ARCHIVES

OF BIOCHEMISTRY

Trehalose

AND

Synthesis

Preparation,

KATHLEEN Boston

Biomedical

Research

170,

BIOPHYSICS

634-643

during Differentiation discoideum l

Stabilization

and Assay Synthetase

A. KILLICK Institute,

(1975)

Department Received

AND

of Trehalose-6-Phosphate

BARBARA

of Developmental April

in Dictyostelium

E. WRIGHT2 Biology,

Boston,

Massachusetts

02114

7, 1975

The developmental profile for the enzyme, trehalose-6-phosphate synthetase was reexamined during morphogenesis in Dictyostelium discoideum, using experimental conditions that appear to optimize enzyme specific activity at each stage of development. The basis of the low-temperature instability of trehalose-6-P synthetase activity was examined and found to be a function of pH. Enzymatic activity could be stabilized at low temperature by preparation of extracts throughout development in pH 7.5 buffer containing 10 mM phosphate and 25 mM trehalose. Synthetase activity was determined by measuring the rates of synthesis of UDP and trehalose with a calorimetric and radiometric assay, respectively. In the latter analysis, UDP-[U-14C]glucose was used as substrate and [“Cltrehalose was isolated and subsequently quantitated using charcoal columns and thin-layer chromatography. With these methods, it was possible to demonstrate that 1) trehalose-6-P synthetase activity is detectable at and prior to 5 h of development, 2) enzyme specific activity increases approximately five- to sixfold during differentiation, 3) total enzyme activity per cell aliquot is constant from about the 10th (aggregation) to the 24th h (sorocarp) of development, and 4) enzyme specific activity is constant from the preculmination (17th h) to the sorocarp (24 h) stage of development. The constancy of enzyme activity (units per cell aliquot) from the 10th to the 24th h of development suggests that a sudden rise in overt trehalose-6-P synthetase activity, resulting from an induction of enzyme synthesis, is not a primary critical variable in the increased rate of trehalose synthesis which occurs in vivo during the culmination process (i.e., 20-22 h).

In the absence of an exogenous source of nutrients (e.g., bacteria or defined media), the unicellular myxamoebae of the cellular slime mold Dictyostelium discoideum stream together to form multicellular aggregates (1). A series of precise and intricate cell movements culminates in the transformation of each aggregate into a fruiting body (sorocarp), which consists of a multispore mass supported by a cellulose-ensheathed stalk. These morphologi’ This paper is paper 2 This investigation lowship (No. lF03 GM by research grants No. the National Institutes

V in the series. was supported by Special Fel53117-01) to K. A. Killick and HD 05357 and HD 04667 from of Health to B. E. Wright. 634

Copyright 0 1975 by Academic Press, Inc. All rights of reproduction in any form reserved.

cal transformations are completed within 24-28 h at 23°C and are accompanied by the synthesis and accumulation of several polysaccharides, as well as the nonreducing disaccharide, a,a’-trehalose (2-5). The latter is thought to serve as a major energy and carbon source during the emergence stage of spore germination (6, 7). Trehalose-6-phosphate (trehalose-6-P) synthetase (UDP-glucose: n-glucose-6phosphate l-a-glucosyltransferase, EC 2.4.1.15) catalyzes the synthesis of trehalose-6-P, the immediate precursor of trehalose (8) from UDP-glucose and glucose-6-P. Changes in the specific activity of trehalose-6-P synthetase during differentiation

TREHALOSE-6-P

SYNTHETASE

in D. discoideum were first described by Roth et al. (g-121, who reported that synthetase activity was barely detectable at or prior to the 5th h (i.e., rippled myxamoebae) and increased at least 150-fold (11) between the 3rd and 16th h of development. After the 18th h, enzyme specific activity decreased during the remainder of these morphogenesis. Subsequently, changes in enzyme specific activity were shown not to reflect changing in uiuo rates of trehalose synthesis (13, 14). Sargent and Wright (14) demonstrated that the rate of trehalose synthesis in uiuo was low and constant from aggregation (10 h) until late in the culmination process, at which time (20-22 h), there occurred an approximate loo-fold increase. Moreover, using various parameters of assessment, it was demonstrated (13) that the maximum rate of trehalose synthesis observed in uiuo would require an enzyme specific activity loo-fold higher than the maximum value reported (10). Because of these observations, it was suggested (13) that activation of latent trehalose-6-P synthetase may be a primary critical variable associated with the rapid increase in the rate of trehalose synthesis that occurs during culmination. Evidence for the existence of a latent form of trehalose-6-P synthetase was subsequently obtained by Killick and Wright (15), who demonstrated that this enzyme was present in a masked form in myxamoebae extracts prepared and maintained in pH 6.5 buffer at 23°C. Additional studies (161, however, demonstrated that enzyme specific activity could be appreciably elevated at both the very early and late stages of development by extract preparation in buffer supplemented with trehalose. Hence, it is possible that the in vitro latency of trehalose-6-P synthetase activity previously observed to occur early in differentiation may in part have been the consequence of the suboptimal conditions used. For this reason, the present communication presents a reexamination of the developmental profile of trehalose-6-P synthetase during morphogenesis in Dictyostehum. Conditions of enzyme preparation, stabilization and assay that appear to op-

