, in press). Likewise, Pi shows an increasing gradient from the apex of the stalk to the base (Rutherford, in press). As mentioned previously, accumulation of Pi would increase glycogen phosphorylase activity but inhibit glycogen synthetase. The levels of glycogen in stalk cells show a striking correspondence to the observed synthetic and degradative activities. Prestalk cells, which have not yet migrated into the area of stalk construction, showed no loss of glycogen (Rutherford, in press). Subsequently, as prestalk cells migrated beneath the apex of the stalk, glycogen degradation occurred rapidly with a decreasing gradient toward the base of the stalk. Thus, the specific cells showing a loss of the glycogen synthetic enzyme exhibit an identical decrease in glycogen levels. Although the regulation of glycogen degradation and resultant synthesis of end products are likely to be under multiple control mechanisms, the observed retention of glycogen synthetase in spore cells and the loss of activity in stalk cells must be included in future models for differentiation of the two cell types. ACKNOWLEDGMENTS This research was supported by the Brown-Hazen Program of The Research Corporation. LITERATURE CITED 1. Firtel, R. A., and J. Bonner. 1972. Developmental control of a-1,4-glucan phosphorylase in the cellular slime mold Dictyostelium discoideum. Dev. Biol. 29:85-103.
J. BACTEIUOL. 2. Gezelius, K., and B. E. Wright. 1965. Alkaline phosphatase in Dictyostelium discoideum. J. Gen. Microbiol. 70:309-327. 3. Hames, B. D., G. Weeks, and J. M. Ashworth. 1972. Glycogen synthetase and the control of glycogen synthesis in the cellular slime mold Dictyostelium discoideum during cell differentiation. Biochem. J. 126:627-633. 4. Jones, T. H. D., and B. E. Wright. 1970. Partial purification and characterization of glycogen phosphorylase from Dictyostelium discoideum. J. Bacteriol. 104:754-761. 5. Liddel, G. U., and B. E. Wright. 1961. The effect of glucose on respiration of the differentiating slime mold. Dev. Biol. 3:265-276. 6. Lowry, 0. H. 1941. A quartz fiber balance. J. Biol. Chem. 140:183-189. 7. Lowry, 0. H., and J. V. Passonneau. 1972. A flexible system of enzymatic analysis. Academic Press Inc., New York. 8. Matschinsky, F. M., C. L. Rutherford, and L. Guerra (ed.). 1968. Proceedings of the 3rd International Congress of Histochemistry and Cytochemistry. Springer-Verlag, New York. 9. Passonneau, J. V., P. D. Gatfield, D. W. Schultz, and 0. H. Lowry. 1967. An enzymic method for measurement of glycogen. Anal. Biochem. 19:315-326. 10. Rosness, P. A., G. Gustafson, and B. E. Wright. 1971. Effects of adenosine 3',5'-monophosphate and adenosine 5'-monophosphate on glycogen degradation and synthesis in Dictyostelium discoideum. J. Bacteriol. 108:1329-1337. 11. Ward, C., and B. E. Wright. 1965. Cell wall synthesis inDictyostelium discoideum. I. In vitro systhesis from uridine diphosphoglucose. Biochemistry 4:2021-2027. 12. Wright, B. E. 1966. Multiple causes and controls in differentiation. Science 153:830-837. 13. Wright, B. E. 1973. Critical variables in differentiation. Prentice-Hall, Inc., Englewood Cliffs, N.J. 14. Wright, B. E., and M. L. Anderson. 1960. Protein and amino acid turnover during differentiation in the slime mold. I. Utilization of endogenous amino acids and proteins. Biochim. Biophys. Acta 43:62-66. 15. Wright, B. E., and D. Dahlberg. 1967. Cell wall synthesis in Dictyostelium discoideum. II. Synthesis of soluble glycogen by a cytoplasmic enzyme. Biochemistry 6:2074-2079. 16. Wright, B. E., D. Dahlberg, and C. Ward. 1968. Cell wall synthesis in Dictyostelium discoideum: a model system for the synthesis of alkali-insoluble cell wall glycogen during differentiation. Arch. Biochem. Biophys. 124:380-385. 17. Wright, B. E., C. Ward, and D. Dahlberg. 1966. Cell wall polysaccharide synthesis in vitro catalyzed by an enzyme from slime mold myxamoebae lacking a cell wall. Biochem. Biophys. Res. Commun. 22:352-356.
