Planta

Planta 149, 130-137 (1980)

9 by Springer-Verlag 1980

Identification and Subcellular Localization of Starch-Metabolizing Enzymes in the Green Alga Dunaliella marina Erich Kombrink and Giinter W6ber Fachbereich Chemie(Biochemie)der Philipps-UniversitM,Lahnberge,D-3550 Marburg, Federal Republicof Germany

Abstract. Enzymes of starch synthesis and degrada-

tion were identified in crude extracts of the unicellular green alga Dunaliella marina (Volvocales). By polyacrylamide gel electrophoresis and specific staining for enzyme activities, 4 multiple forms of starch synthase, 2 amylases, and at least 2 forms of ct-glucan phosphorylase were visible. Using specific a-glucans incorporated into the gel before electrophoresis we have tentatively correlated a-amylase and /?-amylase with both hydrolytic activities. The activities of aglucan phosphorylase and amylase(s) were measured quantitatively in crude extracts, and the concomitant action of a-glucan phosphorylase and amylase(s) was found to account for the fastest rate of starch mobilization observed in vivo. Isolated chloroplasts retained both typical plastid marker enzymes and ADPglucose pyrophosphorylase, starch synthase, amylase(s), and a-glucan phosphorylase to a similar percentage. Gel electrophoretic analysis followed by staining for enzyme activity of a stromal fraction resulted in a pattern of multiple forms of starch-metabolizing enzymes analogous to that found in a crude extract. We interpret the combined data as indicating the exclusive location in vivo of starch-metabolizing enzymes in chloroplasts of D. marina. Key words: ADPglucose pyrophosphorylase - Chlo-

roplast Enzyme compartmentation- Enzymes, multiple- a-Glucan phosphorylase - Starch metabolism. Abbreviations: Chl=chlorophyll; DEAE-dextran=diethylamino-

ethyl-dextran; DDT = dithiothreitol; EDTA= ethylenediaminetetraacetic acid; FBPase=fructose-l,6-bisphosphate phosphatase, EC 3.1.3.11 ; G1P=glucose 1-phosphate; G6P-DH=glucose 6phosphate dehydrogenase, EC 1.1.1.49; HEPES=N-2-hydroxyethylpiperazine-N'-ethanesulphonic acid; MES=2-(N-morpholino)ethanesulphonic acid; Pi=inorganic orthophosphate; RuBP carboxylase= ribulose-1,5-bisphosphate carboxylase,EC 4.1.1.39

0032-0935/80/0149/0130/$01.60

Introduction

The pathway of starch synthesis in those parts of plants containing the photosynthetic apparatus and involving ADPglucose pyrophosphorylase, starch synthase, and branching enzyme has been elucidated in both higher plants and algae (Preiss and Levi 1979). In contrast, the identity of enzymes participating in starch breakdown during the dark period remains to be clarified. While some studies suggest a concomitant action of amylases and a-glucan phosphorylase in starch mobilization (Haapala 1969; Wanka et al. 1970; Pongratz and Beck 1978), other investigations demonstrated that neither a-amylase nor/?-amylase are located in chloroplasts (Levi and Preiss 1978; Stitt et al. 1978). Thus, whereas participation of a-glucan phosphorylase is generally accepted, the contribution of amylolytic hydrolysis of starch is at present unclear. Indeed, multiple forms of phosphorylase have been identified both in various green tissues of higher plants (Steiger and Abel 1975; Kumar and Sanwal 1977; Richardson and Matheson 1977; Steup and Latzko 1979), green algae, and bluegreen bacteria (Fredrick 1967 ; Fredrick 1973 ; Mangat t979). However, the picture is further complicated by the fact that multiple forms of a-glucan phosphorylase seem to be located both within and outside of the chloroplast (Steup and Latzko 1979). Although one might tend to intuitively consider only plastidic enzymes as being relevant to transitory starch metabolism, reports to the contrary must not be summarily dismissed as artifacts of one sort or another. Therefore, we decided to investigate enzymes of starch metabolism and their compartmentation in the unicellular green alga Dunaliella marina. The choice of this organism offers several advantages over higher plants as experimental systems.