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DZCTYOSTELZUM

timize enzyme specific activity at each stage of development will be described, as a knowledge of these conditions is an important prerequisite to both enzyme purification and in-depth analysis of those mechanisms by which trehalose-6-P synthetase activity is unmasked as well as regulated during differentiation. EXPERIMENTAL

PROCEDURES

Chemicals. UDP-glucose (sodium salt), UDP (sodium salt), r+glucose-6-P (disodium salt], phosphoenolpyruvate (monopotassium salt), crystalline pyruvate kinase (type II from rabbit skeletal muscle), morpholinopropane sulfonic acid3 (MOPS), 2[N-morsulfonic acid (ME@, Npholinolethane tris[hydroxymethyl]methylglycine (Tricine), cyrstalline bovine serum albumin, a,a’-trehalose dihydrate and Z-mercaptoethanol were purchased from Sigma Chemical Co.; ultrapure Tris from S&wars-Mann; Kieselguhr G (according to Stahl) from Brinkman Instruments; alkaline phosphatase (Escherichia co&, chromatographically purified) from Worthington; uniformly labeled UDP-[U‘V]glucose and Aquasol from New England Nuclear; [U-“Cltrehalose from International Chemical and Nuclear Corporation; Bio-Gel A-l.5 m from BioRad Laboratories. All other chemicals were reagent grade. Organism and culture conditions. D. discoideum strain NC-4 (ATCC 24697) was grown on nutrient agar with Escherichio coli as the bacterial associate according to methods previously described (17). Initiation ofdifirentiation. With the depletion of the bacterial food source, the vegetative myxamoebae of Dictyostelium enter the stationary phase of their growth cycle. At this time, the myxamoebae were harvested from the nutrient agar surfaces and washed free of any residual bacteria by repeated centrifugation in the presence of cold distilled water. To initiate differentiation, the washed cells were spread onto sheets of 2% nonnutrient agar and allowed to incubate at either 15 or 23°C until the desired stage of morphogenesis was reached (14). Under these conditions cellular differentiation pro’ Abbreviations used: MES, 2[N-morphosulfonic acid; MM buffer, 66 rnM 2[Nmorpholinolethane sulfonic acid plus 5 rnM 2-mercaptoethanol; MMT buffer, 50 rnr.r PlN-morpholinolethane sulfonic acid, 5 mM 2-mercaptoethanol and 25 mM trehalose; MOPS, morpholinopropane sulfonic acid; MPMT buffer, 50 rnM morpholinopropane sulfonic acid-NaOH, pH 7.5, containing 10 mM potassium phosphate, 5 mM 2-mercaptoethanol and 25 rnxr trehalose; Tricine, N-tris[hydroxymetbyllmethylglycine.

linolethane

636

KILLICK

AND

ceeds synchronously over a 24-h period, and hence stages of development described in this manuscript correspond to the morphological time scale depicted by Roth and Sussman (10). Preparation of extracts for trehalosex-P synthetase profile studies. The following procedures were carried out at 4°C unless specifically noted otherwise and all pH values correspond to that temperature at which the buffer was used. Cells were harvested from nonnutrient agar sheets with 50 mM MOPS-NaOH buffer (pH 7.5) which contained 10 rnru potassium phosphate, 5 mM 2-mercaptoethanol and 25 mM trehalose (MPMT buffer). The resulting cell suspension was centrifuged for 2 min at 3OOOg (0°C) and the supematant fluid discarded. The cell pellets were dispersed into a homogenous solution using MPMT buffer, and the cells were then ruptured by freezing in a Dry-Ice/ acetone bath, followed by a gradual thawing. At stages of development beyond preculmination (18 h), the thawed homogenate was subjected to a single passage through a French pressure cell (8000 psi). After the homogenate had been centrifuged for 20 min at 33,OOOg (O”C), the supematant liquid was decanted and recentrifuged for an additional 20 min under the same conditions. The resultant cell-free extract usually had a protein concentration of 8-10 mg/ml. In those studies where extracts were prepared and maintained at room temperature (23”C), MPMT buffer was replaced by either 50 mM MES-NaOH buffer (pH 6.5) which contained 5 mM S-mercaptoethanol (MM buffer) or MMT buffer (MM buffer plus 25 mM trehalose). In some experiments sodium thioglycollate was used in place of 2-mercaptoethanol. Assay of trehalosed-P synthetnse activity. Trehalose-6-P synthetase activity was assayed at 23°C in 0.25 ml of an incubation mixture that contained: 10 mM UDP-glucose, 400 mM KCl, 62.2 mM MgCl,, 1 mM EDTA, 5 mM 2-mercaptoethanol, and 50 mM MES-NaOH (pH 6.5) buffer. Glucose-6-P was added to this mixture at a final concentration of either 50 mM (extracts prepared prior to and including early aggregation, i.e., 6-8 h) or 25 mM (all developmental stages subsequent to aggregation). In the radiometric assay, this mixture was supplemented with UDP-[U-‘*C]glucose. Prior to addition of this labeled substrate, the solvent was removed according to the manufacturer’s instructions and the residue was redissolved in distilled water. The final specific radioactivity of the sugar nucleotide used in the assay varied with the developmental stage being studied: 608 cpm/nmol (myxamoebae), 304 cpm/nmol (rippled myxamoebae and early cell aggregates), and 152 cpm/nmol (preculmination and subsequent developmental stages). The reaction was initiated by addition of enzyme and was terminated by a 3-min incubation in a

