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. 1998 Oct;118(2):581–590. doi: 10.1104/pp.118.2.581

Characterization of Starch-Debranching Enzymes in Pea Embryos1

Zhi-Ping Zhu 1, Christopher M Hylton 2, Ute Rössner 2,2, Alison M Smith 2,*
PMCID: PMC34834  PMID: 9765544

Abstract

Two distinct types of debranching enzymes have been identified in developing pea (Pisum sativum L.) embryos using native gel analysis and tests of substrate preference on purified or partially purified activities. An isoamylase-like activity capable of hydrolyzing amylopectin and glycogen but not pullulan is present throughout development and is largely or entirely confined to the plastid. Activities capable of hydrolyzing pullulan are present both inside and outside of the plastid, and extraplastidial activity increases relative to the plastidial activity during development. Both types of debranching enzyme are also present in germinating embryos. We argue that debranching enzymes are likely to have a role in starch metabolism in the plastid of the developing embryo and in starch degradation during germination.

The aim of this work was to characterize the starch-debranching enzymes of pea (Pisum sativum L.) embryos. Two distinct types of debranching enzymes occur in higher plants. Enzymes that can hydrolyze the α1,6-linkages in the yeast glucan pullulan and in limit-dextrins of amylopectin are known as limit-dextrinases, R-enzymes, or pullulanases (Nakamura, 1996). Enzymes that cannot attack pullulan but can debranch amylopectin and its limit-dextrins are known as isoamylases. Debranching enzymes are involved in the degradation of starch in germinating or sprouting starch-storing organs (Manners, 1985), and evidence from mutant cereals and Chlamydomonas reinhardtii indicates strongly that they are also important in starch synthesis. The maize endosperm mutant sugary1 (su1), the rice endosperm mutant sugary1 (su1), and the sta7 mutant of C. reinhardtii accumulate a highly branched, soluble, α1,4,α1,6-linked glucan polymer, phytoglycogen, in addition to or instead of starch (Mouille et al., 1996; Nakamura, 1996). All three mutants have reduced activity of debranching enzyme (Pan and Nelson, 1984; Doehlert et al., 1993; Mouille et al., 1996; Nakamura et al., 1996b, 1997), indicating that debranching activity is required for normal amylopectin synthesis. Ball and colleagues (1996) recently suggested that the synthesis of amylopectin and its organization to form a starch granule involves the “trimming” by debranching enzyme of highly branched, phytoglycogen-like material synthesized by starch synthase and starch-branching enzyme at the surface of the starch granule.

The precise way in which debranching enzymes are involved in amylopectin synthesis is difficult to assess because of the lack of information about the occurrence and nature of these enzymes in starch-synthesizing organs. First, it is not clear whether a lack of isoamylase, limit-dextrinase, or both types of enzyme is responsible for the phenotypes of the su1 and sta7 mutants. The su1 mutants are reported to have reduced activity of limit-dextrinase, but this enzyme is not encoded at the su1 locus in either maize or rice (James et al., 1995; Nakamura et al., 1996a). The su1 mutation of maize lies in a gene encoding a putative isoamylase (James et al., 1995), but the effect of the su1 mutation on isoamylase activity has not been reported. The genes at the su1 locus of rice and the sta7 locus of C. reinhardtii have not been identified.

Second, relatively little is known about these enzymes in higher plants. Limit-dextrinase activity occurs in leaves, embryos, and endosperms of several species (Nakamura, 1996). There is evidence that the enzyme in developing rice endosperm is plastidial (Nakamura et al., 1996a), and both plastidial and extraplastidial activities have been found in leaves (Okita et al., 1979; Li et al., 1992; Ghiena et al., 1993), but the location of the activity and the number of different gene products involved in other organs are not known. Isoamylase activity has been reported only from maize endosperm, in which it is plastidial (Doehlert and Knutson, 1991; Yu et al., 1998), and potato tuber, in which its intracellular location is not known (Ishizaki et al., 1983). This apparently limited distribution probably reflects the lack of research on other organs.