JOURNAL OF BACTERIOLOGY, JUlY 1976, p. 84-90 Copyright © 1976 American Society for Microbiology
Localization of Glycogen Synthetase During Differentiation of Presumptive Cell Types in Dictyostelium discoideum JAMES F. 'HARRIS AND CHARLES L. RUTHERFORD* Department of Biology, Virginia Polytechnic Institute and State University, Blacksburg, Virginia 24061
Received for publication 6 April 1976
Ultramicrochemical techniques were utilized to assay glycogen synthetase (EC 2.4.1.11) activity in cell samples of Dictyostelium discoideum as small as 0.01 jig (dry weight) in reaction volumes of 0.1 ,ul. The activity was assayed by an amplification procedure employing the enzymatic cycling of pyridine nucleotides. These techniques were used to determine the extent of localization of glycogen synthetase in the two cell types during differentiation of D. discoideum. Localization studies in developing spore cells revealed decreasing enzyme activity to the culmination stage. During this phase of development, the enzyme required the presence of soluble glycogen for activity. From culmination to sorocarp stage, enzyme activity increased and was independent of the soluble glycogen. In developing stalk cells, synthetase showed a decreasing gradient of activity. In sorocarps, the cells in the stalk apex showed synthetase activity similar to that of the spores. The cells at the bottom of the stalk had no detectable activity.
The cellular slime mold Dictyostelium discoideum offers several advantages as a model system for studying the spatial, temporal, and biochemical relationships during differentiation. Some of the characteristics of the life cycle include the following. (i) The differentiation process represents a closed system, being induced by deficient nutritional conditions and proceeding in the absence of growth. (ii) Two major cell types are involved. (iii) Differentiation is accompanied by the synthesis of distinct carbohydrate end products synthesized from a common glycogen precursor. (iv) Energy metabolism is separated from end product synthesis in that protein is utilized as the primary energy source and carbohydrate is preserved. The enzymes regulating glycogen levels in D. discoideum have been studied frequently since glycogen degradation supplies glucose units for the end products of differentiation. The activity of the degradative enzyme glycogen phosphorylase (EC 2.4.1.1) is tightly coupled to the differentiation cycle (1, 4). The enzyme is synthesized during development, with a peak of activity occurring at the culmination stage. The peak activity of the degradative enzyme coincides precisely with the initiation of net glycogen degradation. Glycogen synthetase (EC 2.4.1.11), on the other hand, shows no apparent relationship to the time period of development. When the enzyme is assayed in the presence of glycogen, which in most systems is required as a primer, the activity is nearly the
same at all stages of development (3, 15). This lack of correlation between the activity of glycogen synthetase and the stages of differentiation is only superficial, however. Studies on primer dependency of glycogen synthetase over the time period of development showed a sharp loss in requirement for glycogen primer at the culmination stage (16). Thus, the loss in primer requirement coincides precisely with the initiation of glycogen degradation. Ward and Wright (11) investigated a cell wall-associated glycogen synthetase from cell husk extracts prepared at the culmination stage of development. The glycogen product of the enzyme activity was found to be associated with the cell wall material. Wright et al. (17) showed that soluble glycogen synthetase obtained from myxamoebae will adhere to cell wall material prepared from a later stage of development. The enzyme could be eluted from the cell wall material with glycogen. Subsequently, the cell wall material was used as a source of both enzyme and primer to synthesize insoluble cell wall-associated glycogen from uridine 5'-diphosphate glucose (UDPG). In this report we utilize ultramicrochemical techniques to investigate the extent to which glycogen synthetase is localized in two cell types during development of the organism. MATERIALS AND METHODS Chemicals. All reagents were purchased from Sigma Chemical Co., St. Louis, Mo. The enzymes 84