WRIGHT boiling-water bath. Initial velocities were determined by measuring the rates of synthesis of UDP (calorimetric assay) or trehalose (radiometric assay) at lo-15min intervals over a 60-75min period and were linear functions of time and protein concentrations for extracts prepared at all developmental stages studied (with the exception of UDP formation with smooth myxamoebae extracts). One unit of enzymatic activity is defined as that amount which catalyzes the synthesis of 1 nanomole of UDP (or 1 nanomole of trehalose) per minute at 23°C. The specific activity is expressed as units per milligram of protein. Measurement of UDP. UDP concentrations were measured calorimetrically by use of the methods of Pontis and Leloir (18). Measurement of trehalose. After termination of the synthetase assay, the contents of each tube was adjusted to pH 8.3 by the addition of Tris-HCl buffer to a final concentration of 100 mM. Alkaline phosphatase (10) was then added and the mixture was incubated under a drop of toluene for 20-24 h at 37°C. Following this treatment, two volumes of 95% ethanol were added and the precipitate was removed by centrifugation and washed several times with 70% ethanol (19). The supematant liquid and washings were combined and evaporated to dryness. The residues were dissolved in distilled water and aliquots desalted on charcoal-Celite columns (14). After elution of the sugars with l-propanol (5%), those fractions containing trehalose were pooled, lyophilized to dryness and redissolved in distilled water. Aliquots were spotted onto glass plates (20 x 20 cm) of Kieselguhr G (14) 2 cm from the edge of the plate and run ascending in l-butanol/acetone/O.l M phosphate (pH 6.0) (4:5:1, v/v) to 16 cm above the origin: Each plate was run twice in the solvent, with drying between runs. After identification of the [“Cltrehalose area by its Rr of migration compared to that of standard [‘4C1trehalose run under identical conditions on a companion strip, this region of the Kieselguhr plate was scraped off and the sugar eluted with 70% ethanol. Aliquots of the sugar solution were mixed with 10 ml of Aquasol and their content of radioactivity was measured in a Beckman LS 200 liquid scintillation counter. The amount of trehalose was calculated from the specific radioactivity of UDP-[U-14C]glucose used as substrate. In a separate series of experiments, it was demonstrated that when the radioactivity in the trehalose area of the thin-layer plate was eluted with water, desalted, and subsequently incubated with partially purified trehalase (see below) more than 85% of the eluted radioactivity could be recovered as [‘*Clglucose as determined by thin-layer chromatography. Preparation of trehalase from Dictyostelium for trehalose determination. Cell-free myxamoebae (2-hstarved) extracts were prepared in 50 mM

TREHALOSE-6-P

SYNTHETASE

MES-NaOH (pH 6.5) buffer by a single freeze-thaw of the cells in a Dry Ice/acetone bath followed by centrifugation of the homogenate at 33,OOQg for 20 min (0°C). The supernatant liquid was cooled to 0°C and absolute ethanol was added with stirring to a final concentration of 40%. After stirring for 15 min at 0°C the solution was centrifuged at 33,OOOg for 15 min and the precipitate discarded. Absolute ethanol was added to the supernatant fluid with stirring to a final concentration of 60%. After stirring for 15 min at 0°C the solution was centrifuged for 15 min at 33,OOOg. The supernatant liquid was discarded and the precipitate was solubilized with a minimal amount of 25 mM Tris-maleate (pH 6.5) buffer. The resulting solution was applied to a column (3 x 60 cm) of Bio-Gel A-l.5 m equilibrated with 25 mM Tris-maleate (pH 6.5) buffer. The column was eluted with the same buffer at a flow rate of 15 ml/h. Those fractions containing the greatest trehalase activity were pooled and after overnight dialysis (at 4°C) versus water were lyophilized. Prior to its usage in the assay of trehalose, the above powder was solubilized in 25 mM Tris-maleate (pH 6.5) buffer and the resultant solution clarified by centrifugation for 20 min at 33,OOOg. The final specific activity of this trehalase preparation was approximately loo-fold higher than that of crude extract enzyme and was usually 1000-2000 units/mg of protein (1 unit = 1 nanomole of glucose per minute at 35°C when enzymatic activity was assayed as previously described (16)). Protein assay. Protein was determined by the method of Lowry et al. (20) with crystalline bovine serum albumin as standard.