E. Kombrink and G. W6ber: Starch-Metabolizing Enzymes in Dunaliella (1) U n i c e l l u l a r g r e e n a l g a e , b e i n g m i c r o o r g a n i s m s , can be cultured under standardized conditions. This may help to preclude ambiguous or misleading results because of tissue inhomogeneity, diurnal fluctuation, or seasonal variation of enzyme activity with higher plants. (2) M o s t i m p o r t a n t l y , D. m a r i n a as a n a t u r a l p r o t o plast lends itself to a novel approach for the isolation o f i n t a c t c h l o r o p l a s t s b y c h e m i c a l l y i n d u c e d lysis o f cells t h e r e b y a v o i d i n g a n y m e c h a n i c a l stress. A f t e r purification by isopycnic centrifugation in a sucrose density gradient, this method produces chloroplasts r e t a i n i n g t h e i r s t r o m a l e n z y m e s in a 3 0 - 4 0 % y i e l d ( K o m b r i n k a n d W 6 b e r 1980).

Materials and Methods Organism and Culture Conditions. Dunaliella marina (LB 19-4, Culture collection, Universitfit G6ttingen) was cultivated in the synthetic medium according to McLachlan (1960), as specified previously (Kombrink and W6ber 1980). Cell-J?ee Extracts. Algae were harvested from the culture medium (600 ml) by centrifugation (10 rain, 500 g) and washed twice with a solution containing 0.5 M sorbitol and 5 mM HEPES-NaOH buffer, pH 7.5, to remove salts. The cells were then suspended in 15 ml chilled buffer (50 mM HEPES-NaOH, pH 7.5) and homogenized with an Ultra-Turrax (3 30-s pulses). The clear supernatant solution obtained after centrifugation (30 rain, 30,000 g) was used in enzyme assays and electrophoretic experiments. In some cases the sediment was extracted again by homogenization, as described above, to obtain any enzyme activity possibly associated with the pellet. Isolation of Chloroplasts and Subcellular Fractions. Chloroplasts were isolated and purified by DEAE-dextran-induced lysis of cells followed by isopycnic centrifugation in a sucrose density gradient (Kombrink and W6ber 1980). The lysed preparation was separated by centrifugation (1 min, 500 g) into a crude chloroplast fraction and a supernatant fraction containing the remainder of the cytoplasmic constituents. After centrifugation (30 min, 30,000g), this supernatant fraction was adjusted to 15 mM and 0.5 mM in HEPES-NaOH buffer, pH 7.5, and DTT, respectively, for the assay of enzyme activities. The four fractions obtained after sucrose density gradient centrifugation of crude chloroplasts and containing, in the order of increasing buoyant density, soluble material, broken chloroplasts, intact chloroplasts, and aggregated fragment at the bottom of the tube were adjusted to 25mM and 0.5mM in HEPES-NaOH buffer, pH 7.5, and DTT, respectively. They were then treated with an Ultra-Turrax (3 30-s pulses) to disrupt the organelles. The pellet after centrifugation (30 min, 30,000 g) was re-extracted with buffer as above and centrifuged again, after which the respective supernatant solutions were combined. These fractions were concentrated to a few ml and dialized against 20 mM HEPES-NaOH buffer, pH 7.5, in an Amicon pressure dialysis ceil (PM-10 membrane). Polyacrylamide Gel Electrophoresis. Electrophoreses were performed in 5% cylindrical gels with a discontinuous buffer system as described by Davis et al. (1967). The buffer in the separation gel was 30 mM HC1 adjusted to pH 7.9 with 2 M Tris base. The electrode buffer was 35 mM L-asparagine in 10% sucrose, adjusted to pH 7.3 with 2 M Tris base. Enzyme solutions were diluted 1 : 1

131

with a sample buffer containing 20% sucrose and imidazole-HC1, pH 5.8, such that the final concentration of chloride ions was 0.48 M. Bromphenol blue was used as a tracking dye. Electrophoreses were run without any sampling or stacking gel at 4~ and 2 mA per tube.

