Trehalase from the dormant spore ofDictyostelium discoideum

EXPERIMENTAL MYCOLOGY9, 108-115 (1985)

Trehalase from the Dormant Spore of Dictyostelium KATHLEEN St. John’s Universizy, Department

of

discoideum

A. KILLICK

Biological

Accepted for publication

Sciences, New York, New York 11439 December 18, 1984

KILLICK, K. A. 1985. Trehalase from the dormant spore of Dictyostelium discoideum. Experimental Mycology 9, 108- 11.5. Following purification of dormant spores of the cellular slime mold Dictyostelium discoideum, trehalase (a,-cr’-trehalose-I-D-glucohydrolase, EC 3.2.1.28) activity was solubilized by a freeze-thaw cycle followed by passage of the crude homogenate through a French pressure cell. Enzyme specific activity for spore trehalase was 1.8 nmollminimg, a value no greater than 10% of that observed for the crude myxamoebal enzyme. Assay of samples of myxamoebal and spore enzyme singly and after they were mixed together resulted in values additive for trehalase activity from the two cell types. Enzymatic activity was stable to repeated freeze-thaw cycles, storage at 6”C, and temperatures up to about 50°C. Based upon analysis of kinetic data with Arrhenius plots, the Q,, and energy of activation were estimated (between 30 and 40°C) as 1.95 and 12.0 2 0.5 kcal/mol, respectively. The pH and temperature optima for maximal activity were 5.5 and 46 to 5O”C, respectively. The apparent K, for spore trehalase was 1.2 mM trehalose as estimated from Lineweaver-Burk double-reciprocal plots of initial velocity data versus substrate concentration. Electrophoretic characterization of the spore enzyme indicated the presence of a single peak of trehalase and that the latter had a relative mobility equal to that of the enzyme purified from vegetative myxamoebae (i.e., isozyme I). The developmentally regulated form of trehalase, i.e., isozyme II, was not detectable. o 1985Academic PESS, IX. INDEX DESCRIPTORS:trehalase; Dictyostelium; spore; germination; latency; trehalose; cell type specificity.

The life cycle of the cellular slime mold Dictyostelium discoideum consists of growth, differentiation, cellular aging, and maturation. In the absence of an exogenous source of nutrients, development is initiated and the myxamoebae stream together to form multicellular aggregates, each of which is subsequently transformed over a period of 24 h (22°C) into a mature fruiting body or sorocarp consisting of a sorus or spore mass which is supported by a cellulose-ensheathed stalk. Activation of the dormant spores of Dictyostelium (Cotter and Raper, 1970; Cotter, 1975) results in germination and the release of single-cell myxamoebae. The emergent cells, under appropriate nutritional conditions, undergo mitosis and reenter the growth phase of the life cycle. During the above morphological processes, there is an accumulation of struc-

tural polysaccharides, i.e., cell wall glycogen, acid mucopolysaccharide, and cellulose and the nom-educing disaccharide, ol,ol’-trehalose (Ceccarini and Filosa, 1965; Killick and Wright, 1974). The latter sugar serves as a major energy and carbon source during spore germination, at which time its hydrolysis to glucose is catalyzed by the enzyme trehalase (ol,cx’-trehalose 1-D-glucohydrolyase, EC 3.2.1.28). Although trehalase activity is detectable at all stages of the Dictyostelium life cycle, dramatic changes in the specific activity occur during morphogenesis. During the first 12-14 h of development, enzyme specific activity decreases about lo-fold as a result of trehalase secretion. During the remainder of morphogenesis, at a time when fruiting body formation occurs, the specific activity of the intracellular enzyme increases so that the amount of trehalase ob108

0147-.5975/85 $3.00 Copyright Q 1985 by Academic Press, Inc. All rights of reproduction in any form reserved.