Temperature

tracts could be stabilized to low temperature (4°C) in pH 7.5 buffers. Of four buffers examined (i.e., MOPS-NaOH, TricineNaOH, Tris-HCl and potassium phosphate), the only one in which full activity was retained after 24 h at 4°C was potassium phosphate (50 mM). In several independent experiments, it was frequently found that enzyme incubated in phosphate buffer under these conditions (pH 7.5 at 4°C for 24 h) showed from lOO-115% of the original activity. With respect to the other buffers, enzymatic activity appeared to be lost somewhat more rapidly in TricineNaOH and Tris-HCl as opposed to MOPS-NaOH buffer. Because of the stability of trehalose-6-P synthetase in phosphate buffer at low temperature, experiments were conducted to determine the minimal phosphate concentration required for maximal recovery of activity under these conditions. In 50 mM MOPS-NaOH (pH 7.5) buffer, the minimal phosphate concentration which afforded maximal stability at 4°C was 10

4’

RESULTS

Low

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FROM

Instability

Since previous studies (15) had suggested that trehalose-6-P synthetase activity was cold labile at pH 6.5, the stability characteristics of this enzyme in crude extracts were examined at 4 and 23°C as a function of pH. As indicated in Fig. 1, stability as a function of pH was markedly different at these two temperatures. The pH for maximal stability, under the conditions described in the legend to Fig. 1, were 6.4-6.8 at 23°C and about 7.3-8.3 at 4°C. At pH 6.5, full activity was recovered after a 24-h incubation of the crude extract at 23°C. This was in contrast to the‘extract incubated at 4°C which retained approximately 35% of the original activity at this PH. Studies were conducted to determine whether synthetase activity in crude ex-

z’ -

60-

.I.

60

70

80

PH FIG. 1. Stability of trehalose-6-P synthetase as a function of pH and temperature. Extracts from cells at preculmination were prepared in 1 mM MOPS-NaOH (pH 7.5) buffer which contained 7.3 mM Na-thioglycollate. Aliquots (4 mg/ml protein) were incubated in 50 mM of each buffer, MESNaOH, 0; MOPS-NaOH, n ; Tricine-NaOH, X ; containing Na-thioglycollate (7.3 mr.r) for 24 h at either 4°C (- --) or 23” (-) under a drop of toluene. Assays of initial and final activity were carried out as indicated under Experimental Procedures.

KILLICK

638

AND WRIGHT

mM. Maximal stabilization of enzymatic activity appears to be specific for the phosphate anion, since potassium chloride (10 rn& did not affect stability, while both potassium nitrate and potassium sulfate at the same concentration appeared to increase instability at pH 7.5 (4°C). In contrast to the enzyme from silk moth fat body (211, trehalose-6-P synthetase from Dictyostelium was not stabilized by 10 mM Mg2+ (added as MgS03 under the above conditions of incubation. pH for Optimum Activity At 23”C, optimum activity was obtained from pH 6.5-7.0. Of several buffers tested, highest activity was obtained with MESNaOH and MOPS-NaOH (50 mM) buffers. At comparable concentrations, both phosphate and Tris buffers appeared to be inhibitory. Trehalose Requirement for Extract Preparation

TABLE I ACTIVITY OF TREHALOSE-6-P SYNTHETASE EXTRACTS

PREPARED OF

IN OR ABSENCE

IN THE PRESENCE TREHAUXIE”

Developmentalstage

Aggregation (9 h) Preculmination (16 h) Sorocarp (24 h)

Specificactivity Extract - trehalose

Extract + trehalose

12 31 23

16 40 40

D Extracts were prepared (see Experimental Procedures) from cells at three developmental stages in 50 mM MOPS-NaOH buffer (pH 7.51, which contained 10 mM potassium phosphate and 5 mre 2mercaptoethanol. When present, the final trehalose concentration was 25 and 2.5 mM in the harvesting buffer and assay mixture, respectively. Activity was assayed immediately after extract preparation as described under Experimental Procedures.

absence of 2.5 mM trehalose, the enzyme specific activity was 23 units/mg of protein in both cases. Enzyme prepared in the presence of trehalose (and, hence, subsequently assayed in a 2.5 mM of this sugar) had a specific activity of 40 unitslmg of protein. Therefore, trehalose does not appear to activate enzymatic activity during the assay, but presumably serves a critical role at the time of cell breakage and subsequent extract preparation.