Procedures for Activity Staining of Gels. After electrophoresis, gels were washed with distilled water and stored at room temperature for up to 20 h in one of the following incubation mixtures, e-Glucan phosphorylase was revealed with a solution containing 30 mM G1P, 50raM MES-NaOH buffer, pH 6.5, 0.I mM DTT, and 0.01% maltodextrin mixture (average chain length 7 glucosyl units) or 0.1% soluble starch (Lee 1972). When soluble starch, maltodextrin mixture, or shell fish glycogen were polymerized into the gel, the primer in the incubation solution was omitted. Amylase activity was detected by incubating the gels in 50 mM HEPES-NaOH buffer, pH 7.5, containing 1% soluble starch. When polysaccharides had been added to the gels before polymerization, the gels were incubated in carbohydrate-free buffer. Starch synthase was revealed according to Schiefer et al. (1973) with a solution containing 1 mM ADPglucose, 1 mM DTT, 2 mM EDTA, 25 mM sodium citrate, 50 mM HEPES-NaOH buffer, pH 8.0, and 0.1% starch. After incubation, the gels were rinsed with distilled water and placed in an iodine solution (0.1% J2, 0.03 % K J, 0.05 M HC1). Polysaccharides synthesized by c~-glucan phosphorylase and starch synthase were visualized as brown, purplish, or blue bands. The action of amylases could be seen as unstained bands in a colored background. An alternative c~-glucan phosphorylase activity stain based on the visualization of the P~ formed (Davis et al. 1967) was also used. The incubation mixture contained 30 mM G1P, 0.2 M calcium chloride, 5% maltodextrin mixture, 50 mM sodium maleate buffer, pH 6.5, and 0.1 mM DTT. Controls of all procedures for activity staining either without substrate in an otherwise identical incubation or without incubation immediately after electrophoresis were routinely done to reveal unspecific or arteficial staining of the gels with iodine. Enzyme Activity Assays. ~-Glucan phosphorylase (EC 2.4.1.1) was measured in the phosphorolytic direction according to Lee and Braun (1973). The reaction mixture contained, in a final volume of 1 ml, 10 gmol sodium phosphate buffer, pH 7.0, 20 gmol MgC12, 2 gmol EDTA, 0.25 ~tmol NADP § 0.7 IU G6P-DH, 0.8 IU phosphoglucomutase, 2 mg soluble starch, and 50 gmol MES-NaOH buffer, pH 7.0. To obtain correct estimates of enzyme activity it was necessary to free the auxiliary enzymes of ammonium sulphate by dialysis. In a final volume of 1 ml, amylase (EC 3.2.1.1 ; EC 3.2.1.2) activity was determined by incubating 50 gmol HEPES-NaOH buffer, pH 7.5, and 2 mg soluble starch for 30 60 min at 30~ C. The activity was calculated from the amount of reducing end groups liberated by the enzyme and measured quantitatively with the alkaline copper reagent (Robyt and Whelan 1968) The starch synthase (EC 2.4.1.2I) activity test was modified after Preiss and Greenberg (1967) and Hawker et al. (1974), measuring the transfer of [U-J4C]glucosyl units from radioactively labelled ADPglucose to starch. The reaction mixture contained, in a total volume of 0.2 ml, 20 gmol Bicine-NaOH, pH 8.5 (N,N-bis(2hydroxyethyl)glycine), 0.16 ~tmol ADP-[U-14C]glucose (1.2.104 Bq gmol-1), 5gmol KC1, 2gmoI DTT, 1 ~tmol EDTA, and 1 mg soluble starch. After incubation for 15 min at 30 ~ C, the reaction was stopped by heating the mixture in a boiling-water bath for 1 min. The mixture was applied to an anion exchange resin (Dowex 1 x 4) in a Pasteur pipette equilibrated with 0.1 ml of a carrier starch solution (25 mg m1-1) for the separation of starch from unreacted ADPglucose. After washing with 3 ml distilled water, the eluate was counted for radioactivity by liquid scintillation counting. A control experiment without enzyme was run in parallel to check the capacity of the ion exchange column. Less than 0.5%

132

E. Kombrink and G. W6ber: Starch-Metabolizing Enzymes in Dunaliella

of the radioactivity in the incubation mixture was found in these control eluates. ADPglucose pyrophosphorylase (EC 2.7.7.27) activity was assayed by following the formation of the sugar nucleotide (Sanwal and Preiss 1967). The reaction mixture contained, in a final volume of 0.2ml, 20pmol Bicine-NaOH buffer, pH 8.5, 0.1 gmol [U14C]G1P (3.8- 10~ Bq gmol 1), 0.5 gmol ATP, 1.5 gmol MgC12, 0.4 ~tmol 3-phosphoglycerate, and 50 gg bovine serum albumin. In some experiments 2 gmol DTT was added to activate the enzyme. After incubation for 15 min at 30 ~ C, the reaction was terminated by heating the mixture in a boiling-water bath for 1 min. After the addition of 2 IU alkaline phosphatase (Boehringer), the incubation was continued for 30 min at 30 ~ C. With this treatment, unreacted [U-14C]G1P was hydrolyzed while the product ADPglucose remained intact. An aliquot of the reaction mixture was then adsorbed on a DEAE-cellulose paper disk and [14C]glucose was washed off with 4 150-ml portions of water. The ion exchange paper was dried and counted for radioactivity. A control incubation without algal extract made sure that unreacted [14C]G1P was not carried over to the DEAE-cellulose separation step, since less than 0.5% of the radioactivity in the incubation mixture remained on the paper after the wash. All other enzyme activity assays were performed according to documented methods as described previously (Kombrink and W6ber 1980).