SPORE TREHALASE

served in extracts from mature sorocarps is about 40% of that found in extracts from vegetative cells on an enzyme unit per cell sample basis. The most unique aspect of this increase in trehalase activity during sporulation is that it occurs during a period of development when trehalose-6-P synthase activity in vitro and the rates of both trehalose synthesis and accumulation in vivo are maximal. Although trehalose turnover during development may be essential to stalk cell development and maturation, accumulation of this saccharide in the apparent absence of turnover is unique to the spore cells, where it serves as the major carbohydrate reserve during dormancy. Although trehalase from vegetative cells and the extracellular fluid of mature sorocarps of Dictyosteliznm (Killick, 1981, 1983a,b) has been purified and characterized, little attention has been paid to the enzyme from dormant spores. As part of an ongoing series of investigations involved with determining various mechanisms critical to maintaining the in vivo latency of spore cell trehalase, this enzyme has now been purified and characterized from dormant Dictyostelium spores. The results of these studies form the basis of the present report. MATERIALS

AND METHODS

Materials. Trehalose dihydrate, maltose, cellobiose, glycine, streptomycin sulfate, maleic acid, 2-[N-morpholinolethanesulfonic acid,’ N-tris(hydroxymethyl)methyl-glycine (Tricine), crystalline bovine serum albumin, and 4-morpholinopropanesulfonic acid (Mops) were purchased from Sigma Chemical Company; Tris (enzyme grade) and Ultrapure ammonium sulfate from Schwartz/Mann; DEAE (DE-.52)-cellulose, microgranular anion exchanger, ’ Abbreviations used: Mes, 4-morpholineethanesulfonic acid; Tricine, N-[2-hydroxy-l.l-bis(hydroxymethyl)ethyl]glycine; Mops, 4-morpholinepropanesulfonic acid.

FROM Dictyosteliurn

from Whatman; purified acrylamide, N,N’methylenebisacrylamide, ammonium persulfate, and N,N,N’, N’- tetramethylethylenediamine (TEMED) from Miles Labs; and sucrose from Pfanstiehl. All other chemicals were of the highest analytical grade available. Organism and culture conditions. D, discoideum strain NC-4 (ATCC 24697) was grown on nutrient agar with Esc~e~ic~ia coli as the bacterial associate according to methods previously described (Killick, 1983a). Initiation of differentiation. Upon depletion of the bacterial food source, the stationary phase amoebae were harvested from the nutrient agar surfaces and washed free of residual bacteria by repeated centrifugation in the presence of cold distilled water. To initiate differentiation, the washed cells were spread onto sheets of nonnutrient agar and incubated at 22°C either for 2 h (2-h starved myxamoebae) or for 1 to 3 days (mature sorocarp stage of development). Preparation of cell-free myxamoebal extracts. After 2 h starvation, cells were harvested from the nonnutrient agar sheets with 50 mM Mes-NaOH (pH 6.5) buffer. The resulting cell suspension was lightly homogenized and then centrifuged for 2 min at 3000g (o”C) and the supernatant fluid was discarded. The cell pellets were dispersed into a homogenous solution using SO mM Mes-NaOH (pH 6.5) buffer and the cells were then ruptured by freezing in a dry-ice/acetone bath followed by a gradual thawing at 23°C. The resulting homogenate was centrifuged at 33,000g (30 mm) and the supernatant liquid served as the source of trehalase activity for all subsequent manipulations, which were performed at 4°C unless otherwise indicated. Isolation of spores. Sorocarps were harvested from nonnutrient agar sheets with IO mM potassium phosphate buffer (pH 6.5). After the organisms were lightly homogenized with a Teflon pestle, the cell suspen-

110

KATHLEEN

sion was centrifuged at 2000 rpm for 5 min in an International portable refrigerated centrifuge, Model PC-2 (4°C). The supernatant fluid was discarded and the cell pellet was resuspended in 10 mM potassium phosphate buffer (pH 6.5). The cell suspension was vortexed in order to dislodge the spore cells from the stalks. Subsequent separation of the two cell types was effected by filtration of the suspension through two layers of very fine nylon mesh. The filtrate was collected and the spores were pelleted by low-speed centrifugation in a Sorvall RC-2B centrifuge (4°C). The supernatant liquid was discarded, and the spores were washed twice with 10 mM potassium phosphate buffer (pH 6.5) and frozen in a dryice/acetone bath. Phase-contrast microscopy of the spore preparation prior to the freezing step indicated the absence of stalk material. Assay of trehalase activity. Trehalase activity was assayed at 35°C in an incubation mixture that contained 25 mM trehalose and 50 mM potassium citrate (pH 5.5) buffer as previously described (Killick, 1983b). One unit of enzymatic activity is defined as that amount of enzyme that catalyzes the synthesis of 1 umol of product/ 30 min at 35°C unless specified otherwise. The specific activity is expressed as enzyme units per milligram of acid-precipitable protein. Measurement of glucose. After termination of the trehalase reaction by boiling, the samples were chilled in an ice bath. The pH of the solution was adjusted to 7.0 with 100 mM potassium phosphate (pH 10.5) buffer and the final volume was adjusted to 2.4 ml with distilled water. Aliquots of solution were analyzed for glucose content as previously described (Killick, 1983b). Measurement of protein. Protein was measured according to the methods of Lowry et al. (1951). Analytical polyacrylamide gel electrophoresis. Electrophoresis was performed at 4°C according to the procedures described