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used in the cycling reaction mixtures were Norite treated before use to remove any nucleotide contamination. The enzymes were suspended in 2% Norite in 100 mM phosphate buffer (pH 7.4) and incubated in an ice bath for 10 min. After the Norite was removed by low-speed centrifugation, ethylenediaminetetraacetate (EDTA) and bovine serum albumin were added to the supernatant to give final concentrations of 5 mM and 0.1%, respectively. Both glucose-6-phosphate dehydrogenase (G6PDH) and glutamic dehydrogenase (GDH) were found to contribute significantly to the cycling blank without prior Norite treatment. Agar, yeast extract, and peptone were obtained from Difco Laboratories, Detroit, Mich. Growth and harvesting conditions. The amoebae of D. discoideum NC-4 were grown with Escherichia coli on nutrient agar surfaces, as previously described by Liddel and Wright (5). The amoebae were spread onto 12.5-cm disks of Whatmal no. 50 filter paper, placed on non-nutrient agar in petri plates. The non-nutrient agar contained 20 g of agar and 1.0 mM EDTA in 1.0 liter of 0.01 M potassium phosphate buffer, pH 6.5. The petri plates were covered and maintained at 23 C. The filter paper was removed from the agar and placed on dry ice at hourly intervals during the life cycle. The filter paper was placed in a freeze-dry apparatus (model 10-800, Virtis Research Equipment, Gardiner, N.Y.) for 36 at -40 C. After lyophilization, the filter papers were cut into portions (2 by 4 cm). Each sample was stored in a lyophilization flask (screw-cap vial no. 10-159-10, Virtis Co., Gardiner, N.Y.) under vacuum at -30 C. Tissue stored in this manner has shown no loss of activity over a 2year period. On the day an assay was made, the vacuum flask was allowed to reach room temperature, the vacuum was released, and the tissue was removed. Microtechniques. When specific cell types were assayed, the freeze-dried individual organism was dissected. Cutting was done free-hand under a dissecting microscope. A microscapel was constructed from a small section of a razor blade edge anchored to a dowel rod with a short piece of copper wire. The two cell types were easily separated at all stages of development. Specific enzyme activity and substrate or product level were expressed per unit of dry weight as measured by a quartz-fiber balance (6). The balances were calibrated as described by Lowry and Passonneau (7). Dry weights ranging from 0.01 ,ug (sections) to 10 ,ug (whole organisms) were assayed. When sections of an individual organism were assayed, the initial reactions were carried out under a mineral oil surface (7). The reaction vessel was a 4mm well drilled into Teflon block (20 by 120 by 5 mm). Glycogen synthetase assay. When sections of an organism were assayed, the following protocol was used. To the bottom of a well was added a 0.1-,,l reaction mixture containing 50 mM glycylglycine buffer (pH 8.0), 10 mM UDPG, 5 mM EDTA, 18 mM glycogen, and 0.016% (vol/vol) Triton X-100. The section of organisms was added to the reaction mix-
85
ture at the bottom of the well. The well was filled with a hexadecane-mineral oil (3:7, vol/vol) mixture and incubated in a 37 C water bath for 90 min. The reaction was stopped by heating for 10 min at 90 C. The uridine-5'-diphosphate (UDP) produced in the glycogen synthetase reaction was assayed by adding 2.5 p.l of a UDP reaction mixture to the reagents in the well. The UDP reaction mixture contained 50 mM glycylglycine buffer (pH 8.0), 0.1 mM phosphoenol pyruvate, 65 mM KCI, 15 mM MgCl., 0.04 mM reduced nicotinamide adenine dinucleotide (NADH), 15 U of pyruvate kinase per ml, and 5 U of lactate dehydrogenase per ml. The well was incubated at 37 C for 40 min. The reactions were stopped, and excess NADH was destroyed by the addition of 2.5 ,ul of 0.35 N HCI. The low level of NADI produced was amplified by enzymatic cycling. The entire reaction volume was removed from the well and was added to a 3-ml test tube containing 50 jl of cycling reaction mixture. The cycling reaction mixture contained 100 mM phosphate buffer (pH 7.4), 0.02% albumin, 8 mM sodium acetate, 3.5 mM ammonium acetate, 0.2 mM adenosine 5'-diphosphate, 2 mM a-ketoglutarate, 2.5 mM mercaptoethanol, 2 mM glyceraldehyde-3-phosphate (GAP), 27 U