Analytical Methods. Radioactive products of the starch synthase and ADPglucose pyrophosphorylase reaction were identified by descending paper chromatography on Schleicher and SchiilI, Nr. 2043b paper. The polymer product as obtained in the starch synthase assay after elution from the ion exchange resin was sub jected to paper chromatography in n-bntanol-pyridine-water (6:4:3, by volume) for 48 h before and after complete hydrolysis by glucoamylase (Diazyme, Miles). Reducing sugars were detected with the AgNO3-NaOH-Na2S20 3 reagents (Trevelyan et al. 1950). To reveal polysaccharides, the paper chromatogram was sprayed with a glucoamylase solution before developing. The identity of enzymically formed presumptive ADPglucose was confirmed by reference to an authentic sample after paper chromatography with the solvent 95% ethanol-1 M ammonium acetate, pH 7.5 (5:2), for 18 h and the same solvent as used above. Elution of the radioactive zone corresponding to ADPglucose and hydrolysis by 0.05 M HC1 gave rise to [l~C]glucose that was again authenticated. Aqueous radioactive solutions were counted by liquid scintilla tion (Packard Tri-Carb, Model 3380) in a cocktail containing per litre: 5 g butyl-PBD, 200 mt Scintisol solubilizer (Isolab-Biolab, Brussels), 800 ml toluene. Radioactivity on paper disks was counted directly in a cocktail without solubilizer. Radioactive zones on paper chromatograms were located with a scanner (Packard, Model 7201). Soluble protein was determined according to Lowry et al. (1951). Chlorophyll was determined according to Arnon (1949), and sucrose concentration was measured refractometrically. Chemicals. Auxiliary enzymes were purchased from Boehringer, Mannheim, [U-~*C]G1P, and ADP-[U-14C]glucose were obtained from Amersham. ADPglucose and DEAE-dextran were purchased from Sigma; all other chemicals were of the highest grade available.

Results

Formation of Starch in Dunaliella marina. Enzymes involved in the synthesis of starch, viz. ADPglucose pyrophosphorylase and starch synthase, have been characterized in both higher plants and the alga Chlo-

rella pyrenoidosa, while nothing is known about these enzymes in D. marina. To confirm that the chosen assays for ADPglucose pyrophosphorylase and starch synthase are applicable to extracts of D. marina, the linearity of enzyme action with time and protein concentration was studied (Fig. 1 A and B). Furthermore, the products formed in each enzyme reaction were fully characterized. The nature of the product formed by the transfer of glucosyl units from ADP-[Ut4C]glucose to a primer in the starch synthase reaction was confirmed in the following way. Enzymic degradation by glucoamylase of the radioactive reaction product that was insoluble in 66% aqueous ethanol to [14C]glucose as the only sugar demonstrated that the polysaccharide is a homopolymer comprising eglycosically linked glucose units. If a mixture of the ADPglucose pyrophosphorylase assay at about 50% turnover of reactants after prolonged incubation was inspected by paper chromatography, radioactivity was distributed equally between ADPglucose and glucose by reference to authentic samples. ADPglucose itself was identified by re-chromatography in 2 solvents (see Methods), and hydrolysis with 0.05 M HC1 gave complete conversion into only 1 radioactive product that was again authenticated as glucose. Thus, ADPglucose pyrophosphorylase and starch synthase have been demonstrated to occur in D. marina, and the quantitative measurement of enzyme activity (see below) is reliable. Breakdown of Starch in D. marina. ~-Glucan phosphorylase found in D. marina has a neutral pH-optimum and acts equally well with either soluble starch, shell fish glycogen, or maltodextrin mixture as a substrate (results not shown). Amylase from D. marina has a pH-optimum around 8.0 in contrast to a value around pH 6 for amylase from other sources (Wanka et al. 1970; Pongratz and Beck 1978 ; Stitt et al. 1978), and the activity is not affected by either Ca 2+, EDTA, or DTT (Bergmann 1976). Other enzymes which may conceivably be involved in starch degradation, e. g., debranching enzyme or ~-glucosidase, could not be detected. Polyacrylamide gel electrophoresis is a well-established technique for the detection of multiple forms of starch synthase or c~-glucan phosphorylase by means of staining for enzyme activity. Fig. 2 is a composite representation of results obtained after numerous electrophoretic analyses of crude algal extracts. Two amylolytic activities can be seen in gel A, a white band 1 and a purplish band 2 against a dark blue background (for experimental details cf. Materials and Methods). The addition of Pi to the incubation medium did not alter this pattern, thus, precluding any interference by c~-glucan phosphory-