A. KILLICK

by Ornstein (1964) and Davis (1964). Separating gels were cast at total acrylamide (%T) and bisacrylamide (%C) concentrations of either 8.7 and O.S%, respectively, or 5 and 3.2%, respectively. Stacking gels consisted of 3.1% total acrylamide plus 20% bisacrylamide. Gels were cast in glass tubes and 50- to 100~~1 samples of enzyme containing 10% (w/v) sucrose were layered onto the gel tops together with bromphenol blue as the tracking dye. The run was initiated and maintained at 1 mA/gel until the dye front was about 0.5 cm from the end of the tube. At the end of the run, the gels were removed from the tubes and sliced into fractions which were then transferred to, and emulsified in, 0.5-1.0 ml of 25 mM Tris-maleate (pH 6.5) buffer. Aliquots of gel eluate were then assayed for trehalase activity. RESULTS

Trehalase Activity in Spore Extracts To effect maximal release of soluble enzymatic activity, spores were subjected to a single freeze-thaw cycle followed by passage of the cell suspension through a French pressure cell (20,000 psi) (Table 1). The latter treatment increased recoverable trehalase activity about loo-fold over that achieved with a freeze-thaw cycle alone. With these extraction procedures, at least 95% of the trehalase activity in the crude homogenate was recoverable in the 33,000g supernatant fraction. With this source of enzyme, product formation was linear with TABLE 1 Trehalase Activity versus Method of Spore Rupture Rupture method Freeze-thaw Freeze-thaw (2300 psi) Freeze-thaw

Soluble activity0 (% maximum) OS

+ Yeda press + French press

2.8 100.0

’ Soluble activity recoverable in the supernatant fraction following centrifugation of the crude homogenate (33,000g, 15 mm).

SPORE

TREHALASE

time for 90 to 120 minutes of incubation after correction for endogenous glucose production in the absence of trehalose addition to the assay system, and the initial reaction rate was directly proportional to the amount of protein assayed. Examination of product stability under the conditions used for the trehalase assay indicated that, with a glucose concentration of 0.4 mM, about 94% of the initial amount was recoverable after a 90-minute incubation at 35°C. Values usually obtained for specific enzyme activity (i.e., 1.8 nmol/min/mg) were in good agreement with those previously obtained by Ceccarini (1966) and Cotter and Raper (1970). The values for specific activity for the enzymes from myxamoebae, sorocarps, and dormant spores were 13.3, 40, and 1.8 nmol/min/mg, respectively. Since the low specific activity observed for the spore enzyme relative to enzyme from myxamoebae may have resulted from the presence of soluble trehalase inhibitors in the spore extract and/or to the presence of soluble trehalase activators in the amoebae preparation, samples of myxamoebae and spore enzyme were assayed singly and after they had been mixed together. The results indicated that the values for enzymatic activity for the two cell types were additive. Partial

Purification

of Trehalase

Prior to the study of some of the properties of trehalase from dormant spores, the enzyme was partially purified (Table 2). Extracts were prepared from dormant spores by a freeze-thaw cycle followed by a passage of the homogenate through a French pressure cell. Insoluble material was removed by centrifugation (33,OOOg, 30 minutes) and streptomycin sulfate was added with constant stirring to a final concentration of 2% (w/v). Following stirring for 1 h on ice, the suspension was centrifuged (33,OOOg, 15 minutes). Enzyme recovered from the previous step was fractionated with absolute ethanol. Alcohol was added

