Enzyme-inhibitor interactions as a basis for heterogeneity of extracellular cyclic AMP phosphodiesterases in Dictyostelium discoideum

Biochimica et Biophysica Acta, 802 (1984) 413-422

413

Elsevier

BBA 21934

E N Z Y M E - I N H I B I T O R INTERACTIONS AS A BASIS FOR H E T E R O G E N E I T Y OF EXTRACELLULAR CYCLIC AMP P H O S P H O D I E S T E R A S E S IN D I C T Y O S T E L I U M DISCOIDEUM D A V I D P. T O O R C H E N * and ELLEN J. H E N D E R S O N **

Department of Chemisto:, Massachusetts Institute of Technology, Cambridge, MA 02139 (U.S.A.) (Received June 25th, 1984)

Key words: cyclic AMP," Enzyme inhibitor protein; Phosphodiesterase," (D. discoideum)

We have previously characterized three forms of cyclic-AMP phosphodiesterase obtained after dithiothreitol activation of the enzyme from the extracellular medium during late vegetative growth of Dictyostelium discoideum (Toorchen, D. and Henderson, E.J. (1979) Biochem. Biophys. Res. Commun. 87, 1168-1175). This communication presents evidence supporting the earlier hypothesis that the observed heterogeneity of enzyme species is due to formation of complexes between an endogenous inhibitor protein and a common catalytic polypeptide. Dithiothreitol inactivates the inhibitor, but does not cause its release from the catalytic unit. Additional evidence is presented for the presence of a similar catalytic polypeptide in the extracellular phosphodiesterase produced during the first 8 h of development, except that this species is a phosphoprotein.

Introduction Amoebae of Dictyostelium discoideum undergo a developmental process triggered by nutrient deprivation. Early in this process the amoebae release pulses of cAMP into the medium. These pulses induce chemotactic responses, mediated by cell surface receptors for the signalling agent, cAMP, such that the amoebae then aggregate into multicellular masses (reviewed in Ref. 1). The amoebae also elaborate cyclic-AMP phosphodiesterases (cyclic-adenosine 3': 5'-monophosphate 5'nucleotidohydrolase, EC 3.1.4.17) for maintenance

* Current address: Department of Pathology, School of Medicine, The University of North Carolina at Chapel Hill, Chapel Hill, NC 27514, U.S.A. ** To whom correspondence should be addressed at (current address): Department of Biology, Georgetown University, Washington, DC 20057, U.S.A. Abbreviations: PMSF, phenylmethylsulfonyl fluoride; SDS, sodium dodecyl sulfate; Mes, 4-morpholineethanesulfonic acid. 0304-4165/84/$03.00 © 1984 Elsevier Science Publishers B.V.

of a proper signal-to-noise ratio. Extracellular phosphodiesterase activity is found during vegetative growth, but is dramatically reduced in early starvation by production of a heat-stable inhibition protein (reviewed in Ref. 2). cAMP pulses suppress inhibitor production and induce the appearance of new extracellular as well as membrane-bound phosphodiesterase activity [3-5]. The first report of D. discoideum extracellular phosphodiesterase by Chang [6] described an enzyme with a molecular weight of 2 - l0 s and a high Michaelis constant (2 mM) for cAMP. Pannbacker and Bravard [7] subsequently showed that freshly secreted enzyme had a higher affinity for cyclic AMP ( K , , = 15 ~M) and could therefore be a significant regulator of in vivo cAMP concentrations [8]. This enzyme had a molecular weight of 65 000 [7,9]. Chassy [9] showed that treatment of phosphodiesterase from the extracellular medium with dithiothreitol resulted in large increases in enzyme activity and that the activity gave a very heteroge-

414

neous profile on ion-exchange columns [10]. The activities were subsequently resolved into three species of differing molecular weights and isoelectric points, the larger forms being progressively more acidic [11 13]. It was shown that the polypeptide inhibitor forms a stable complex with the enzyme [12,14-16], which converts the low Kn, of the M r 65000 phosphodiesterase to mM ranges (15) and that dithiothreitol causes an inactivation of the inhibitor protein [11,15]. Since the phosphodiesterase inhibitor is acidic [2] and the M r 65 000 phosphodiesterase basic [12,13], the heterogeniety of the various phosphodiesterase species observed could be explained by a model in which of a single phosphodiesterase catalytic polypeptide forms high K,,, enzyme-inhibitor complexes of differing stoichiometries and according to which dithiothreitol activates the enzyme (reduces the K,,) by inactivation of the inhibitor without causing its release from the complex. It has generally been assumed that inactivation of the inhibitor by dithiothreitol causes its release from extracellular phosphodiesterase. However, we report here that the larger phosphodiesterase species contain a polypeptide of the same size as the inhibitor, that the inhibitor can generate all of the extracellular phosphodiesterase species in mixtures and that dithiothreitol does not block or reverse this process even though it eliminates the inhibitory activity. Methods