E. Kombrink and G. W6ber: Starch-Metabolizing Enzymes in Dunaliella

100

100

A

80

.o

'~

/ Q ~

80

/i/j

60

0

Fig. 1 A and B. Linearity of the starch synthase ( - e e - ) and the ADPglucose pyrophosphorylase assay (ax-zx-) with protein concentration (A) and incubation time (B). The composition of the assay is described in Materials and Methods; incubation time 15 min for A, protein concentration 40 p,g for B

40

2O

O

133

/

20

40 Protein

I

80 { iJg )

I

120

0

o

I

,o

2o

Incubation

Fig. 2A-E. Separation by polyacrylamide gel electrophoresis of a crude cell homogenate of Dunaliella marina followed by specific staining for enzyme activities. For amylases by incubation with 1% soluble starch for 18 h (A), for ~-glucan phosphorylase by incubation with 30 mM G1P and 0.1% starch for 6 h (B) or for 18 h (C), for starch synthase by incubation with 1 mM ADPglucose and 0.1% starch for 6 h (D) or for 18 h (E). Migration towards the anode. The product of enzyme action was revealed with an iodine solution

lase. The latter enzyme could easily be detected as a starch-synthesizing activity after incubation with G1P and either soluble starch or a maltodextrin mixture (Fig. 2B and C). e-Glucan phosphorylase did not coincide with either amylase. Four multiple forms of starch synthase are distinguishable in Fig. 2D and E. The bands differed in the color of the carbohy-

," time

20 ( rain

50

)

drate-iodine adduct, some turning from blue after a short incubation to a reddish color after long incubation periods. This behavior suggests an intimate association of starch synthase with branching enzyme such as that observed previously (Schiefer et al. 1973). An attempt was made to differentiate between the 2 amylases by adding specific e-glucans to the gels before polymerization (Fig. 3). Although the overall pattern of enzyme activity was similar, enzymes were considerably retarded by interaction with substrates during electrophoresis (compare Schiefer et al. 1973). Gels A and B had been loaded with 0.02% soluble starch and each showed 2 amylases after 0.5 h and 4 h incubation, respectively. The same result was found with other e-glucans such as amylose and amylopectin (not shown). In contrast, if 0.035% amylopectin /?-amylase limit dextrin was included in the gels, only one amylase was seen after 0.5 h incubation (Fig. 3 C). Zone 2 was only revealed after 4 h incubation in the presence of amylopectin /?-amylase limit dextrin (Fig. 3D). Although corroborative evidence is clearly needed, we can tentatively assign zones 1 and 2 to e-amylase and/?-amylase, respectively. The primer (snbstrate) specificity of e-glucan phosphorylase was also studied by electrophoretic techniques. Amylopectin and shell fish glycogen were ineffective in priming the reaction, presumably because these macromolecules did not diffuse into the gel matrix within the incubation period. If the primer was polymerized into the gel at different concentrations, additional multiple forms of c~-glucan phosphorylase became apparent. In Fig. 4A, one band is seen in the gel that had been primed with 0.02% soluble starch similar to the incubation with externally added starch (cf. Fig. 2B and C). When 0.2% real-

134

E. Kombrink and G. W6ber: Starch-Metabolizing Enzymes in Dunafiella

Fig. 3A-D. Separation by polyacrylamide gel electrophoresis of a crude cell homogenate of Dunaliella marina followed by staining for amylolytic activity, c~-Glucan (0.02% starch with A and B or 0.035% amylopectin /?-amylase limit dextrin with C and D) was added to the gel tube before polymerization. After electrophoresis, gels were incubated in buffer for 0.5 h (A and C) or 4 h (B and D) and then stained with iodine solution