To provide more information about the occurrence and possible role of debranching enzymes in starch synthesis, we studied these enzymes in developing pea embryos. These are among the best characterized starch-synthesizing organs, but, to our knowledge, the occurrence of debranching enzymes has not previously been investigated. Limit-dextrinase is known to be present in mature seeds of legumes such as pea (Yellowlees, 1980), broad bean (Gordon et al., 1975), and mung bean (Morinaga et al., 1997). In this paper we describe the identification, characterization, and localization of isoforms of debranching enzyme present during starch synthesis in the developing pea embryo. These isoforms are then compared with those of the germinating embryo in which starch is being degraded.

MATERIALS AND METHODS

Plant Material

All pea (Pisum sativum L.) plants were of the BC1/RR or BC1/rr lines derived from JI 430 (John Innes Centre germplasm collection) by Hedley et al. (1986) and used in our laboratory for previous work on the characterization of starch synthesis in pea embryos. The BC1/rr line was used in the preparation of plastids, and the BC1/RR line was used in all other experiments. For developing embryos, pea plants were grown in a greenhouse in the conditions described by Smith (1988). Seeds were harvested on ice, the testas were removed, and the embryos were used immediately.

For imbibed and germinating embryos, seeds were soaked in running tap water overnight (imbibition), and were then germinated between damp paper towels in the dark at 25°C. Testas were removed from dry, imbibed, and germinating seeds before use in experiments.

Assay of Starch-Debranching Enzyme

Embryos were extracted with a mortar and pestle in approximately 3 volumes of extraction medium (100 mm Mes, pH 6.0, 50 mL L−1 ethanediol, and 5 mm DTT). The extract was centrifuged at 15,000g for 10 min. The supernatant was desalted on a column of Sephadex G-25 (Pharmacia) equilibrated with extraction medium and assayed as follows.

The assay medium for limit-dextrinase (pullulanase, EC 3.2.1.41), for glycogen-hydrolyzing activity during partial purification of isoamylase, and for determination of the glucan substrate specificities of purified and partially purified enzymes contained 100 mm Mes, pH 6.0, 20 g L−1 glucan substrate, and 25 to 75 μL of extract in a final volume of 0.1 mL. After incubation at 37°C for 1.5 to 2 h (during which time activity was linear with respect to time), reducing sugars were assayed with dinitrosalicylic acid reagent according to the method of Bernfeld (1951). The standard curve contained known amounts of maltose and the same volume of extraction medium as the assays. Control assays were stopped by the addition of dinitrosalicylic acid reagent immediately after addition of the extract. Assays using Red Pullulan were carried out according to the manufacturer's instructions (Megazyme International, Bray, County Wicklow, Ireland) on extracts derived as described above.

Assays in which maltoheptaose was the substrate were performed as described above, except that they contained 18 mm maltoheptaose instead of glucan. After various periods of incubation (from 15 to 90 min), assays were stopped by boiling and assayed spectrophotometrically for Glc according to the method of Lowry and Passonneau (1972).

Analysis of Products of Enzymic Digestion

When glucan substrates were digested to completion, the assay contained 100 mm Mes, pH 6.0, and 2 g L−1 glucan substrate. Incubation was for 8 h at 37°C. β-Limit-dextrin used in these experiments was prepared from amylopectin according to the method of Enevoldsen and Manners (1994).

For analysis of the products of digestion of amylopectin, 0.5 mL of potato amylopectin at 1 mg mL−1 was incubated at pH 4.0 with 5000 units of commercial isoamylase from Pseudomonas amyloderamosa (Sigma) or at pH 5.4 with either 5 units of α-amylase from pig pancreas (Boehringer Mannheim) or partially purified isoamylase from pea embryos (25 nmol min−1, determined by assay for glycogen-hydrolyzing activity, as described above). After incubation at 37°C for 16 h, samples were subjected to high-performance anion-exchange chromatography (Carbopac PA100 column with pulsed amperometric detection, Dionex Ltd., Camberley, Surrey, UK), as described by Tomlinson et al. (1997).