Strain and culture conditions. All experiments were performed with axenic strain Ax3 grown on autoclaved suspensions of Escherichia coil B / r to late exponential or early stationary phase for preparation of vegetative extracellular phosphodiesterase. For studies using the inhibitor protein or developmental phosphodiesterase, the cells from cultures as above were harvested by centrifugation at ( 5 0 0 - 7 0 0 ) x g and were washed once with and resuspended at ( 1 - 2 ) × 107/ml in 20 mM potassium phosphate (pH 6,1)/0.5 mM CaC12. These starvation cultures were shaken at 200 rpm, 22°C, for 15 18 h. Supernatants from vegetative or starvation cultures were obtained following centrifugation at 500-700 × g for 15 rain. Preparation of vegetative extracellular phos-

phodiesterase forms. The procedure involves two a m m o n i u m sulfate precipitations with an intervening activation of phosphodiesterase by dithiothreitol and subsequent fractionation on DEAE cellulose. Details are in the report of Chassy and Porter [10] as modified by Toorchen and Henderson [11]. This procedure resolves three phosphodiesterase species with molecular weights on Sephacryl S-200 columns of M r 65000 (phosphodiesterase 65), 95000 (phosphodiesterase 95) and 185 000 (phosphodiesterase 185). Assay of cAMP phosphodiesterase activity. The assay was as previously described [11] and is based on spectrophotometric determination of inosine formed in a coupled assay with 50 ffM cAMP, phosphodiesterase, alkaline phosphatase and adenosine deaminase at 30°C (pH 7.5). Units are expressed as nmol 5'-AMP formed/rain. Since the concentration of dithiothreitol required to activate the phosphodiesterase also causes strong kinetic inhibition [7], the enzyme was always diluted to minimally inhibitory concentrations before assay, i.e., < 0.1 mM [17]. Assay of inhibitor protein. This was as previously described [11]. 1 unit of inhibitor is defined as the amount which reduces 2 units of phosphodiesterase activity to 1 unit. This definition is somewhat arbitrary in that the affinity of the inhibitor is not the same for all phosphodiesterase species [11]. In all experiments reported here the inhibitor was assayed against a single phosphodiesterase preparation obtained after the second (NH4)2SO 4 precipitation.

Preparation of crude inhibitor protein and developmental extracellular phosphodiesterase. Supernatant from starved amoebae (approx. 350 ml) was concentrated 5-fold by ultrafiltration (Amicon stirred-concentration cell and a PM 10 membrane). The concentrate was clarified by centrifugation at 20000 × g for 45 min. If inhibitor was to be prepared, the solution was adjusted to 0.5 M KC1 by the addition of 5 M KC1, placed in a water bath at 80°C for 15 20 min and then transferred to an ice-bath, The chilled material was clarified by centrifugation at 20000 x g for 1 h and dialyzed against 21 of 50 mM Tris-HC1 (pH 7.5 at 5°C), with one change of dialysis buffer. This treatment resulted in 6-fold purification and a 3-fold increase in total inhibitor activity, presumably re-

415 flecting its release from thermally-inactivated phosphodiesterase. The resulting solution is referred to as heat-treated starvation supernatant. If developmental extracellular phosphodiesterase activity was required, the addition of KCI and the heat-step were omitted. Instead, the solution was dialyzed against 50 mM Tris-HC1 as described above, and the resulting material was made 5 mM in dithiothreitol by addition of the solid reagent. This solution was incubated at 22°C for 1 h, clarified by centrifugation at 20000 × g for 30 min, and then dialyzed against 2 l of 50 mM Tris-HC1 (pH 7.5 at 5°C), containing 0.2 mM dithiothreitol, with one change of buffer. Purification of inhibitor protein. Columns of DEAE-cellulose (Whatman DE-52, 1.6 x 50 cm) were equilibrated with 10 mM Tris-HC1 (pH 7.5 at 5°C). A 30 ml aliquot of heat-treated starvation supernatant in the same buffer was applied at a flow rate of 1 ml/min, and 5-ml fractions were collected. After washing the column with 100 ml of the loading buffer, elution was with a linear 500 ml gradient from 10 mM Tris-HC1 (pH 7.5 at 5°C), to 50 mM Tris-HC1 (pH 7.5 at 5°C), 0.5 M KCI. Inhibitor activity eluted between 0.1-0.2 M KC1. Fractions containing activity were pooled, concentrated by ultrafiltration and dialyzed against 0.1 M Tris-HC1 (pH 7.5 at 5°C). The concentrated preparation was further fractionated on a calibrated Sephacryl S-200 column as described [11], except omitting dithiothreitol from the buffer• Total purification was 146-fold.