Although trehalose-6-P synthetase activity was stable at 4°C in extracts prepared in 50 mM MOPS-NaOH (pH 7.5) buffer, which contained 10 mM phosphate plus 5 mM 2-mercaptoethanol, it was observed that the specific activity could be further increased by approximately 1.3-2.0-fold (depending on the developmental stage) by preparation of the extracts in the presence of trehalose (Table I). The minimal trehaKinetics and Stoichiometry lose concentration required for maximal specific activity was 25 mM. This observaThe kinetics and stoichiometry of prodtion is in agreement with that reported uct formation by trehalose-6-P synthetase previously (16) which demonstrated that were examined with crude, cell-free extrehalose elevated the specific activity of tracts prepared in MPMT buffer (see Exenzyme prepared at 23°C in pH 6.5 buffer perimental Procedures) at five stages of during the culmination process. differentiation (Fig. 2). The products were Since extracts prepared in the presence measured with a calorimetric assay (UDP) of trehalose were usually assayed for syn- and a radiometric assay (trehalose). For thetase activity in the presence of 2.5 mM maximal recovery of [‘4Cltrehalose during trehalose, it was conceivable that the pri- the latter assay, it was necessary to incubate the trehalose-6-P synthetase assay mary effect of this sugar was the activation of synthetase activity during the as- mixture (after termination of the synthesay, as opposed to enzyme stabilization at tase reaction by boiling) with bacterial althe time of extract preparation. In a con- kaline phosphatase, suggesting the limitatrol experiment, sorocarp extracts were tion of phosphatase activity during the synprepared in the presence and absence of 25 thetase assay. mM trehalose. When extracts lacking treFor each stage of development examhalose were assayed in the presence and ined, initial velocities were determined by

TREHALOSE3-P

SYNTHETASE

measuring the rates of synthesis of UDP and trehalose at lo-15min intervals over a 60-75min period of time. The amount of carrier trehalose present during the assay was either 2.5 or 5.0 mM, depending upon the amount of crude extract being assayed. Control experiments demonstrated that neither of these trehalose concentrations affected enzymatic activity. In addition, it was also demonstrated with extracts prepared from cells at approximately 4-24 h of development that the initial velocity was a linear function of the protein concentration being assayed. Initial velocities measured with the colorimetric assay were comparable to those determined with the radiometric assay when extracts prepared from cells between approximately the 5th or 6th to the 24th h I

I

I

1

.C

.b

./ /

0

20

40

I 60

I 80

MINUTES

FIG. 2. Kinetics of [Wltrehalose synthesis. Extracts were prepared in MPMT buffer at five stages of development and assayed for trehalose-6-P synthetase activity using the radiometric assay described under Experimental Procedures. The amount of protein used in each of the assays was (a) smooth myxamoebae (3.5 h), 270 pg; (b) rippled myxamoebae (5.0 h), 270 Fg; (c) early cell aggregates (7.5 h), 162 pg; (d) cells at preculmination (18 h), 155 pg; and (e) sorocarps (24 h), 220 fig. The specific radioactivity of UDP-[Wlglucose used as substrate varied with developmental stage as described under Experimental Procedures.

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of differentiation were used as the source of enzyme. Hence, only during this developmental period was it possible to demonstrate that UDP and trehalose were synthesized in equivalent amounts during the trehalose-6-P synthetase reaction. However, with extracts prepared prior to the rippled myxamoebae stage of differentiation (i.e., 5-6 h), no apparent correlation between the rates of UDP and trehalose production could be detected. Hence, it appears that, under the conditions used for the preparation of such extracts (see Experimental Procedures), the only valid method for measuring synthetase activity was with the radiometric assay. In the case of each of the stages studied between approximately the 4th and 24th h of development, the rate of trehalose formation was usually linear for at least the first 30 min of the assay (Fig. 2). For sorocarp extracts and those prepared prior to 5 h of differentiation, the rate of trehalose synthesis appeared to decrease during the remaining 20-30 min of the assay. Since with sorocarp extracts the rate of UDP synthesis was linear for at least 60 min under the same conditions, it appears that this decrease in amount of radioactive trehalose was probably due to the activities of both endogenous trehalase and alkaline phosphatase during the synthetase assay, as the specific activities of these two enzymes undergo dramatic increases during the culmination process (9, 16, 22). Although the specific activity of alkaline phosphatase is low in myxamoebae extracts (221, the level of trehalase (units per cell aliquot) is about 2.5 times that observed with sorocarp extracts (16). Since trehalase catalyzes not only the hydrolysis of trehalose but also apparently the conversion of trehalose-6-P to glucose and glucose-6-P (231, it is possible that the hydrolysis by this enzyme of trehalose and trehalose-6-P may at least in part be responsible for the decreased rate of [14C]trehalose production observed to occur during the latter half of the assay period. Developmental

Profile

When the specific activity of trehalose6P synthetase was examined in extracts pre-