FROM

Dictyostelium

111

dropwise to a final level of 70% (v/v> with constant stirring of the solution. After 15 minutes of additional stirring, the solution was centrifuged (15 minutes, 33,OOOg) and the supernatant fraction was discarded. The pellet was solubilized with 25 m&I Trismaleate (pH 6.5) buffer and dialyzed overnight against the same buffer. Insoluble ma.terial was removed by centrifugation and aliquots of the supernatant flui plied to columns of DE-52 cellulose ion exchanger previously equilibrated in the same buffer. Enzymatic activity was eluted from the column with a linear salt gradient (O- Z M NaCl). Fractions containing the majority of high-specific-activity enzyme were pooled and concentrated by ultra~ltrat~o~ in an Amicon apparatus equipped with a PM-10 filter. The partially purified enzyme was stored at - 12°C. Properties

of the Enzyme

Studies on the stability of spore trehalase at low temperature demonstrated that enzymatic activity was stable (1) to several repeated freeze-thaw cycles, (2) to incubation at - 12°C for at least 2 months, and (3) for at least 2-4 days at 5°C in a cold room. Enzyme stability to temperatures over the range of 2.5 to 60°C was investigated by subjecting aliquots of partially purified enzyme in 25 mM Tris-maleate (pH 6.5) buffer to a 5-minute incubation at each of several designated temperatures. FoIlowing heat treatment, the enzyme was stored on ice for 30 minutes and the residual trehalase activity was measured. Full enzymatic activity was recovered following incubation of trehalase at temperatures up to about 48-50°C. At higher temperatures, trehalase instability was apparent. For example, after 5 minutes at 55 and 60°C 45 and 15% of the original activity were recovered, respectively. Parallel studies with enzyme purified from myxamoebae indicated comparable results. The kinetic properties of trehalase were examined at 35°C in the presence of 50

KATHLEEN

112

A. KILLICK

TABLE 2 Partial Purification of Trehalase from Dormant Spores of Dictyostelium Fraction 1. 2. 3. 4.

Crude extract Streptomycin sulfate Ethanol,fractionation DE-52 cellulose

Specific activity” (pmol/30 min/mg)

% Recovery

0.053

100 100 81

0.080 0.231 2.32

Fold purification 1.0 1.5 4.4

32

44.0

a Enzymatic activity was measured as described under Materials and Methods.

potassium citrate (pH 5.5). With all assays employed for trehalase activity, product formation was linear with time and the rate was directly proportional to the amount of protein assayed. Examination of the initial velocity of the trehalase reaction as a function of the trehalose level indicated a typical hyperbolic response. The apparent Michaelis constant was 1.2 mM trehalose. The effect of pH on the rate of product formation (using the coupled, discontinuous spectrophotometric trehalase assay) by the partially purified spore trehalase preparation was determined using 100 mM potassium citrate, Mes-NaOH, MopsNaOH, and Tricine-NaOH buffers. The pH for optimal activity was 5.5. Parallel studies using a purified myxamoebal enzyme preparation indicated that enzyme from these cells was likewise maximally active at pH 5.5. Measurement of trehalase activity as a function of temperature over the range of 21 to 67°C indicated maximal activity at about 46 to 50°C. Analysis of kinetic data with Arrhenius plots demonstrated that the Q,, was 1.9 (between 30 and 40°C) (Fig. l), and that the energy of activation (i.e., E,) calculated from the slope of the straight line resulting from a plot of log V,,, versus l/ T(“K) was 12 -t 0.5 kcal/mol. Parallel studies with purified myxamoebal trehalase indicated comparable values for the Q,, and Ea. Further characterization of the spore enzyme was achieved by subjecting aliquots of partially purified enzyme to nondenaturing electrophoresis in polyacrylamide

gels. Following electrophoresis, the gels were sectioned into slices, each of which was then emulsified in 25 mA4 Tris-maleate (pH 6.5) buffer. Trehalase activity in the gel eluates was measured as described under Materials and Methods. The results indicated the presence of a single major peak of trehalase activity (Fig. 2). The relative mobility of the latter enzyme was equal to that of the trehalase from vegetative myxamoebae. When mixtures of the amoebae and spore enzyme preparations were subjected to electrophoresis under the same conditions as described under Materials and Methods, the trehalases from the two sources coelectrophoresed and were not resolvable into multiple forms. As demonstrated previously (Killick, 1983a) and subsequently confirmed in the present report, isozyme I (i.e., the amoebae and spore enI

8 4

2.9

3.0

3.2

3.1 I/T

3.3

34

PK-‘1

FIG. 1. Arrhenius plot of the effect of the assay temperature on the initial velocity of the trehalase reaction.