gels was as described by Davis [18] with the LKB Instruments modification for use with the Multiphor flat-bed apparatus. Activity staining for phosphodiesterase was performed as described by Goren et al. [19]. For visualization of the high-K,, phosphodiesterase-inhibitor complex, the cAMP concentration was raised to 2 mM, twice that described in the cited procedure.

Protein standards and molecular weights were phosphorylase a (9.4. ]04), bovine serum albumin (6.8 • 104), IgG subunit (5.0. 104), ovalbumin (4.3 • 104), carbonic anhydrase (2.8. 104), IgG subunit (2.3 • 104) and cytochrome c (1.7 - 104). A mixture of these proteins was radioiodinated as described below for detection by fluorography or autoradiography. Radioiodination of proteins• Soluble protein samples were iodinated using 1,3,4,6-tetrachloro3a,6a-diphenylglycouril (Iodogen, Pierce Chemical Co.) as described by Markwell and Fox [21]. Typically, 100 /~g of protein in 50 mM Tris-HC1 (pH 7.5 at 5°C), or in Tris-glycine buffer (20 mM glycine, adjusted to pH 8.9 at 5°C with Tris) was placed in a test tube plated with 10 /~g of the iodinating reagent and 500 /~Ci of 12sI (carrierand reductant-free, 100 mCi/ml) added to initiate the reaction. If retention of inhibitor or phosphodiesterase activity was desired, the reaction was conducted on ice; in other cases it was conducted at room temperature. The reaction proceeded for 30-60 min and was quenched by addition of 2.5 M NaI to a final concentration of 0.25 M. Free 12sI was separated from protein-bound radioactivity either by extensive dialysis at 5°C or by gel filtration. In this latter method a 10 ml. Sephadex G-25 column was equilibrated with 0.1 M ammonium bicarbonate and the sample chromatographed at room temperature. Fractions of 1 ml were collected, and the first two radioactive fractions were saved for analysis. Autoradiography and fluorography. Kodak XOmat film was used. Depending upon the experiment, gels were either fixed and dried onto filter paper for exposure or were wrapped in clear plastic (Saran Wrap) and exposed to film, while the gel was still hydrated. Fluorography was performed as described by Laskey and Mills [22] on gels of 125I-labeled proteins with a low input radioactivity.

Sodium dodecyl sulfate-polyacrylamide gel electrophoresis. Electrophoresis under denaturing

Coupling of heat-treated starvation supernatant to CNBr-activated Sepharose. Heat-treated starva-

conditions was performed as described by Laemmli [20]. For autoradiography or fluorography, gels were fixed after electrophoresis in 10 vol. of 25% isopropanol, 10% acetic acid and then dried under vacuum onto Whatman 3MM chromatography paper.

tion supernatant (5 ml, 2 mg/ml, in 0.1 M NaHCO3/0.5 M NaCI (pH 8)) was mixed with 40 mM PMSF in 95% ethanol to give a final concentration of 2 mM. After 45 min on ice, centrifugation (20000 x g, 30 min) removed the precipi-

Nondenaturing polyacrylamide gel electrophoresis. Electrophoresis in 3.5% polyacrylamide slab