of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) per ml, and 31 U of GDH per ml. The test tubes were capped and incubated for 60 min at 37 C. Glutamate and 3-P-glycerate accumulated as a result of the cycling reactions. After the cycling reactions were completed, 6.5 pA of 3% hydrogen peroxide was added to destroy excess a-ketogluterate (8). The enzymes were denatured by boiling for 3 min. The glutamate produced in the cycling reaction was assayed by adding 1.0 ml of the glutamate reaction mixture, containing 50 mM hydrazine-HCl (pH 9.3), 0.02% albumin, 0.3 mM adenosine-5'-diphosphate, 0.4 mM NADI, and 5 U of GDH per ml, to the reagents in the test tube. Incubation was for 30 min at room temperature. The fluorescence of the NADH produced was read on a fluorometer (model A-4, Farrand Optical Co., New York, N.Y.). The fluorometer was equipped with a Corning no. 5840 filter for the incident light and filter no. 3387 plus no. 4308 for the emitted light. UDP standards, in triplicate, were carried through all steps of the assay. The concentration of stock UDP solutions was determined in the UDP reaction mixture. At least 90% recovery of the standards was obtained. A complete range of standards was included each time the assay was done. Enzyme specific activities based on UDP standards are reported as millimoles per hour per kilogram of
dry weight. Glycogen was assayed by the method of Passonneau et al. (9). A sample of the glycogen synthetase reaction mixture was added to an equal volume of a glycogen reaction mixture containing 100 mM imidazole buffer (pH 7.0), 2 mM magnesium acetate, 3.5 mM EDTA, 0.25 mM NADP+, 0.34 mM adenosine5'-monophosphate, 10 mM K.,HPO4, 0.07% albumin, 1.8 mM dithiothreitol 1.7 ,uM glucose-1,6-diphosphate, 10 U of phosphoglucomutase per ml, 4 U of G6PDH per ml, and 7.7 U of phosphorylase a per ml. Incubation was for 60 min at 37 C. The reactions
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HARRIS AND RUTHERFORD
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stopped and excess NADP+ was destroyed by adding 100 ,l of 6 N NaOH and incubating for 30 min at 60 C. A 100-IAI portion was added to 500 Al of 6 N NaOH containing 0.03% H2O., in a 3-ml test tube. The mixture was incubated for 10 min at 60 C. The final volume was attained by adding 500 ,lp of distilled water, and the fluorescence was measured. were
RESULTS
Glycogen synthetase assay. Initially, satisfactory conditions for each reaction in the assay were accomplished by using 10-- M quantities of standards. The proportions of reagents were then scaled down to 10-" M quantities and eventually to 10-'2 M levels where fluorometric techniques were used to measure reaction rates or product levels. The glycogen synthetase assay system was dependent upon the complete reaction mixture as described in Materials and Methods (Table 1). The low activity found when soluble glycogen was omitted from the reaction mixture was probably due to the presence of endogenous glycogen. This cellular glycogen may be of sufficient level to act as primer for the reaction catalyzed by the enzyme. The reaction was linear for at least 90 min at 37 C. UDP and glycogen were both used as standards for the glycogen synthetase assay. The reaction for UDP and glycogen was linear to at least 80 pmol. The product levels produced by the glycogen synthetase in sections of tissue fell within this range. Both UDP and glycogen were produced in these extracts, in a near 1:1 stoichiometry. Over this range of concentrations, 500-fold amplification by the cycling reagent was needed. The addition to the glycogen reaction mixture of NaOH to final concentrations ranging from 0.08 to 0.10 N resulted in maximum destruction of residual NADP+. The TABLE 1. Assay conditions ofglycogen synthetase activity Components present in glycogen synthetase reaction mixture
NADH oxidized (10-7 M) 30 mine"
Complete"
6.2 0 1.5 0 0
60 mim
12.3
Without cell extract 0 Without glycogen 2.3 Without UDPG 0.8 Without glycogen and UDPG 0 a Incubation time at 37 C. b Complete reaction mixture contained 50 mM glycylglycine buffer (pH 8.0), 10 mM UDPG, 5 mM EDTA, and 18 mM glycogen (standardized as glucose units). Late-aggregation cell extract (75 ,ug, wet weight) was added to 50 ,lp of the reaction mixture. Assayed in 1 ml of UDP reaction mixture as described in the text.