Purification of Pullulanase

All steps were carried out at 0°C to 4°C. About 400 g of developing embryos (individual embryos of 400–600 mg fresh weight) was homogenized in a blender in 600 mL of medium A (20 mm Bis-Tris propane, pH 7.0, and 5 mm DTT). The homogenate was centrifuged at 10,000g for 15 min, and the supernatant was subjected to ammonium sulfate fractionation. Protein precipitating between 30% and 40% saturation was collected by centrifugation, redissolved in a small volume of medium A, and then dialyzed against medium A. The dialyzed sample was applied at a flow rate of 0.5 mL min−1 to a column (21 cm high, 2.6 cm internal diameter) of DEAE-Sepharose Fast Flow (Pharmacia) equilibrated with medium A. The column was washed with medium A and then eluted at a flow rate of 2.5 mL min−1 with a 250-mL linear gradient of 0 to 1 m NaCl in medium A.

Five-milliliter fractions were collected and assayed for pullulanase activity. Fractions with high activity were pooled and dialyzed against medium B (20 mm Mes, pH 6.0, and 5 mm DTT). The dialyzed sample was applied at a flow rate of 0.25 mL min−1 to a column (10 cm high, 1.5 cm internal diameter) of cyclohexa-amylose-Sepharose (prepared from epoxy-activated Sepharose 6B, according to the method of Vretblad [1974]) equilibrated with medium B. The column was washed with medium B, eluted with 80 mL of 20 mm Hepes, pH 7.9, 0.5 m KCl, and 5 mm DTT, and 2-mL fractions were collected.

Partial Purification of Isoamylase

All steps were carried out at 0°C to 4°C. About 400 g of developing embryos (individual embryos of 400–600 mg fresh weight) or about 250 g of embryos on the 3rd d of germination was homogenized in a blender in 600 mL of medium D (50 mm Mes, pH 6.0, 10 mm calcium acetate, 5 mm DTT, and 50 mL L−1 ethanediol). The homogenate was centrifuged at 10,000g for 15 min, and the supernatant was adjusted to pH 5.0 by the addition of 1.2 m acetic acid with stirring. Precipitated protein was collected by centrifugation, resuspended in 80 to 100 mL of medium D, and then stirred for 16 h. This sample was applied at a flow rate of 0.5 mL min−1 to a Mono-Q anion-exchange column (Pharmacia) equilibrated with medium D. The column was washed with medium D and eluted with a 60-mL linear gradient of 0 to 0.6 m NaCl in medium D. One-milliliter fractions were collected, analyzed on native gels, and assayed for the production of reducing sugars from rabbit-liver glycogen (Sigma). Those with isoamylase activity were pooled and dialyzed against medium E (50 mm Mops, pH 7.0, 10 mm calcium acetate, 5 mm DTT, and 50 mL L−1 ethanediol). The dialyzed sample was reapplied to a Mono-Q column equilibrated with medium E. Elution and assay conditions were identical to those used for the first Mono-Q column except that medium E was used throughout. Fractions were analyzed on native gels and assayed for the production of reducing sugars from glycogen.

Determination of pH Optima

The pH optima of purified and partially purified enzymes were determined by assay at five pH values between 5.5 and 8.0. Buffers were Mes (pH 5.5–6.6), Mops (pH 6.6–7.2), and Hepes (pH 7.2–8.0).

Preparation of Plastids

Plastid-enriched fractions were prepared from samples of about 40 developing embryos according to the method of Denyer and Smith (1988) for crude plastid preparations. Embryos were from a line lacking one isoform of starch-branching enzyme (BC1/rr; Bhattacharyya et al., 1990) but otherwise isogenic to the line used in other experiments (BC1/RR). The low starch content of the rr line facilitates the preparation of plastids. No differences were observed between developing embryos of the RR and rr lines in the pattern or intensity of bands on amylopectin- or Red Pullulan-containing gels. Marker enzymes for plastids (ADP-Glc pyrophosphorylase, EC 2.7.7.27) and cytosol (alcohol dehydrogenase, EC 1.1.1.1) were assayed according to the method of Denyer and Smith (1988).