416

tate which had formed. The supernatant proteins were then coupled to cyanogen bromide-Sepharose 4B (Pharmacia) and washed according to the manufacturer's instruction. The final matrix was washed several times with 50 mM Tris-HC1, 0.02% sodium azide (pH 7.5 at 5°C), and stored until needed at 5°C. Affinity chromatography. In order to purify extracellular phosphodiesterase fractions on heattreated starvation supernatant-Sepharose 4B, the factions were added to portions of a 1 : 1 slurry of matrix and buffer in 1.5 ml microcentrifuge tubes. These were incubated for 30 min at 30°C with frequent agitation. The tubes were then centrifuged, and the supernatant was removed by aspiration. The matrix was then washed 3-5-times with 5 vol. of 0.3 M D-galactose/50 mM Tris-HCl (pH 7.5) by resuspension and centrifugation. To release material specifically bound by the heattreated starvation supernatant-Sepharose, 1 vol. of 2% SDS was added to the matrix, and the tubes were mixed and heated at 100°C for 2 min. Following centrifugation and retention of the supernatant, this step was repeated. The supernatants were combined, lyophilized, and resuspended in Tris-glycine buffer for SDS-polyacrylamide gel electrophoresis. Protein cross-linking. For protein cross-linking [23] with dimethylsuberimidate (Pierce Chemical Co.), [125I]-labeled phosphodiesterase and 125Ilabeled inhibitor were dialyzed against 5 mM sodium phosphate buffer (pH 7.0). 20 ~1 of 125Ilabeled inhibitor (0.8 /~g protein, 2.5.105 cpm) were added to 20 /~1 of 125I-labeled phosphodiesterase 65 or the same volume of 5 mM phosphate buffer. The mixture was incubated at 30°C for 30 min at which time 5~1 of 1.5 M Tris-HC1 (pH 8.5) and 5 ~1 of dimethylsuberimidate/0.15 M Tris-HCl (pH 8.5) was added, and the reaction was allowed to proceed for 3 h at room temperature. The dimethylsuberimidate in the reaction was varied from 1 to 5 mg/ml. At the end of a 3-h period the samples were subjected to SDS-polyacrylamide gel electrophoresis. ~:P-labeling of developing amoebae. Amoebae were grown in HL-5 axenic media [24] and harvested by centrifugation at 700 × g for 10 rain. They were washed once with cold Mes buffer (per liter: 1.5 g Mes, 1.5 g KC1, 0.6 g MgSO 4 (pH 6.5

with KOH)). The washed amoebae were distributed on a Nuclepore filter (47 mm diameter, 0.2 /~m pore diameter, 2- 108 amoebae per filter) resting on an agar plate (3% agar in Mes buffer) and starved for 0, 4 or 8 h. At the appropriate times the filter was transferred to a drop of carrier free [32p]orthophosphoric acid (500 #Ci) in an otherwise empty petri dish, which was then covered with wet filter papers. After 60 min at 22°C the amoebae were harvested, washed once with 17 mM potassium phosphate (pH 6.1), and the supernatants were lyophilized preparatory to purification on inhibitor-Sepharose. Sephacrvl-S200 gel filtration. This was as previously described [11]. Protein determination. Protein was determined by the method of Bradford [25] (using Bio-Rad protein reagent and gammaglobulin fractions as standard) or spectroscopically by the method of Christian and Warburg [26]. Results

We have previously reported [11] fractionation of phosphodiesterase from late exponential or early stationary phase cultures into three major species with molecular weights of 65 000 (phosphodiesterase 65), 95 000 (phosphodiesterase 95) and 185 000 (phosphodiesterase 185) on Sephacryl S-200. The negative charge-to-mass ratio of these species increased progressively with size by the criteria of affinity for DEAE cellulose and mobility in nondenaturing polyacrylamide gel electrophoresis. In order to determine the polypeptide composition of these species and to test extracellular phosphodiesterase-inhibitor interactions, each form and the phosphodiesterase inhibitor had to be at least partially purified. Since extracellular phosphodiesterase species and the inhibitor do not stain well on gels with Coomassie blue (Ref. 16, and our unpublished observations), we used radioiodination or activity staining for detection on gels. The phosphodiesterase fractions and inhibitor could be radioiodinated with Iodogen, typically with 50 and 75% retention of activity, respectively.