SPORE TREHALASE

FROM Dictyostelhm

113

prestalk cell migration and subsequent stacking into the cellulose sheath intensified, resulting in the eventual construction of a rigid stalk, intra- and intercellular trehalase gradients became very striking. The results of Jefferson and Rutherford (1976) are enlightening since they demonstrate that (1) the 8- to IO-fold increase in trehalase specific activity occurring between the 14th and 24th h was entirely a consequence of the 30-fold increase in enzyme activity in the prestalkistalk cells; and (2) trehalase activity was specificahy localize in the stalk cells of the slime mold with no enzymatic activity observable in mature, DISTANCE FROM ORIGIN (mm) dormant spores. It is reasonable to assume FIG. 2. Polyacrylamide discontinuous gel electrothat the basis for the observed absence of phoresis of trehalase. Enzyme was partially purified from spores and samples were subjected to electrotrehalase activity stemmed from the small phoresis. Trehalase activity was assayed after gel sec- amounts of biological material used and the tioning and extrusion as described in the text. The low enzymatic activity resultant from the arrow indicates the position of the tracking dye. essentially in situ assay methods employed. zyme) was electrophoretically distinguishRecently, electrophoretic analysis of treable from the developmentally regulated halase from Dictyostelium has demonform of trehalase (i.e., isozyme II). Under strated the presence of multiple activities all conditions, detection of enzymatic ac- of the enzyme during the culmination tivity in gel eluates was dependent upon the cess (Killick, 1983a). Although a single inpresence of trehalose and product formatracellular trehalase activity (isozyme I, the tion was a function of the incubation time vegetative form) was detectable prior to 14 and the enzyme level subjected to electroh, during culmination (fruiting body forphoresis. mation) a second form of the enzyme, isozyme II, accumulated. Since the abservaDISCUSSION tions of Jefferson and Rutherford (1976) The present studies confirm the original have shown that trehalase activity accuobservations of Ceccarini (1966) and Cotter mulated by cells during culmination is speand Raper (1970) concerning the specific cifically localized in stalk tissue, and elecactivity of trehalase in extracts from dor- trophoretic analysis of the spore enzyme mant D. discoideum spores, but are at vari(Fig. 2) has shown only isozyme I to be ance with those reported by Jefferson and present, it may be concluded that the acRutherford (1976). The latter investigators cumulation of isozyme II is restricted to employed ultramicrochemical techniques stalk tissue. and enzymatic cycling to investigate the inTrehalase activity may be modulated in tercellular compartmentation of trehalase Dictyostelium by (1) alterations in enzyme between spore and stalk cells during develconcentration at the cellular, organelle. opment. Their results indicated that, at the and/or microenvironmental levels, (2) intrapseudoplasmodial stage, trehalase specific cellular and intercellular comp~rtme~tatio~ activity was low and nearly identical in the phenomena, and (3) post-translational (e.g., two cell types. At later stages (i.e., preculcovalent) modification mechanisms during mination, culmination, and sorocarp) as germination. Trehalase specific activity is

114

KATHLEEN

maximal during the myxamoebal emergence stage, which is characterized by the dissolution of the innermost cellulosic wall layer. In extracts from dormant spores, the enzyme has a specific activity of 0.05-0.08 pmo1/30 min/mg, while in extracts from either myxamoebae, prepared 5 h after heatshock treatment of the spores, or from vegetatively growing cells, the average value is 0.45 pmol/30 min/mg. The increase in trehalase activity during germination is sensitive to inhibition by both actinomycin D and cycloheximide as is also cell emergence from the spore case (Demsar et al., 1983). The effects of cycloheximide on trehalase activity are reversible by washing the cells. When this is done, however, an overshoot in enzyme specific activity occurs. The superinduction of trehalase activity that occurs during reversal of cycloheximide-inhibited germination suggests that, although an increase in the rate of trehalase synthesis during germination may be a principal factor underlying the observed increase in enzyme activity, an alteration in the rate of enzyme turnover during this time period may also play a contributory role. To more fully understand the basis for the alterations in trehalase specific activity during germination, the rates of synthesis and degradation of this enzyme prior to, during, and following the germination process must be studied utilizing standard radioimmunochemical methodology. Analysis of this aspect of metabolic regulation in Dictyostelium is presently being pursued in this laboratory. Since trehalase activity is maximal in vitro during the emergence stage of germination (i.e., between the 3rd and 5th h), it has been suggested that the rate of trehalose degradation in vivo is also maximal at this time. To determine whether the increases in trehalase specific activity are accompanied by corresponding alterations in the endogenous trehalose pool, Jackson et al. (1982) assayed trehalase activity and trehalose levels at different stages of germi-