Fig. 4A-E. Separation by polyacrylamide gel electrophoresis of a crude cell homogenate of Dunaliella marina followed by staining for c~-glucan phosphorylase. Different c~-glucans were added to the gel tubes before polymerization (0.02% starch, A; 0.2% maltodextrin mixture, B and C; 0.2% shell fish glycogen, D and E). After electrophoresis, the gels were incubated with 30 mM G1P for 4 h (B and D) or 14 h (A, C and E) and then stained with iodine solution

todextrin mixture (Fig. 4B and C) or shell fish glycogen (Fig. 4D and E) had been included in the gel, a faster moving band with c~-glucan phosphorylase activity was revealed. Only these ~-glucans could be tested at such high concentration because they do not stain with iodine appreciably.

The cross-contamination of fraction 3 containing intact chloroplasts with cytoplasm, mitochondria, and microbodies was 3.8%, 4.7%, and 28%, measured as pyruvate kinase, fumarase, and catalase, respectively (not shown in Table 1). e-Glucan phosphorylase and amylase were located in chloroplasts of similar percentages to the marker enzymes, FBPase and RuBP carboxylase. The corresponding figures for starch synthase and ADPglucose pyrophosphorylase are somewhat lower, the reason for which is unknown. The values of fraction 1 and 3 (Table 1) were recalculated to afford specific enzyme activities and are shown in Table 2. A ratio of specific activities of enzymes in fractions 1 and 3 (ratio 3/1) should give a similar figure for all enzymes supposedly of plastidial origin, given an unavoidable rupture of organelles during ultracentrifugation in a sucrose density gradient with a consequent leakage of chloroplast enzymes. For this to be true, selective leakage from or binding of some enzymes to chloroplasts is tacitly excluded. Indeed, the figures for enzymes either established or presumed to occur in chloroplasts are within the same order of magnitude, whereas this value is very different for enzymes known to be located in other organelles.

Subcellular Distribution of Enzymes. We used a novel technique for the isolation and purification of intact chloroplasts from D. marina (Kombrink and W6ber 1980) to study the subcellular distribution of starch-metabolizing enzymes that had been identified in crude cell homogenates. Using chemically induced lysis followed by isopycnic centrifugation in a sucrose density gradient, the cellular components were separated into 5 fractions: fraction 1 comprising soluble material, fraction 2 broken chloroplasts, fraction 3 intact chloroplasts, fraction 4 aggregated sediment, and fraction 5 the lysis supernatant suspension containing cytoplasmic components, except chloroplasts. The identity and purity of these fractions has been established previously (Kombrink and W6ber 1980). The distribution of some plastid marker enzymes and of starch-metabolizing enzymes is shown in Table 1. In addition, figures are given in Table 1 for the recovery of each enzyme after cell fractionation in relation to the activity in a crude extract.

E. Kombrink and G. W6ber: Starch-Metabolizing Enzymes in Dunaliella

135

Table 1. Distribution of enzymes in subcellular fractions of Dunaliella marina. Cells were chemically lysed by DEAE-dextran and centrifuged to separate crude chloroplasts from the lysis supernatant fraction. The crude chloroplast-containing pellet was purified by isopycnic density gradient centrifugation (4 h, 100,000g) to give 4 distinct fractions corresponding to soluble components, 1, broken chloroplasts, 2, intact chloroplasts, 3, and aggregated sediment, 4. The sum of enzyme activities (nmol substrate used per min) in these 5 fractions is given in column 2; and related, in terms of percentage recovery, to the enzyme activity in a crude cell extract Lysis supernatant fraction Protein (mg) (%)

Chlorophyll (mg)

Fraction 1

Fraction 2

23.2

9.2

16.0

6.1

8.2

39.1

15.5

26.9

10.3

--

--

-

7,877

(%)

7.2

7.9

3.2

39.3

43.4

17.5

420.4

5,886

55.6

FBPase

159.4

(%)

c~-Glucanphosphorylase (%)

Amylase

3,716

3.0

41.5

19.7

2,370 36.3

3.9

185.8

8.2

134.8

19.3

3.6

51.5

2.3

47.9

64.7

ADPglucose pyrophosphorylase (%)