Gel Electrophoresis

SDS-PAGE was performed according to the method of Laemmli (1970) on 6% or 7.5% acrylamide gels 1.0 or 0.75 mm thick. Native gel electrophoresis was carried out in the same way except that SDS was omitted from all solutions. The separating gel contained potato amylopectin (Sigma) at 2 g L−1. After electrophoresis, native gels were rinsed in medium C (100 mm Mes, pH 6.0, 5 mm DTT, and 50 mL L−1 ethanediol), incubated in this medium for 2 to 4 h at 37°C, and then stained with a solution of 13 mm iodine and 40 mm KI. To inhibit amylase activity in gels, 0.1 mg mL−1 aplanin (BAY e4609, Bayer AG, Wuppertal, Germany) and 0.1 mg mL−1 acarbose were included in the separating gel and in the incubation medium. The incubation medium also contained 5 mm EDTA. Aplanin and acarbose were kind gifts from Bayer AG.

Proteins from identical amylopectin gels were transferred by electroblotting at 36 V for 2 h onto native gels containing Red Pullulan at 10 g L−1 according to the method of Wegrzyn and MacRae (1995). Gels were then incubated in medium C for 6 to 16 h at 37°C until the bands were visible.

Protein Assay

Protein was assayed with the Bio-Rad Protein Assay Dye Reagent with a standard curve of BSA.

RESULTS

Identification and Localization of Debranching Enzymes

We used native polyacrylamide gels containing amylopectin to detect debranching enzymes of both types in a nonquantitative manner. Incubation of gels at an appropriate pH after electrophoresis followed by staining with iodine solution revealed debranching activities as blue bands against a brown background. To distinguish debranching enzymes capable of hydrolyzing pullulan, proteins were transferred by electroblotting from amylopectin-containing gels onto gels containing Red Pullulan, on which pullulan-hydrolyzing debranching enzymes appear as clear bands. We also measured the capacity of embryo extracts to release reducing sugars (pullulan is not a substrate for other starch-hydrolyzing enzymes) and dye from Red Pullulan. Assay by release of reducing sugars is subject to interference by enzymes capable of hydrolyzing the products of the action of debranching enzyme on pullulan (maltotriose and maltotetraose). Isoamylase cannot be measured in crude extracts because its substrates are also substrates for amylases and/or limit-dextrinase. Because we identified debranching enzymes according to their ability to use pullulan as a substrate, we use the term “pullulanase” for isoforms that can use this substrate and “isoamylase” for isoforms that cannot.

Electrophoresis of crude, soluble extracts of developing pea embryos on native gels containing amylopectin revealed three main groups of bands that stained blue with iodine solution (Fig. 1, groups I, II, and III; A and B show typical results from two separately grown batches of plants). Other bands visible on these gels were either clear or stained pinkish-brown or reddish. Group I (usually a single band) and group II bands were always clearly visible, but group III bands were relatively faint and not easily distinguished in some preparations. To determine whether these bands were likely to be pullulanases or isoamylases, proteins were transferred immediately after electrophoresis to native gels containing Red Pullulan, which revealed clear bands in positions corresponding to groups II and III, but no bands corresponding to the slower-migrating group I (Fig. 1, panels at right). Therefore, it seemed likely that the group II and group III bands represented pullulanases, whereas the group I band represented isoamylase. On a fresh-weight basis, group I bands changed little in intensity during embryo development, but group II and group III bands increased in intensity (Fig. 1).

Figure 1.