Polypeptide composition of enzyme and inhibitor preparations The polypeptide composition of the various ex-

417

tracellular phosphodiesterase preparations and of the purified inhibitor was determined by SDSpolyacrylamide gel electrophoresis. Fig. 1A shows the phosphodiesterase preparations. The phosphodiesterase 65 from Sephacryl-S200 (lane 2) shows a highly purified band referred to as p65. While preparations of phosphodiesterase 95 (lane 3) and 185 (lane4) from DEAE cellulose were crude, both contain a p65 band. Also, both contain p38 and p95 bands, which are not detectable in phosphodiesterase 65. Fig. 1B (lanes 3 and 4) shows the purified inhibitor which gives a band of 38000 (p38). The major contaminant (lane 3) is a band at 63000 (p63), which is not detected in samples of vegetative stage phosphodiesterase 65, 95 and 185. By several criteria, considered later, p63 may be the phosphodiesterase produced during early development. l:SI-labeled crude phosphodiesterase 185 from DEAE cellulose was further purified by nondenaturing electrophoresis and localized by a stain for phosphodiesterase activity. The band of lead phosphate precipitate was excised from the gel and

A

B 95 68 50

eluted into Tris-glycine buffer containing 5% flmercaptoethanol. The eluted material on SDSpolyacrylamide gel electrophoresis gave the autoradiogram of Fig. 1B, lane 5. While recovery from the lead phosphate is poor, there was a selective enrichment of p38, 65 and 95. The presence of p95 band in both phosphodiesterase 95 and 185 suggested that it could be a stable complex of p65 with p38. The following experiments support, but do not prove, this possibility. First, crude inhibitor was covalently coupled to cyanogen bromide-activated Sepharose for use as an affinity matrix. After adsorption of radioiodinated phosphodiesterase fractions from DEAE and extensive washing with galactose-containing buffers, bound protein was eluted by boiling in SDS and analyzed by SDS-polyacrylamide gel electrophoresis. Both phosphodiesterase 95 and 185 generated an enriched p95 band (not shown). Second, we attempted to generate a p95 from mixtures of purified inhibitor and phosphodiesterase 65 in the presence of the bifunctional, covalent, protein crosslinker, dimethylsuberimidate, to trap the species for gel-electrophoretic analysis. The results are shown in Fig. 2. In the absence of the crosslinker (lane 2) only a trace of p95 was observed. In the presence of the crosslinker, p95 was substantially enhanced (lanes 3-5). The same figure shows crosslinker-dependent enhancement of p95 in mixtures of phosphodiesterase 185 and inhibitor (lane 6 vs. 7-9).

43

37

Inhibitor effects on electrophoretic mobility of native extracellular phosphodiesterase fractions

29

24 17

LANE

I

2

3

4

LANE

1

2

3

4

5

Fig 1. Polypeptide composition of purified phosphodiesterase and inhibitor fractions. The phosphodiesterase and inhibitor preparations were radioiodinated and fractionated by SDS-polyacrylamide gel electrophoresis. The autoradiogram lanes are as follows. (A) 1, molecular weight standards; 2, phosphodiesterase 65 after the Sephacryl S-200 step; 3 and 4, phosphodiesterase 95 and 185, respectively, from DEAE cellulose. (B) 1, molecular weight standards; 2, phosphodiesterase 185 from DEAE cellulose; 3 and 4, two loads of inhibitor; 5, the phosphodiesterase 185 used for lane 2 after elution from a nondenaturing gel.

If native phosphodiesterase 95 and 185 are formed by complexes of p65 and p38, then addition of the purified inhibitor to the lower molecular weight, less acidic forms should convert them to the larger more acidic species. Further, according to the model, treatment with dithiothreitol should activate the enzymes, but should not regenerate the input enzyme species if the inhibitor remains bound. These predictions are supported by the data in Fig. 3, which shows activity stains in nondenaturing polyacrylamide gels for various enzyme fractions with and without addition of inhibitor and with and without dithiothreitol treatment. Lanes la, 2a and 3a show the electrophoretic

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95 68 50 43 37 29 17 LANE

I

2

5

4

5

6

7

8

9

Fig. 2. Covalent crosslinking of inhibitor with phosphodiesterase 65 and 185. Dimethylsuberimidate was used to induce covalent crosslinking in protein complexes, and the products were analyzed by SDS-polyacrylamide gel electrophoresis. The lanes of the autoradiogram are: 1, molecular weight standards; 2, phosphodiesterase 65 plus inhibitor; 3 5, phosphodiesterase 65 plus inhibitor plus 1, 3 and 5 m g / m l dimethylsuberimidate, respectively; 6, phosphodiesterase 185 plus inhibitor: 7-9, phosphodiesterase 185 plus inhibitor plus 1, 3 and 5 m g / m l dimethylsuberimidate, respectively.