A. KILLICK

nation. Most of the trehalose was catabolized during the swelling stage (i.e., the first 2 h of germination), namely about 2 to 3 h before the maximal increase in trehalase activity was detected in vitro. Thus, these investigators concluded that (1) the trehalase, apparently synthesized de nova during the emergence stage, does not play a role in spore germination since the endogenous trehalose pool has been depleted by the time that this stage of germination is reached, and (2) endogenous (i.e., basal) trehalase activity associated with dormant spores is responsible for trehalose catabolism during germination. The physiological function of endogenous spore trehalase appears to be that of catalyzing the hydrolysis of stored trehalose during the swelling stage. Experimental evidence has indicated that during the latter germination stage not only does de IZOVOtrehalase synthesis occur but that the trehalase level is also regulated by alterations in its rates of turnover and secretion. In contrast to endogenous trehalase, the function of the enzyme synthesized de nova during emergence is unknown. Whether the activity of this enzyme is expressed in vivo during either germination or vegetative growth is likewise unknown. Studies in this laboratory are presently concerned with determining the mechanisms underlying the alteration in trehalase specific activity during the emergence stage as well as the physiological function(s) of this enzyme during the germination and growth phases of the life cycle. ACKNOWLEDGMENTS These studies were supported by Research Grants AG 04316 and GM 32094 from the National Institute of Health. REFERENCES CECCARINI,C. 1966. Trehalase from Dictyostelium discoideurn: Purification and properties. Science (Washington, D.C.) 151: 454-456. CECCARINI, C., AND FILOSA, M. 1965. Carbohydrate

SPORE TREHALASE content during development of the slime mold, Dietyosteiium discoideum. J. Cell Comp. Physiol. 66: 135-140. COTTER, D. A. 1975. Spores of the cellular slime mold Dictyostelium discoideum. Spores VI: 61-72. COTTER, D. A., AND RAPER, K. B. 1970. Spore germination in Dictyostelium discoideum: Trehalase and the requirements for protein synthesis. Dev. Bioi. 22: 112-128. DAVIS, B. J. 1964. Disc electrophoresis. II. Method and application to human serum proteins. Ann. N. Y. Acad. Sci. 121: 404-427. DEMSAR,

1. H.,

COTTER,

D. A.,

AND CHAN,

A. H.

1983. Ultraviolet light-induced inhibition of enzyme synthesis during Dictyostelium discoideum spore germination. Exp. Mycol. 7: 95-108. JACKSON, B. BL., CHAN, A. H., AND COTTER, D. A. 1982. Utilization of trehalose during Dktyosrelium discoideum spore germination. Dev. Biol. 90: 369374.

JEFFERSON,B. L., AND RUTHERFORD, C. L. 1976. A

FROM Dietyostelium

115

stalk-specific localization of trehalase activity in Dictyostelium discoideum. Exp. Cell Res. 103: 127134.

KILLICK, K. A., AND WRIGHT, B. E. 1974. Regulation of enzyme activity during differentiation in Dictyostelium discoideum. Annu. Rev. Microbial. 28: 139166. KILLICK, K. A. 1981. Methods for the rapid preparation of trehalase from Dictyostelium discnideum. Prep. Biochem. 11: 547-557. KILLICK, K. A. 1983a. Multiple forms of trehalase during development in Dictyostelium discoideum. Exp. Mycol. I: 66-73. KILLICK, K. A. 1983b. Trehalase from the cellular slime mold, Dictyostelium discoideum: Purification and characterization of the homogeneous enzyme from myxamoebae. Arch. Biochem. Biophys. 222: 561-573.

ORNSTEIN, L. 1964. Disc electrophoresis. I. Background and theory. Ann. N. Y. Acad. Sci. 121: 321349.