Figure 1

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Isoforms of debranching enzymes in developing pea embryos. A and B show results from two independent batches. Crude, soluble extracts of developing embryos of the fresh weights (in milligrams) indicated above the lanes were loaded onto duplicate amylopectin-containing native gels so that each lane contained material from approximately 2.5 mg fresh weight. After electrophoresis one gel was incubated at 37°C for 2 h at pH 6.0, and then stained with iodine solution (left panels), and the other was electroblotted onto a Red Pullulan-containing native gel that was then incubated at 37°C for 16 h at pH 6.0 (right panels). The positions of three bands or groups of bands that stained blue on amylopectin-containing gels are labeled as I, II, and III.

To determine whether the putative debranching enzymes were plastidial, crude homogenates of developing embryos were separated into plastid-enriched pellets and supernatant fractions by established techniques (Denyer and Smith, 1988). When gels of homogenate, supernatant, and plastid-enriched fractions were loaded so that each lane contained an equal activity of the plastidial enzyme ADP-Glc pyrophosphorylase, the putative isoamylase band (group I) and the faster migrating of the two pullulanase bands in group II were present at approximately equal intensities in all of the lanes. However, the slower-migrating pullulanase band in group II and the group III bands were visible only in supernatant and homogenate lanes and not in lanes containing plastid-enriched fractions. Figure 2 shows typical results from young embryos (Fig. 2A) and older embryos (Fig. 2B). The group III bands were visible only in the latter. These results suggest that most or all of the putative isoamylase and the faster-migrating pullulanase in group II are plastidial, whereas the slower-migrating pullulanase in group II and the group III pullulanases are extraplastidial. The results also indicate that the ratio of extraplastidial to plastidial pullulanase activity may increase during development. On Red Pullulan-containing gels the slower-migrating pullulanase in group II and the group III pullulanases were much more prominent in homogenates of older than younger embryos, but the faster-migrating pullulanase in group II was not.

Figure 2.

Figure 2

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Intracellular localization of isoforms of debranching enzymes in developing pea embryos. A, Preparation from young embryos of approximately 200 mg fresh weight. B, Preparation from older embryos of approximately 400 mg fresh weight. Plastid-enriched (P) and supernatant (S) fractions were separated by centrifugation from homogenates (H) of developing embryos. Fractions were loaded onto duplicate amylopectin-containing gels so that each lane contained the same activity of the plastidial enzyme ADP-glucose pyrophosphorylase. After electrophoresis, one gel was incubated and then stained with iodine (left panels) and the other was electroblotted onto a Red Pullulan-containing native gel (right panels) (for details, see legend to Fig. 1). Red Pullulan-containing gels were treated with ethanol after incubation to enhance the contrast between the bands and the background. The positions of group I, group II, and, where visible, group III bands are indicated.

The intracellular location of pullulanase was further investigated by assay of the enzyme in homogenate and plastid-enriched fractions of developing embryos. Total pullulanase activity, measured either by release of reducing sugars from pullulan or by release of dye from Red Pullulan, was low in the early stages of embryo development and increased during development (Table I and data not shown).

Table I.

Activity of pullulanase in developing, dry, imbibed, and germinating embryos

Stage Pullulanase Activity
μmol min−1 seed−1
Developing embryos (fresh wt)
 0.2 to 0.3 g 0.027  ± 0.006
 0.3 to 0.4 g 0.071  ± 0.011
 0.45 to 0.55 g 0.104  ± 0.006
Dry embryos 0.109  ± 0.003
Imbibed embryos
 1 DAIa 0.135  ± 0.005
 3 DAI 0.115  ± 0.006
 5 DAI 0.145  ± 0.007
 7 DAI 0.134  ± 0.005
 9 DAI 0.095  ± 0.005
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Values are means ± se of six measurements, each made on a separate extract of a single seed. Pullulanase activity was assayed by the reducing-sugar method.

a

DAI, Days after imbibition. 

The low activity in the early stages of development was not attributable to the presence of inhibitory substances in extracts of young embryos. In two separate experiments in which old (about 500 mg fresh weight) and young (100–200 mg fresh weight) embryos were co-extracted, activity in the mixed extract was 88% and 89%, respectively, of that predicted from separate extractions of the two kinds of embryo when assayed by the reducing-sugar method, and 81% and 83%, respectively, of that predicted when assayed by the Red-Pullulan method.