LANE

4 d 4 c 4 6 4 a 3 d 3 c 3 b 3a2d 2 c 2 b 2 a l d lc lb la

Fig. 3. Effects of the purified inhibitor on electrophoretic mobilities of native phosphodiesterase species. The phosphodiesterase species from DEAE-cellulose columns were incubated with or without the purified inhibitor, with or without prior dithiothreitol treatment of the inhibitor and with or without dithiothreitol treatment of the enzyme-inhibitor complexes. The lanes are: l a - d , phosphodiesterase 65; 2a-d, phosphodiesterase 95; 3 a - d , phosphodiesterase 185; 4 a - d developmental phosphodiesterase. Lanes a are the native enzyme species. Lanes b are after preincubation with active inhibitor. Lanes c are the result of dithiothreitol treatment of the preincubated mixtures of enzyme and inhibitor. Lanes d are the result of mixing the inhibitor after dithiothreitol treatment with the various enzyme species.

mobilities of phosphodiesterase 65, 95 and 185 prepared from DEAE cellulose columns. All the bands are diffuse, suggesting microheterogeneity within the species. When these species were incubated with purified inhibitor prior to electrophoresis, two results occurred. First, the activities were inhibited, since the stain intensity was considerably reduced (lanes lb, 2b and 3b). Second, as seen in the same lanes, the reduced activity of phosphodiesterase 65 migrated further into the gel as an apparent mixture of phosphodiesterase 95 and 185 (lane lb). The broad activity band of partially purified phosphodiesterase 95 (lane 2a) is compatible with its content of some phosphodiesterase 65 on gel filtration columns [11]. The inhibitor caused the disappearance of species migrating as phosphodiesterase 65 and generated a new band with the mobility of phosphodiesterase 185 (lane 2b). In the presence of the inhibitor, phosphodiesterase 185 formed a more tight band of inhibited activity at the higher mobility range of untreated phosphodiesterase 185 (lane 3b). These results show that the inhibitor can generate, progressively, all of the higher mobility species from the lower mobility species. The effects of the inhibitor on mobility are most pronounced on phosphodiesterase 65, which has no endogenous inhibitor, and are less dramatic on species postulated to contain endogenous but inactive inhibitor. The observation that the activity of all species is reduced suggests that active inhibitor can at least partially replace complexed, inactive inhibitor. Addition of dithiothreitol to the enzyme plus inhibitor mixtures prior to electrophoresis (lanes lc, 2c and 3c) restored enzymic activity to uninhibited levels, but did not prevent the inhibitor-induced mobility shifts. Similarly, treatment of the inhibitor with inactivating concentrations of dithiothreitol prior to its addition to the enzyme species did not prevent the inhibitor-induced mobility changes but did prevent inhibition of enzymic activity (lanes ld, 2d and 3d). Similar experiments were performed with crude, unfractionated, dithiothreitol-activated developmental extracellular phosphodiesterase, which migrated as a diffuse doublet close to phosphodiesterase 185 (lane 4a). The activity was sensitive to the inhibitor which also caused the doublet to collapse into a single band of about phos-

419 phodiesterase 185 mobility (lane 4b). The same effect on mobilities was observed when the inhibitor was inactivated by dithiothreitol after (lane 4c) or before (lane 4d) mixing with the enzyme.

Are vegetative and developmental catalytic subunits related? A small amount of developmental extracellular phosphodiesterase survives the heat inactivation step during the isolation of the inhibitor (not shown), and the major contaminant of the p38 inhibitor is a p63 polypeptide which is not detectable in any of the vegetative phosphodiesterase forms (Fig. 1). In control experiments to test possible use of disulfide-containing protein crosslinkers, we investigated the effects of/3-mercaptoethanol on polypeptide mobility during SDS-polyacrylamide gel electrophoresis. Both p65 and the p63 in the inhibitor preparation showed a marked change in apparent molecular weight in the absence of fl-mercaptoethanol (Fig. 4), migrating to apparent molecular weights of 13 000 less than the

p63 and 65 values observed in the other gels. The derivation of the smaller from the larger species is made clear by the transition in mobilities of these polypeptides in the gel lanes between those with and without /3-mercaptoethanol. We assume that these transitional lanes are the result of diffusion of the reducing agent from the sample wells of the gel slab. The simplest interpretation of these mobility shifts is that these proteins contain disulfide bonds which, in the absence of a reducing agent, stabilize a highly compact structure. The inhibitor p38, which is known to contain many half-cystine residues [16], also shifts to lower molecular weight in the absence of the reducing agent. The mobility of p95 is unaffected though its amount diminishes in the presence of fl-mercaptoethanol.