remaining NADPH was amplified by adding 6 N NaOH containing 0.03% H.,O., to a final NaOH concentration of 1.0 N, or by enzymatic cycling. Glycogen synthetase activity was measured in specific cell types by enzymatic cycling involving the coupled reduction-oxidation of NAD+. The level of NAD+ produced by 0.1 ,ug (dry weight) of cells ranged from 10-12 to 10-'5 mol. Both cycling enzymes, GAPDH and GDH, contained NAD+ as a contaminant, which was removed by treatment with Norite. The maximum velocities, in micromoles per minute per milligram of protein, for GDH and GAPDH before and after Norite treatment were 5.83 and 13.1 and 5.00 and 16.8, respectively. Glutamate accumulation in the cycling reaction was determined by the levels of GDH and GAPDH and by the amount of NAD+ added to the cycling reaction mixture. An optimum rate of cycling was achieved with a GAPDH/GDH unit ratio of approximately 70:30. The steady-state level of NADH occurring during cycling was observed directly on the fluorometer in a preliminary test of the cycling reaction mixture (7). The ratio of the reduced to oxidized nucleotide was found to increase with the ratio of GAPDH to GDH. Maximum cycling coincided with a nucleotide ratio of 40:60, reduced to oxidized form. Figure 1 shows that the cycling reaction was linear with increasing amounts of NAD+. UDP levels produced during the glycogen synthetase reaction fell within this concentration range. Localization of total enzyme activity at the midculmination (20-h) stage of development. Figure 2 shows the localization of total glycogen synthetase specific activity in prespore cells and along the vertical axis of the developing stalk from apex to base. This is the first stage in the developmental cycle in which stalk and spore cells are sharply separated and in which cell specific enzyme activity could be detected. For the individual depicted in Fig. 2, the stalks were dissected into five equal sections. The prespore cells and the apical stalk cells showed similar specific activities of 105 + 24 and 115 mmol/h per kg, respectively. There was a decreasing gradient of specific activity from apex to the bottom section of the stalk, which had a specific activity of approximately 30 mmol/h per kg. Localization of total enzyme activity at the late-culmination (23-h) stage of development. The distribution of total glycogen synthetase specific activity between prespore cells and stalk cells at 23 h is shown in Fig. 3. At the late culmination stage of development, the spore mass had lifted off the substratum. The
LOCALIZATION OF GLYCOGEN SYNTHETASE.
VOL. 127, 1976
12
0
E 0
>W6
o
I
z
0
87
Glycogen synthetase dependence on soluble glycogen primer during differentiation of spores. Since changes in primer dependence of glycogen synthetase have been shown during the differentiation of D. discoideum (12, 16), assays were made to determine whether this also occurred in spore cells free from stalk materials. Soluble glycogen primer dependence of enzyme activity was found in prespore cells of preculmination, but not in the mature, spore cells (Table 2). This primer dependency was demonstrated by an eightfold increase in specific activity when prespore tissue was incubated in the presence of soluble glycogen. There was little difference, however, in the specific activities of sorocarp spores incubated with or without soluble glycogen. The preculmination prespore cells showed an increase in specific activity from 19 to 147 mmol/h per kg when soluble glycogen was added to the incubation mixture. The spore cells from sorocarp showed no change with specific activity remaining at about 200 mmol/h per kg with or without soluble glycogen in the incubation mixture.
10 8 6 NAD+ LEVEL (pmoles) 2
4
FIG. 1. Linearity of cycling with increasing amount of NAD+. NAD+ was added to 50 ,d of cycling mixture.
stalk was nearly completed and visibly protruding from beneath the spores. When the dry individual was dissected at this stage, the spore mass neatly separated from the stalk, leaving the intact stalk with the apical cells attached. The prespore cells and the apical stalk cells showed similar mean specific activities of 162 and 150 mmol/h per kg, respectively. As we found at midculmination, the stalk showed a decreasing gradient of activity with the bottom section having the lowest activity. Localization of total glycogen synthetase activity at sorocarp (24 h). Figure 4 shows the localization of total glycogen synthetase specific activity in the spore cells and along the vertical axis of the completed stalk from apex to base. As was the case with the mid- and late-culmination stages, the spore cells and apex of the stalk had similar specific activities of approximately 200 mmol/h per kg. There was a decreasing gradient of activity from the apex to the first section of stalk located immediately below the sorus. At this point in the stalk, the specific activity was approximately 30 mmol/h per kg. Beyond this section of stalk and continuing to the base of the stalk, there was little or no measurable glycogen synthetase activity remaining.
e
>,120
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LAU
wL a-
(A
601
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1o
0+ PRESPORE
STALK
FIG. 2. Total glycogen synthetase activity in prespore and stalk cells at the midculmination (20-h) stage of development. Measured as UDP production with enzymatic cycling for amplification. Prespore point represents mean + standard deviation of 11 replications. Stalk points represent one determination from indicated location in the stalk. These data are representative of two assays. Specific activity is expressed as millimoles per hour per kilogram ofdry
weight.