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AND

pared in either MPMT or MMT buffer (as described under Experimental Procedures) comparable developmental profiles were obtained, suggesting equivalent stability under the two distinctly different conditions of extract preparation. Comparison between the UDP (colorimetric) and trehalose (radiometric) assays of synthetase activity (Table II) demonstrates that excellent agreement between the two assays is observed for enzyme prepared between the 5th or 6th (rippled myxamoebae) and 24th (sorocarp) h of development. The minimal specific activity observed was obtained for enzyme prepared from cells during the first 5-6 h of differentiation and usually amounted to 6-7 units/mg of protein. The maximal specific activity achieved during differentiation was 35-42 unitslmg of protein and occurred from the 17th to the 24th h of differentiation. During the first 5-6 h of development, enzyme specific activity appeared to be relatively constant. From about the 5th to the 17th h (preculmination) of differentiation, there was an approximate five- to sixfold increase in specific activity, which appeared to occur in a linear fashion. At subsequent stages of differentiation (i.e., 18-24 h), enzyme specific activity usually remained constant. During the course of TABLE DEVELOPMENTAL

Developmental

II

PROFILE OF TREHALOSE-6-P SYNTHETA~E" Specific activity

Stage

UDP assay Smooth myxamoebae (3.5 h) Rippled myxamoebae (5 h) Early cell aggregates (7.5 h) Preculmination (18 h) Sorocarp (24 h)

7.0 14 39 35

Trehalose assay 6.7 6.7 14 38 34

a Extracts were prepared in MPMT buffer and maintained at 4°C at each of live developmental stages. Trehalose-6-P synthetase activity was assayed (23°C) as described under Experimental Procedures. The specific activity is expressed as nanomoles of product per minute per milligram of protein.

WRIGHT

several studies on the developmental profile of trehalose-6-P synthetase, it was found that enzymatic activity on a unitsper-cell-aliquot basis increased approximately three- to fourfold during morphogenesis and was fairly constant from about the 10th (aggregation) to the 24th (sorocarp) h of differentiation (Table III). Changes in enzyme specific activity during this period of morphogenesis appear to be due to the net decrease in amount of buffer-soluble extract protein, which presumably reflects catabolism of endogenous protein during differentiation in Dictyostehum. DISCUSSION

Mechanistically, very little is known about the regulation in uiuo of trehalose synthesis. Since trehalose-6-P synthetase catalyzes the synthesis of trehaloseSB, the immediate precursor of trehalose, attention has been focused on the developmental behavior of this enzyme, as regulation of its activity in viuo might be expected to play a particularly prominent role in the control of both trehalose synthesis and accumulation during differentiation in Dictyostelium (24, 25, 26). The experimental bases for the discrepancies between the present analysis and those of previous investigators (9, 10, 11) concerning the developmental profile for trehalose-6-P synthetase appear to fall within three categories: 1) Culture conditions of organism, 2) Preparation and stabilization of enzyme, and 3) Assay methods. Culture conditions. Previous investigators (10) have grown D. discoideum NC-4 on nutrient agar in association with Aerobatter aerogenes. Morphogenesis was initiated by respreading the cells on black Millipore filters, supported by adsorbent pads saturated with a phosphate buffer/salts SOlution containing streptomycin sulfate. The present study employed E. coli as the bacterial associate and morphogenesis occurred on 2% nonnutrient agar, containing 10 mM phosphate buffer (pH 6.5) and 1 mM EDTA. Although it has been well documented (27-31) that the developmental profile for an enzyme can be influenced by various

TREHALOSE-6-P TABLE

SYNTHETASE

III

CONSTANCY OF TREHALOSE-6-P SYNTHETASE ACTIVITY FROM THE AGGREGATION TO THE SOROCARP STAGE OF DEVELOPMENT" EXperiment number

Stage

Trehalose6-P synthetase (units/ cell aliquot)

1

Aggregation (10 h) Pseudoplasmodium (14 h) Preculmination (18 h)

57 58 54

2

Preculmination (18 h) Late Culmination (22 h) Sorocarp (24 h)

54 52 57

’ At live stages of development, equal cell aliquots were prepared and assayed for synthetase activity der Experimental Procedures.

extracts from in MPMT buffer as described un-

environmental perturbations, little attention has apparently been paid to the effect (if any) of the bacterial associate on the detection of specific developmental enzymes in extracts from either vegetative myxamoebae or cells undergoing the early stages of morphogenesis. For this reason, it is interesting to compare the profile for trehalose-6-P synthetase from wild-type D. discoideum with that from axenic strain Ax-2 (32), which can be grown in the absence of bacteria. In extracts prepared from the latter organism, trehalose-6-P synthetase undergoes changes in specific activity during the first 18 h of development, which are comparable to those described for strain NC-4 in this paper (i.e., specific activity increases about five- or sixfold during morphogenesis and enzyme is detectable prior to 5 h of development). The kinetic constants obtained for enzyme from axenic D. discoideum (32) are in excellent agreement with those reported for the corresponding enzyme from wildtype strain NC-4 (10). The ease of detection of synthetase activity in myxamoebae extracts prepared from several axenic strains (331in addition to strain Ax-2, in contrast to the difficulty which has long been encountered with wild-type Dictyostelium,