Developmental phosphodiesterase is a phosphoprotein Amoebae were given 1-h pulses of [3Zp]orthophosphate at three stages of early development. Extracellular proteins were harvested as described in Methods and fractionated on a crude inhibitorSepharose matrix. The material which could only

95 68

95

5O

68 5O

43 37

43 37

29 17

24 17

LANE

1

2

3

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7

Fig. 4. Effect of fl-mercaptoethanol on electrophoretic mobilities of p65, p63 and p38. The autoradiogram shows the electrophoretic mobilities of radioiodinated polypeptides when flmercaptoethanol had been either omitted (lanes 2-5) or included (lanes 1 and 6) in the sample preparation prior to electrophoresis by SDS-polyacrylamidegel electrophoresis. The samples are: lanes 1-3, purified inhibitor; lanes 4-6, phosphodiesterase 65; lane 7, molecular weight standards.

LANE

1

2

3

4

Fig. 5. Phosphorylation of p63. Amoebae were metabolically pulse labeled with [32P]orthophosphate. Secreted proteins were fractionated on an inhibitor-Sepharose matrix and the species eluted only by boiling in SDS were analyzed by SDS-gel electrophoresis. The lanes are: 1, molecular weight standard; 2, 3 and 4 samples metabolicallylabelled during the 0-l-h, 4-5-h and 8-9-h periods of starvation, respectively.

420 be eluted by boiling in SDS was subjected to SDS-polyacrylamide gel electrophoresis. An autogradiogram of such a gel is shown in Fig. 5. The major species obtained from the matrix had the mobility of p63 and showed a progressively increasing amount of 32p incorporation during 8 h of starvation and early development. Discussion

The major focus of this report has been to test our earlier suggestion that heterogeneity in physical properties among species of low°K m extracellular phosphodiesterase from vegetative cells could be due to varying stoichimetries of enzyme-inhibitor complexes in which the inhibitor has been inactivated by dithiothreitol but not released from the enzyme. The inhibitor is a polypeptide of M r 38000, a value similar to that reported by others [16, 27, 28]. On denaturing gels, the polypeptide of pure phosphodiesterase 65 is p65, and a peptide of this size is also present in the more complex forms, phosphodiesterase 95 and 185. The latter also contain a polypeptide the same size as the inhibitor (p38) and a larger species (p95). When crude phosphodiesterase 185 is purified on nondenaturing gels, it contains p65, 95 and 38. Purification of phosphodiesterase 95 and 185 on an inhibitor affinity column highly enriches p65 and 95. Crosslinking of mixtures of purified phosphodiesterase 65 and inhibitor protein with dimethylsuberimidate generates p95. Therefore, p95 may be a complex of p65 and p38. Since both of these polypeptides may be exceptionally rich in half-cystine residues, the detection of p95 in the absence of dimethylsuberimidate could be due to extensive disulfide bond formation or interchange. The stoichiometry of phosphodiesterase 185 is not known, but it is unlikely to be a simple dimer of p95 since phosphodiesterase 185 has a larger charge-to-mass ratio [11] and lower isoelectric point [12] than phosphodiesterase 95. We have no definite proof that p65 is identical among the species. However all species bind the phosphodiesterase inhibitor, which is highly specific for D. discoideum phosphodiesterase [29] and all bind conconavalin A [30]. We have compared tryptic fingerprints of p65 prepared from SDS gels of phosphodiesterase 95 and 185 with phos-