Plastid-enriched fractions were prepared from homogenates of embryos of about 200 mg fresh weight. The percentage of pullulanase activity in the plastid-enriched fraction was greater than that of the cytosolic marker enzyme alcohol dehydrogenase, but considerably less than that of the plastidial marker enzyme ADP-Glc pyrophosphorylase (Table II). We estimate from these data that 17% of the pullulanase activity is plastidial in embryos of about 200 mg fresh weight (calculation described by Denyer et al., 1996).

Table II.

Intracellular localization of pullulanase activity

Enzyme Activity in Plastid-Enriched Fraction Recovery of Activity
%
ADP-Glc pyrophosphorylase 19.8  ± 2.0 98.3  ± 4.2
Alcohol dehydrogenase 0.14  ± 0.02 106.1  ± 3.4
Pullulanase 3.55  ± 0.68 90.2  ± 7.1
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Activities of pullulanase, the plastidial enzyme ADP-Glc pyrophosphorylase, and the cytosolic enzyme alcohol dehydrogenase were measured in a homogenate of young embryos (approximately 200 mg fresh weight), in a plastic-enriched pellet fraction, and in a supernatant fraction derived from the homogenate by centrifugation. The activity in the plastid-enriched fraction is expressed as a percentage of that in the homogenate. The recovery of activity during the fractionation is calculated by expressing the sum of the activities in the supernatant and plastid-enriched fractions as a percentage of that in the homogenate. The high values indicate that no losses of activity occurred during the fractionation. Values are means ± se of measurements made on six separate preparations.

Purification of Pullulanase from Developing Embryos

Pullulanase was purified to near homogeneity from developing embryos by ammonium sulfate precipitation, anion-exchange chromatography, and affinity chromatography on cyclohexa-amylose-Sepharose. Activity was monitored during the purification as the production of reducing sugars from pullulan. Purification was approximately 1500-fold, to a final specific activity of 33 to 44 μmol min−1 mg−1 protein (range of values from three separate purifications). A typical purification is shown in Table III.

Table III.

Purification of pullulanase from developing embryos

Assay Total Protein Total Activity Specific Activity Purification Recovery
mg μmol min−1 μmol min−1 mg−1 protein -fold %
Initial supernatant 6464 166.8 0.0258 1 100
Ammonium sulfate 1115 61.7 0.0553 2.1 37
DEAE-Sepharose 249 32.4 0.130 5.0 19.4
CHA-Sepharose 0.106 4.14 39.1 1516 2.5
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Embryos (about 400 g, 400–600 mg fresh weight each) were homogenized and centrifuged to give an initial supernatant. A fraction precipitating at between 30% and 40% saturation with ammonium sulfate was subjected to further chromatography on columns of DEAE-Sepharose Fast Flow and cyclohexa-amylose (CHA)-Sepharose. Pullulanase activity was assayed by the reducing-sugar method. Recovery of activity is the activity remaining after a given step expressed as a percentage of that in the initial supernatant.

On SDS-polyacrylamide gels stained with Coomassie brilliant blue R, the purified preparation usually appeared as two bands of approximately 100 kD (Fig. 3, lane 2). On gels containing amylopectin, the purified pullulanase from pea embryos appeared as a blue-staining band that co-migrated with group II bands from crude extracts (Fig. 3, lanes 3–5). On gels containing Red Pullulan, the purified enzyme appeared as a clear band (Fig. 3, lane 6) that comigrated with the slower-migrating, extraplastidial group II pullulanase, and not with the faster-migrating, plastidial pullulanase (Fig. 3, lanes 7–9). In some preparations, bands that comigrated with the group III bands of crude extracts were also faintly visible. The purified enzyme had a pH optimum of 6.0, hydrolyzed pullulan very readily, and also hydrolyzed amylopectin; it had very little activity with glycogen (Table IV).

Figure 3.