1,203

Catalase

4.7

5.6

164.7

10t .4

10.7

6.6

89.0

171.0

4.7

86.7

22.3

23.8

45.8

1.3

23.2

6.0

Fraction 1

Fraction 3

Ratio 3/1

Crude cell extract

340.0

11,120.0 32.71

30,972

13.2

1.1

0.08

5.4

49.3

3.5

0.07

145.4

84,8

73.2

0.86

91.4

RuBP carboxylase

339.5

367.9

1.08

241.6

EBPase

160.2

148.4

0.93

156.2

c~-Glucan phosphorylase

8.0

8.4

1.05

11.3

Amylase

5.4

7.8

1.44

9.6

51.9

10.3

0.20

48.9

7.4

5.4

0.74

374.5

ADPglucose pyrophosphorylase

47.4 0.4

Pyruvate kinase

Starch synthase

5.3

12.4

78.1

Fumarase G6P-DH

37.4

125.1

4.2

Table 2. Specific enzyme activities (nmol min- 1 mg- 1 protein) and the ratio of specific activities in fractions of a chloroplast preparation compared with those in a crude cell extract. Data from Table 1 were recalculated, and the designation of fractions corresponds with Table 1 Enzyme

257.0

12.9

-

If a gross recovery of a b o u t 50% intact chloroplasts together with a loss of s t r o m a l enzymes from the r e m a i n i n g chloroplasts into the s u p e r n a t a n t fraction is taken into account, it can be c o n c l u d e d that ~-glucan phosphorylase, amylase, A D P g l u c o s e pyro-

Crude cell extract

93

100

83

100

59.4 1O0

63.8

18.3 100

22.0 I5,430

100

0.3

126.3

S

14,180

57.0

(%) (%)

-

2.4

-

Starch synthase

Fraction 4

4.88

(%)

RuBP carboxylase

Fraction 3

92

6,523 100

66

360.9 100

100

721.0 50

100

43

100

1,540

3,123

263.8 100 I00

612.6

49

373.7 100

100

9,966

100

23,890 1.6

I00

phosphorylase, and starch synthase are m o s t likely restricted in d i s t r i b u t i o n to the chloroplast of D. marina. Characterization o f Chloroplast Starch-metabolizing E n z y m e s . Evidence for multiple forms of starch-meta-

bolizing enzymes present in chloroplasts of D . m a r i n a is given in Fig. 5. S t r o m a l enzymes o b t a i n e d after r u p t u r e of intact chloroplasts, dialysis a n d c o n c e n t r a tion were subjected to disc gel electrophoresis a n d stained for activity. Two amylases were seen either when gels were i n c u b a t e d with 0.5% soluble starch (Fig. 5A) or when 0.02% starch had been polymerized into the gels ( i n c u b a t i o n for 0.5 h, Fig. 5B, or for 4 h, Fig. 5C). Two multiple forms of c~-glucan p h o s p h o r y l a s e were seen with 0.2% glycogen in the gels (incubation in the presence of G 1 P for 10 h, Fig. 5D, or for 21 h, Fig. 5E). W i t h starch synthase, only enzymes 2 and 3 ( c o m p a r e Fig. 2 D a n d E) could be detected when A D P g l u c o s e a n d 0.1% starch were added externally in i n c u b a t i o n s for 6 h (Fig. 5 F ) or for 21 h (Fig. 5 G). T h a t enzyme 3 formed a red b a n d a n d enzyme 2 a p u r p l i s h - b l u e b a n d subscribe to the fact that starch synthase a n d b r a n c h i n g enzyme were still firmly associated. Band 4 was only faintly visible because of the interference of a fairly active amylase in this zone. W e suggest that some multiple forms of starch synthase were inactivated or lost dur-

136

E. Kombrink and G. W6ber: Starch-MetabolizingEnzymesin Dunaliella

Fig. 5. Separationby polyacrylamidegel electrophoresis of stromal enzymes of Dunaliella marina followed by staining for enzymeactivities.For amylases (A-C): 0.5% starch was added externally to the incubation medium (A) or 0.02% starch was added to the gel tube before polymerization(B and C). Incubation time was 21 h (A), 0.5 h (B) or 4 h (C). For c~-glucanphosphorylase (D, E) : 0.2% shell fish glycogenwas added to the gel tube before polymerization.Incubation with 30 mM G1P was for 10 h (D) or 21 h (E). For starch synthase (F, G) : After electrophoresis,gels were incubated with 1 mM ADPglucose and 0.1% starch for 6 h (F) or 21 h (G). The product of enzyme action was revealed with an iodine stain

ing the time-consuming preparation of chloroplasts and a stromal extract.