88
H
HARRIS AND RUTHERFORD
J. BACTERIOL.
1801-
H H H
135H
1
H
lL H LLJ ()
125-
901U U_
45k
FIG 4.Ttlgyoe
4ytets ciiyi/pr
0 ol L
,I
PRESPORE
I
STALK
FIG. 3. Total glycogen synthetase activity in prespore and stalk cells at the late culmination (23-h) stage ofdevelopment. Prespore point represents mean + standard deviation of five replications. Stalk points represent one determination from indicated location in the stalk. These data are representative of three assays. The variation irn values taken from similar areas ofprespore or prestalk cells from different individuals was always less than 20% for all stages of development. Specific activity is expressed as millimoles per hour per kilogram of dry weight.
In addition, further tests were made to determine whether the prespore enzyme could utilize the primer from sorocarp spore cells. In an experiment in which primer-independent activity was measured in prespore cells and sorocarp spore cells separately and mixed, no additive effect was found when the two cell types were mixed (Table 2). This result showed that under the experimental conditions the enzyme from the prespore cells could not utilize the primer employed by the enzyme in the sorocarp spores. Furthermore, prespore extract did not contain an inhibitor that would reduce the activity of the enzyme from sorocarp extract.
DISCUSSION Cell-specific events occurring during spore differentiation. Some of events known to be associated with the differentiation of prespore cells include: (i) the accumulation of inorganic phosphate, (ii) the degradation of soluble glycogen, (iii) the conversion of glycogen synthetase from a soluble to a particulate form, and (iv)
0 L-r1i
I
SPORE
STALK
FIG. 4. Total glycogen synthetase activity in spore and stalk cells at the sorocarp (24-h) stage of development. Spore point represents mean ± standard deviation offive replications. Stalk points represent one determination from indicated location in the stalk. These data are representative of two assays. Specific activity is expressed as millimoles per hour per kilogram of dry weight. TABLE 2. Dependence ofglycogen synthetase activity on soluble glycogen primer in prespore and spore cells Sp acP' (mmol/h per kg, dry wt) Stage of development
-Glycogen +Glycogenb Preculmination (18 19 + 4 (20Y 147 ± 5 (13) h') Sorocarp (24 h) 198 ± 16 (13) 221 ± 30 (6) Preculmination plus 191 ± 17 (7) sorocarpY' a Measured as UDP produced as described in the text. Enzymatic cycling was used for amplification. b Glycogen added to a final concentration of 18 mM (as glucose units). ¢ Hours after removing bacterial food source. d Mean + standard deviation. Number of replications in parentheses. " Preculmination prespore cells (0.13 + 0.01 ,ug, dry weight) added to sorocarp spore cells (0.15 + 0.03 gg, dry weight).
the accumulation of trehalose. In the development of spore cells, inorganic phosphate (Pi) accumulates to an in vivo concentration of 30 mM at the sorocarp stage of development (C.