Figure 3

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Purified pullulanase from developing embryos. Pullulanase activity was purified as described in Table III. Left panel, SDS-polyacrylamide gel. Middle panel, Amylopectin-containing native gel; the positions of group I and group II bands are marked. Right panel, Red Pullulan-containing native gel. Lane 1, Molecular markers (in kilodaltons); lanes 2, 3, and 6, purified pullulanase; lanes 4 and 7, plastid-enriched fraction from young embryos (as in Fig. 2); lanes 5 and 8, supernatant fraction from young embryos (as in Fig. 2); and lane 9, mixture of the samples in lanes 6 and 7. Note that on the Red Pullulan-containing gel the purified pullulanase comigrates with the upper, extraplastidial group II band in the supernatant fraction, and not with the plastidial group II band.

Table IV.

Substrate preferences of pullulanase and isoamylase

Preparation Release of Reducing Sugars
Amylopectin Glycogen Pullulan
%
Developing embryos
 Pullulanase 100 3.4 384
 Isoamylase 100 73 nda
Imbibed embryos
 Isoamylase 100 74 nd
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Release of reducing sugar by preparations of pullulanase (as in Table III) and isoamylase (as in Figs. 4 and 7) was measured with amylopectin, glycogen, and pullulan as substrates. Activity is expressed as a percentage of that in assays containing amylopectin. Substrate concentration in all assays was 20 g L−1. For a given enzyme preparation, all assays contained the same amount of enzyme protein. Activity was measured for 90 min, and was linear with respect to time during this period. Values are means of duplicate assays made on single preparations of each enzyme and are typical of values obtained from several preparations.

a

nd, Not detected. 

Purification of Isoamylase from Developing Embryos

The putative isoamylase was partially purified from developing embryos by precipitation at pH 5.0 followed by anion-exchange chromatography. As reported previously for the enzyme from potato (Ishizaki et al., 1983), activity was effectively precipitated at pH 5.0, whereas the activity of other starch-hydrolyzing enzymes was not. Activity of the putative isoamylase was monitored throughout the purification by analysis on native gels containing amylopectin. Fractions from the final Mono-Q column gave blue-staining bands that comigrated with the group I band of crude extracts and contained an activity that released reducing sugars from glycogen. They gave no bands on gels containing Red Pullulan (Fig. 4). SDS-polyacrylamide gels revealed that each of the active fractions contained several proteins (not shown).

Figure 4.

Figure 4

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Partially purified isoamylase from developing embryos. Activity was purified by precipitation at pH 5.0 from crude, soluble extracts of embryos, followed by chromatography on Mono-Q anion-exchange columns. Top panel, Elution of glycogen-hydrolyzing activity (micromoles of reducing sugar produced per minute per fraction) from the final Mono-Q column. NaCl concentration increased linearly from 0.23 to 0.34 m across the fractions shown. Middle panel, Amylopectin-containing native gel of unfractionated extract (left lane) and fractions from the Mono-Q column shown in the top panel. The lower of the two bands in fractions 23, 24, and 25 stained reddish-brown with iodine solution; all other bands stained blue. Bottom panel, Red Pullulan-containing native gel of unfractionated extract (left lane) and of fractions from the Mono-Q column shown in the top panel.

Further evidence about the identity of the purified activity was sought from studies of its substrate preferences. The activity could release reducing sugars from β-limit-dextrin, but to a lesser extent than pancreatic α-amylase. In a typical experiment in which the substrate was digested to completion, the amount of reducing sugar produced by the putative isoamylase from β-limit-dextrin was 23% of that produced by α-amylase from the same amount of substrate. The putative isoamylase did not release Glc from maltoheptaose, a linear glucan of seven glucosyl units from which α-amylases can release Glc. Its activity with glycogen was slightly less than that with amylopectin, and it had little or no activity on pullulan (Table IV). When allowed to digest amylopectin to completion, the products of the putative isoamylase were much more similar to those of bacterial isoamylase than to those of pancreatic α-amylase. Fractionation of the products by high-performance anion-exchange chromatography with pulsed amperometric detection revealed that although the putative isoamylase produced more chains of fewer than six glucosyl units than did the bacterial isoamylase, both enzymes produced a range of longer chains with a maximum abundance at about 12 to 15 glucosyl units. In contrast, pancreatic α-amylase produced almost exclusively Glc and chains of six or fewer glucosyl units in length (Fig. 5). The activity of the putative isoamylase was maximal at pH 7.0.