phodiesterase 65 (not shown) and all were very similar, but not of sufficient resolution to establish absolute identity. A genetic approach may be required to unequivocally establish whether these species are the product of a single or several closely related genes for p65. It is clear, however, as shown in Fig. 3, that in vitro the inhibitor can convert the less acidic into the more acidic species, and that treatment with dithiothreitol does not reverse or prevent this effect, although it does abolish the effects of the inhibitor on catalysis. This behavior can, therefore, explain the existence of major enzyme structural heterogeneity in vivo during late vegetative and early starvation phases. If there is a single gene for p65, one could speculate that this same gene is the source of developmental phosphodiesterase, and that the same phenomenon could account for the heterogeneity of this enzyme. The vegetative and developmental extracellular phosphodiesterase polypeptides have a number of features in common. Both bind the inhibitor and concanavalin A. For both vegetative and developmental phosphodiesterase, the enzyme is activated by inactivation of the inhibitor with dithiothreitol. Both generate three major species with a similar relationship of size and isoelectric point [11,12] to the vegetative stage species. As shown in Fig. 4, both p63 and p65 display a large mobility shift on SDS gels in the absence of/~-mercaptoethanol. Very few proteins exhibit such pronounced sensitivity to reducing agents [31]. This behavior, along with the copurification of a small amount of p63 with p38, even after heating, and the high enrichment of p63 by a crude inhibitor-Sepharose matrix constitute all of our evidence that p63 is the developmental phosphodiesterase. Assuming from this evidence that this is the case, it is possible that phosphorylation of the same p65 produced by vegetative amoebae could generate p63, account for the small increase in mobility on SDS-polyacrylamide gel electrophoresis, and that the formation of enzyme-inhibitor complexes could explain the existence of heterogeneity. It is known that the catalytic polypeptide produced during development is of similar size and isoelectric point [12,13]. Kessin and co-workers [15,16] mixed this pure enzyme with pure inhibitor and obtained a form equivalent to our phos-

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phodiesterase 95. They did not observe a form equivalent to our phosphodiesterase 185 in these mixtures, but the major phosphodiesterase form produced by starved cells was of this size. Their developmental phosphodiesterase 185 did not contain detectable inhibitor activity [13]. However, it did contain acidic polypeptide material in addition to the catalytic subunit, and pulsed additions of cAMP during starvation, conditions known to suppress inhibitor synthesis, resulted in suppressed appearance of phosphodiesterase 185 along with increased amounts of the free catalytic polypeptide. Rutherford and Brown [32] showed that treatment of large, native developmental extracellular phosphodiesterases with dithiothreitol did release some free (smaller) enzyme from phosphodiesterase 95, but not from their larger form, though both forms were activated by the dithiothreitol. There remain, however, other reports in apparent conflict with our speculation on the source of heterogeneity of developmental phosphodiesterase, and further studies will be required to resolve the issue. Dicou and Brachet [12] have observed three extracellular phosphodiesterase species in starvation supernatants which are identical to phosphodiesterase 65, 95 and 185 forms in size and charge-to-mass ratio behavior, but are from a mutant which does not produce active inhibitor. It remained possible that this mutant produced an inhibitor capable of binding the enzyme but lacking inhibitory activity, since our results show that dithiothreitol-treated inhibitor can still bind phosphodiesterase. These authors [28] also developed a phosphodiesterase affinity column capable of isolating homogeneous inhibitor from total crude supernatants. When the mutant was tested, no inhibitor polypeptide was recovered. However, it remains possible that the mutant does produce an inhibitor polypeptide with both defective inhibition and weaker binding properties than the inhibitor from wild type. Thus, the mutant inhibitor might be eluted from the affinity column by the high salt washes used prior to the conditions which elute the native protein. This is a viable possibility in that the larger phosphodiesterase fractions from the mutant could regenerate some of the smaller forms, whereas we have not observed formation of the smaller species from the larger ones, and

Franke and Kessin [16] did not observe significant dissociation of a 1:1 inhibitor-catalytic subunit complex. McDonald and Sampson [33] have isolated the same three types of phosphodiesterase species from amoebae starved by shaking in suspension in the presence of cAMP to suppress inhibitor synthesis. They studied only species which were inhibitor-free by the criterion of failure of dithiothreitol to cause activation. However, they did not test directly for the presence of the inhibitor polypeptide, and the Coomassie stain used, in our hands, fails to detect this species. Further, it is possible that redox conditions in handling of cultures and enzyme preparations could interfere with detection of inhibitor activity. Therefore, while it is difficult to explain all the published results on these enzymes by our in vitro enzyme interconversions (Fig. 3), reports of the absence of inhibitor from the larger enzymes are all subject to alternate possible interpretations at this time. Reinvestigation of the larger enzyme species is required to firmly resolve the issue, since, in vitro, it is clear that the dithiothreitoltreated inhibitor is still able to interact with the enzyme and that treatment of inhibited complexes with dithiothreitol does not reverse the inhibitorinduced changes in extracellular phosphodiesterase physical properties (size and isoelectric point) despite restoration of catalytic activity.

Acknowledgements This research was supported by N.S.F. grant PCM 77-25378 to E.J.H.

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