Discussion If Dunaliella marina is transferred from the light to the dark, a breakdown of starch with an initially high rate of 64 nmol glucose per min and mg Chl is observed which falls to 16 nmol glucose per min and mg Chl after 2 to 3 h (Bergmann 1976). The activity of a-glucan phosphorylase and amylase in crude extracts of D. marina was found to be 33 and 28 nmol glucose per min and mg Chl, respectively (compare Table 1). Thus, both enzymes seem necessary to account for rapid starch mobilization, if only because a-amylase is thought to be capable of initiating the attack on the starch granule (Levi and Preiss 1978; Bailey and MacRae 1973). When the degration products of radioactively labeled starch upon incubation of isolated spinach chloroplasts under varying reaction conditions were identified, the operation of both an amylolytic and phosphorolytic pathway was inferred from the results (Steup et al. 1976; Levi and Gibbs 1976; Peavey et al. 1977). When enzyme activities were investigated, however, considerable differences were concluded to exist both with regard to the occurrence of starch-degrading enzymes in plant species and their subcellular distribution. Pea chloroplasts were reported to contain only a-glucan phosphorylase, while they lacked a-amylase and/3-amylase (Levi and Preiss 1978; Stitt et al. 1978). In spinach leaves, on the other hand, although chloroplasts apparently contain both amy-

lase and a-glucan phosphorylase, a high percentage of these enzymes has been found outside of the plastids (Pongratz and Beck 1978 ; Okita et al. 1979). Furthermore, an exclusive extraplastidial localization has been reported for certain multiple forms of these enzymes (Steup and Latzko 1979). Our experiments attempt a new approach to the problem of subcellular localization of starch-metabolizing enzymes that differ from the protocol of other investigators in 2 main ways. First, by using a unicellular microorganism instead of a highly differentiated higher plant, complications caused by tissue inhomogeneity do not exist. Second, a replacement of mechanical rupture of tissue by chemical lysis of the cells may avoid systematic artefacts inherent in the former method. Irrespective of the reason for these contradictory results with higher plants, we find that not only all of the starch-degrading enzymes but also all of the multiple forms of amylase and starch phosphorylase are located within the chloroplast of D. marina.

Although both a-glucan phosphorylase and amylase as the plastidial enzymes may participate in the starch breakdown, nothing is known about the signal that brings about the switch from starch accumulation to mobilization as a consequence of a light-dark transition. Neither enzyme was inhibited by DTT, as would be expected for regulatory redox changes of dark-activated enzymes (Buchanan et al. 1979). So far only one example of a light-regulated a-glucan phosphorylase has been reported (Lehmann and W6ber 1978). In contrast, stromal ADPglucose pyrophosphorylase, the allosterically regulated enzyme of starch biosynthesis (Preiss 1978), was found to be

E. Kombrink and G. W6ber: Starch-Metabolizing Enzymes in Dunaliella a c t i v a t e d 5 to 50 fold b y l 0 m M D T T even in the p r e s e n c e o f 2 m M 3 - p h o s p h o g l y c e r a t e a n d 7.5 m M M g 2§ (results n o t s h o w n ) . As w i t h o t h e r r e g u l a t o r y e n z y m e s o f t h e C a l v i n cycle, b o t h k e y m e t a b o l i t e s o f low m o l e c u l a r w e i g h t a n d r e d o x c h a n g e s c a n m o d ulate the enzyme activity concurrently. It r e m a i n s to be seen w h e t h e r the m u l t i p l e f o r m s o f c~-glucan p h o s p h o r y l a s e in D . m a r i n a per se or the a p p a r e n t l y d i f f e r e n t a f f i n i t y to s t a r c h o r m a l t o d e x trins h a v e a n y r e g u l a t o r y significance. T h e i n t e r p r e t a t i o n o f gel p a t t e r n s after e l e c t r o p h o r e s i s o f a c r u d e extract can become complicated and misleading, and e x p e r i m e n t s to p u r i f y these e n z y m e s are u n d e r w a y . T h e q u e s t i o n , a m o n g m a n y others, of w h e t h e r these m u l t i p l e f o r m s are i s o m e r s d i f f e r i n g in charge, size ( H e d r i c k a n d S m i t h 1968), o r s u b s t r a t e specificity may then be answered. Financial support by the Deutsche Forschungsgemeinschaft (SFB 103/A5) is gratefully acknowledged.

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Received 23 September 1979; accepted 27 January 1980