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LOCALIZATION OF GLYCOGEN SYNTHETASE
Rutherford, J. Embryol. Exp. Morphol., in press). Pi is known to exert multiple effects on both glycogen phosphorylase and soluble glycogen synthetase activities. The glycogen phosphorylase activity is stimulated whereas the activity of the soluble glycogen synthetase is inhibited by Pi. The in vivo levels of Pi localized in the prespore cells at culmination are below Km values for glycogen phosphorylase. Thus, the accumulation during spore maturation would result in increased activitv of this enzyme. These changes in phosphorylase and synthetase activities would lead to an increased capacity to degrade glycogen and to a decreased capacity to synthesize glycogen. A corresponding loss of glycogen is actually observed in these prespore cells; as during sorocarp construction, the glycogen level in the developing spore cells decreased from 30 to 3 mM (Rutherford, in press). The conversion of spore cell glycogen synthetase from a soluble to an insoluble form of the enzyme results in a situation favorable for the synthesis of insoluble glycogen and a decrease in the synthesis of soluble glycogen. An important result of soluble glycogen degradation might be the release of glycogen synthetase from a glycogen-enzyme complex. It has been suggested that some of the soluble enzyme becomes entrapped in the cell wall matrix as it is constructed (17). Therefore, some of the released glycogen synthetase may become insoluble. This change from a soluble to an insoluble form results in a loss of the Pi inhibition. An in vivo Pi concentration of 20 mM is known to inhibit soluble glycogen synthetase by 73% (2), whereas insoluble glycogen synthetase is inhibited by only 28% (13). Therefore, it is likely that the association of a fraction of the soluble glycogen synthetase with the spore cell wall protects it from the inhibitory effects of Pi. As glycogen synthetase becomes insoluble, the dependence of the enzyme on G-6-P levels is overcome. Rosness et al. (10) have demonstrated that the soluble form of glycogen synthetase changes from an I form early in development to a D form at culmination. We have shown that in developing spore cells some insoluble glycogen synthetase is in the I form. G-6-P levels decrease in spore cells, presumably as a result of utilization by the cellulose and trehalose synthetic pathways. Since the soluble glycogen synthetase in spore cells is in the D form, the insoluble I form of the enzyme could successfully compete for UDPG even in the presence of low G-6-P levels. Considering the spore-specific characteristics of glycogen synthetase, glycogen phosphoryl-
89
ase, Pi, and glycogen, we propose a preliminary model of the events associated with glycogen degradation during prespore cell differentiation. As Wright (13) has pointed out, a kinetic situation favoring glycogen breakdown is created in spore cells as a result of both an increase in glycogen phosphorylase activity and the accumulation of Pi. The accumulation of Pi would further decrease glycogen synthesis in prespore cells by inhibition of glycogen synthetase. Upon degradation of glycogen, the synthetic enzyme may become associated with the developing spore wall. The spore wall, or a small portion of the soluble glycogen pool now bound to the spore wall, could act as a primer for the synthesis of "insoluble" glycogen. Both insoluble glycogen synthetase and insoluble glycogen may remain entrapped in the cell wall until germination. Upon germination, cellulases may degrade the cellulose wall, thus releasing the enzyme and glycogen from the cell wall matrix. Thus, the newly emerged myxamoeba would contain both soluble glycogen synthetase and the soluble glycogen primer required for further production of soluble glycogen later in development. Glycogen degradation during prestalk cell migration. Glycogen synthetase specific activity decreases from the apex to base of the stalk. Those cells at the base of the stalk were the first prestalk cells to enter the position of stalk construction. Therefore, the decreasing gradient of activity may be a reflection of the degree of maturation of stalk cells. We cannot determine from the present study the reason for the decrease in activity in mature stalk cells. Since protein degradation is thought to provide an energy source for differentiation in D. discoideum (14), the loss of activity could be the result of degradation of the enzyme as well as other proteins in stalk cells. Alternatively, the loss ofactivity could be due to a conversion from an active to a less active form of the enzyme. The enzyme activity obtained from microgram quantities of lyophilized tissue was compared to levels in tissue homogenates and found to be similar when expressed on a dry weight basis. The glycogen synthetase activity from extracts of freeze-dried tissue (both pseudoplasmodium and sorocarp) was also compared with that from extracts prepared with a French pressure cell or by sonic treatment. Results from all methods of extraction were not significantly different. Thus, the preparation of extracts by freezing followed by lyophilization was sufficient to release enzymes and substrates to the assay conditions. Furthermore trehalase, acid phosphate, and Pi levels increase during stalk
90
HARRIS AND RUTHERFORD
cell maturation (Rutherford, in press). Since these enzymes can be assayed in the lyophilized tissue, the freeze-dry treatment apparently disrupts the stalk cells. Thus, the lack of glycogen synthetase activity in stalk cells is not due to an inaccessibility of the enzyme to the reaction mixture. Regardless of the mechanism for the loss of glycogen synthetase activity in stalk cells, the result is a cell-specific kinetic situation favorable for glycogen degradation. Interestingly, glycogen phosphorylase is active in stalk cells (Rutherford, in press). Likewise, Pi shows an increasing gradient from the apex of the stalk to the base (Rutherford, in press). As mentioned previously, accumulation of Pi would increase glycogen phosphorylase activity but inhibit glycogen synthetase. The levels of glycogen in stalk cells show a striking correspondence to the observed synthetic and degradative activities. Prestalk cells, which have not yet migrated into the area of stalk construction, showed no loss of glycogen (Rutherford, in press). Subsequently, as prestalk cells migrated beneath the apex of the stalk, glycogen degradation occurred rapidly with a decreasing gradient toward the base of the stalk. Thus, the specific cells showing a loss of the glycogen synthetic enzyme exhibit an identical decrease in glycogen levels. Although the regulation of glycogen degradation and resultant synthesis of end products are likely to be under multiple control mechanisms, the observed retention of glycogen synthetase in spore cells and the loss of activity in stalk cells must be included in future models for differentiation of the two cell types. ACKNOWLEDGMENTS This research was supported by the Brown-Hazen Program of The Research Corporation. LITERATURE CITED 1. Firtel, R. A., and J. Bonner. 1972. Developmental control of a-1,4-glucan phosphorylase in the cellular slime mold Dictyostelium discoideum. Dev. Biol. 29:85-103.