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would not therefore appear to derive from a genetically induced modification of this enzyme, detectable solely by kinetic analysis. Hence, it may be that the low trehalose-6-P synthetase activity detected in extracts prepared during the first 5 h of development in strain NC-4 by investigators in the past (9-11, 15) was caused (at least in part) by a metabolic product(s) originating from the bacterial associate, which effectively masked synthetase activity, under the conditions which were used for the preparation and assay of this enzyme. Support for such a speculation derives from the studies of Killick and Wright (15) which showed that under certain conditions of enzyme preparation and assay, trehalose-6-P synthetase activity is masked in myxamoebae extracts. Although the mechanism of this in vitro latency is unknown, studies by Roth and Sussman (9) may be interpreted as suggesting the involvement of an endogenous inhibitor. When the latter investigators mixed extracts from 5-h cells with those from preculmination, the activity obtained indicated a 56% inhibition. In the presence of carrier trehalose, inhibition was reduced to 39%. As the above observations suggest a basis for the in vitro latency of trehalose-6P synthetase activity in extracts prepared during the initial stages of differentiation, it is conceivable that the experimental conditions described in the present paper may either prevent or reverse suppression of synthetase activity caused by the endogenous component in question. Preparation and stabilization of enzyme. In previous studies (9-11) enzyme has been prepared from nonwashed cells throughout development by sonication in 10 mM Tris-HCl (pH 7.5) buffer containing sodium thioglycollate (0.5 mg/ml). The resulting homogenate was used directly as the source of enzyme. In the present study, extracts have been prepared in MPMT buffer from washed cells by either a single freeze-thaw (up to and including 18 h of development) or a freeze-thaw, followed by a single passage through a French pres-. sure apparatus (20-24 h of development). The resulting homogenate was centrifuged twice (33,OOOg)prior to the assay of enzymatic activity.

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KILLICK

AND

The specific activity observed for trehalose-6-P synthetase during development is markedly affected by the methods used for cell rupture. In a study of the effects of different methods of cell breakage on trehalose-6-P synthetase activity at preculmination, Roth (11) demonstrated that maximal activity was obtained with extracts which had been prepared by a single freeze-thaw, while use of the French pressure apparatus reduced activity by 71%. At the same stage of development, sonication apparently was not detrimental to enzymatic activity. A major drawback to the use of sonication, however, is that this procedure does not effectively rupture cells during the later stages of development (i.e., 20-25 h) (11). Hence, the decline in specific activity of trehalose-6-P synthetase, which Roth and Sussman (10) observed between the 18th and 24th h of development may have been due in part to inefficient cell breakage. In the present study, cells were effectively broken during the terminal stages of development with the French pressure apparatus. No loss of activity was detected as a result of this operation, due presumably to the stabilizing effect of trehalose present in the buffer (MPMT buffer) used for preparation of the homogenate. In addition to apparently suboptimal conditions of enzyme preparation, two additional factors, both associated with enzyme stabilization, may be responsible for the low trehalose-6-P synthetase specific activities previously observed (9-11). These are 1) the lability of trehalose-6-P synthetase activity at 4°C in pH 7.5 buffer lacking phosphate and 2) the requirement for trehalose for optimal stabilization of enzymatic activity in extracts prepared throughout development (Table I). Methods of assay. In assaying trehalose6-P synthetase activity during differentiation in Dictyostelium, previous investigators (10, 11) have employed 6 mM Tris-HCl buffer (pH 6.6) at 37”C, whereas in the present paper, 50 mM MES-NaOH buffer (pH 6.5) at 23°C has been used. Tris buffers have been reported to inhibit trehalose-6-P synthetase activity (8, 11); another reason for avoiding their use derives from their

WRIGHT

greater variability in pH as a function of temperature as compared to any of the buffers of Good et al. (34). Moreover, measurements of enzymatic activity at 37°C (10) may not be physiologically meaningful (26) as Dictyostelium does not undergo differentiation at this temperature. Because of the pronounced changes in specific activity of trehalase which occur during development in Dictyostelium (9, 16), it is essential when employing the radiometric assay of trehalose-6-P synthetase activity to follow the kinetics of [14C]trehalose synthesis over short intervals of time (i.e., 10 min) in the presence of a concentration of carrier trehalose not inhibitory (either directly or indirectly) to synthetase activity (Fig. 2). In previous studies, investigators have followed the rates of [14C]trehalose synthesis at hourly intervals; in several cases this did not represent the initial velocity (9). Either they did not use carrier or employed concentrations of trehalose (40 mM) which may inhibit synthetase activity, depending upon the developmental stage at which the extract is prepared (9, 10). Hence, erroneous conclusions based on a false kinetic characterization of [14Cltrehalose production at several developmental stages may have been obtained. On a methodological basis, it would appear from the above discussion that several of the discrepancies concerning the developmental profile of trehalose-6-P synthetase have been resolved. As the present studies have demonstrated that synthetase activity on a per-cell-aliquot basis is constant from approximately the 10th h (aggregation) to the 24th h (sorocarp) of development, it is unlikely that a sudden rise in overt trehalose-6-P synthetase activity, resulting from an induction of enzyme synthesis, as suggested by previous investigators (lo), is the rate-limiting event (i.e., primary critical variable) in the initiation of trehalose synthesis in vivo during the culmination process (i.e., 20-22 h). Since the maximum specific activity obtained for trehalose-6-P synthetase in extracts prepared at preculmination is less than that theoretical value required in vivo to achieve the maximum rate of tre-

TREHALOSES-P

SYNTHETASE

halose synthesis (13), it is concluded that neither the developmental profile nor the maximum specific activity obtained are consistent with either the timing or the extent of trehalose accumulation during development. REFERENCES 1. BONNER, J. T. (1971) Annu. Rev. Microbial. 25, 75-92. 2. WHITE, G. J., AND SUSSMAN, M. (1963) Biochim. Biophys. Acta 74, 173-178. 3. WHITE, G. J., AND SUSSMAN, M. (1963) Biochim. Biophys. Acta 74, 179-187. 4. CECCARINI, C., AND FIL~SA, M. (1965) J. Cell. Comp. Physiol. 66, 135-140. 5. ROSNESS, P. A., AND WRIGHT, B. E. (1974) Arch. Biochem. Biophys. 164,60-72. 6. CECCARINI, C. (1967) Biochim. Biophys. Acta 148, 114-124. 7. COTTER, D. A., ANDRAPER, K. B. (1970)Develop. Biol. 22, 112-128. 8. CABIB, E., AND LELOIR, L. F. (1958) J. Biol. Chem. 231, 259-275. 9. ROTH, R., AND SUSSMAN, M. (1966)Biochim. Biophys. Acta 122,225-231. 10. ROTH, R., AND SUSSMAN, M. (1968) J. Biol. Chem. 243,5081-5087. 11. ROTH, R. (1967) Ph.D. thesis, 107 pp., Brandeis University, Waltham, MA. 12. ROTH, R., ASHWORTH, J. M., ANQ SU~~MAN, M. (1968) Proc. Nat. Acad. Sci. USA 59, 12351242. 13. WRIGHT, B. E., AND MARSHALL, R. (1971)5. Biol. Chem. 246,5335-5339. 14. SARGENT, D., AND WRIGHT, B. E. (1971) J. Biol. Chem. 246, 5340-5344. 15. KILLICK, K. A., AND WRIGHT, B. E. (1972) J. Biol. Chem. 247, 2967-2969. 16. KILLICK, K. A., AND WRIGHT, B. E. (1972) Biochem. Biophys. Res. Commun. 48, 14761481.

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17. LIDDEL, G. U., AND WRIGHT, B. E. (1961) Develop. Biol. 3, 265-276. 18. PONTIS, H. G., AND LEU)IR, L. F. (1962) Methods Biochem. Anal. 10, 107-136. 19. ELBEIN, A. D. (1967) J. Biol. Chem. 242, 403406. 20. LOWRY, 0. H., ROSEBROUGH, N. J., FARR, A. L., AND RANDALL, R. J. (1951) J. Biol. Chem. 193, 265-275. 21. MURPHY, T. A., AND WYATT, G. R. (1965)5. Biol. Chem. 240, 1500-1508. 22. GEZELIUS, K., AND WRIGHT, B. E. (1965) J. Gen. Microbial. 38, 309-327. 23. DIXON, M., AND WEBB, E. C. (1964) Enzymes, 2nd ed., p. 747, Longmans Green, London. 24. WRIGHT, B. E. (1973) Critical Variables in Differentiation, Prentice-Hall, Englewood Cliffs, NJ. 25. GUSTAFSON, G. L. AND WRIGHT, B. E. (1972) in Critical Reviews in Microbiology (La&in, A. I., and Lechevalier, H., eds.), Vol. 1, pp. 453478, CRC, Cleveland, OH. 26. KILLICK, K. A., AND WRIGHT, B. E. (1974)Annu. Rev. Microbial. 28, 139-166. 27. NEWELL, P. C., AND SUSSMAN, M. (1970) J. Mol. Biol. 49, 627-637. 28. NEWELL, P. C., LONGLANDS, M., AND SUSSMAN, M. (1971) J. Mol. Biol. 58, 541-554. 29. ELLINGSON, J. S., TELSER, A., AND SUSSMAN, M. (1971) Biochim. Biophys. Acta 244,388-395. 30. NEWELL, P. C., FRANKE, J., AND SUSSMAN, M. (1972) J. Mol. Biol. 63, 373-382. 31. MCMAHON, D., AND FORGAC, M. (1974) Fed. Proc. 34, 1476. 32. GARROD, D., AND ASHWORTH, J. M. (1973) in Microbial Differentiation, (Ashworth, J. M., and Smith, J. E., eds.), pp. 407-435, 23rd Symp. Sot. Gen. Microbial. Cambridge University Press, London. 33. WASHINGTON, A. (1971) Ph.D. thesis, 76 pp., Illinois Institute of Technology, Chicago, Ill. 34. GOOD, N. E., et al. (1966) Biochemistry 5, 467477.