J. BACTEIUOL. 2. Gezelius, K., and B. E. Wright. 1965. Alkaline phosphatase in Dictyostelium discoideum. J. Gen. Microbiol. 70:309-327. 3. Hames, B. D., G. Weeks, and J. M. Ashworth. 1972. Glycogen synthetase and the control of glycogen synthesis in the cellular slime mold Dictyostelium discoideum during cell differentiation. Biochem. J. 126:627-633. 4. Jones, T. H. D., and B. E. Wright. 1970. Partial purification and characterization of glycogen phosphorylase from Dictyostelium discoideum. J. Bacteriol. 104:754-761. 5. Liddel, G. U., and B. E. Wright. 1961. The effect of glucose on respiration of the differentiating slime mold. Dev. Biol. 3:265-276. 6. Lowry, 0. H. 1941. A quartz fiber balance. J. Biol. Chem. 140:183-189. 7. Lowry, 0. H., and J. V. Passonneau. 1972. A flexible system of enzymatic analysis. Academic Press Inc., New York. 8. Matschinsky, F. M., C. L. Rutherford, and L. Guerra (ed.). 1968. Proceedings of the 3rd International Congress of Histochemistry and Cytochemistry. Springer-Verlag, New York. 9. Passonneau, J. V., P. D. Gatfield, D. W. Schultz, and 0. H. Lowry. 1967. An enzymic method for measurement of glycogen. Anal. Biochem. 19:315-326. 10. Rosness, P. A., G. Gustafson, and B. E. Wright. 1971. Effects of adenosine 3',5'-monophosphate and adenosine 5'-monophosphate on glycogen degradation and synthesis in Dictyostelium discoideum. J. Bacteriol. 108:1329-1337. 11. Ward, C., and B. E. Wright. 1965. Cell wall synthesis inDictyostelium discoideum. I. In vitro systhesis from uridine diphosphoglucose. Biochemistry 4:2021-2027. 12. Wright, B. E. 1966. Multiple causes and controls in differentiation. Science 153:830-837. 13. Wright, B. E. 1973. Critical variables in differentiation. Prentice-Hall, Inc., Englewood Cliffs, N.J. 14. Wright, B. E., and M. L. Anderson. 1960. Protein and amino acid turnover during differentiation in the slime mold. I. Utilization of endogenous amino acids and proteins. Biochim. Biophys. Acta 43:62-66. 15. Wright, B. E., and D. Dahlberg. 1967. Cell wall synthesis in Dictyostelium discoideum. II. Synthesis of soluble glycogen by a cytoplasmic enzyme. Biochemistry 6:2074-2079. 16. Wright, B. E., D. Dahlberg, and C. Ward. 1968. Cell wall synthesis in Dictyostelium discoideum: a model system for the synthesis of alkali-insoluble cell wall glycogen during differentiation. Arch. Biochem. Biophys. 124:380-385. 17. Wright, B. E., C. Ward, and D. Dahlberg. 1966. Cell wall polysaccharide synthesis in vitro catalyzed by an enzyme from slime mold myxamoebae lacking a cell wall. Biochem. Biophys. Res. Commun. 22:352-356.