Figure 5.

Figure 5

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Products of hydrolysis of amylopectin by partially purified isoamylase. A, Amylopectin after digestion with bacterial isoamylase. B, Amylopectin after digestion with partially purified isoamylase from pea embryos (25 nmol min−1 incubation−1 from the final Mono-Q column, as in Fig. 4). C, Amylopectin incubated under the same conditions as in B, but without enzyme activity. D, Amylopectin after digestion with pancreatic α-amylase. After digestion of the amylopectin to completion, the products were fractionated by high-performance anion-exchange chromatography and visualized with a pulsed amperometric detector. The system was calibrated with maltooligosaccharides of known degrees of polymerization. Peaks with degrees of polymerization of 10 (A and B) and 6 (D) are indicated.

Pullulanases in Mature and Germinating Embryos

When expressed per embryo, the activity of pullulanase was similar in embryos in late development, dry embryos, and embryos during the first 9 d of germination (Table I). The general pattern of isoforms also differed little between these stages. Red Pullulan-containing native gels revealed the presence of both group II and group III bands in dry, imbibed, and germinating embryos (Fig. 6).

Figure 6.

Figure 6

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Debranching enzymes in developing, dry, imbibed, and germinating embryos. A, Amylopectin-containing native gel (prepared as described for Fig. 1) of crude, soluble extracts of embryos. Each lane contains material from approximately 1/100th of an embryo. Lane 1, Developing embryo of 500 mg fresh weight; lane 2, dry embryo; lane 3, embryo after imbibition; lane 4, embryo after 2 d of germination; lane 5, embryo after 4 d of germination. Group I and group II bands are indicated; these are partly or wholly obscured by bands of amylase activity. B, Amylopectin-containing native gel identical to that in A, except that α-amylase inhibitors were included in the gel and incubation medium. Group I and group II bands are indicated. The differences between lanes in the composition of the samples causes slight differences in the migration of the group I band; this band is indicated on both sides of the panel and is the upper of the prominent doublet of bands in all lanes. C, Red Pullulan-containing native gel of samples electroblotted from a gel identical to that shown in A. Group II and group III bands are indicated.

Isoamylase in Mature and Germinating Embryos

The presence of large amounts of slow-migrating, amylopectin-hydrolyzing activities (probably α-amylases) prevented debranching enzymes from being resolved clearly on amylopectin-containing gels of germinating embryos. Incorporation into the incubation mixture of the α-amylase inhibitors aplanin, acarbose, and EDTA strongly reduced the development of these bands (Fig. 6). The inhibitors had no effect on the appearance of the group I band on gels of developing embryos, showing that they did not inhibit isoamylase. In gels treated with inhibitors, a blue-staining band that comigrated with the group I band of developing embryos was visible in extracts of mature and germinating embryos (Fig. 6).

To provide further information about the identity of the putative isoamylase in mature, imbibed, and germinating embryos, extracts of imbibed embryos were subjected to the purification procedure devised for isoamylase from developing embryos. The final preparation from imbibed embryos was similar to that from developing embryos. Fractions from the final Mono-Q column appeared exclusively as slow-migrating, blue-staining bands on amylopectin-containing gels, and contained an activity that released reducing sugars from glycogen. No bands were visible on Red Pullulan-containing gels of these fractions (data not shown). The substrate preferences of the preparation were indistinguishable from those of the isoamylase from developing embryos (Table IV), and the pH optimum for activity was also the same (7.0).

DISCUSSION

Pullulanases of Developing and Germinating Embryos

Our data show unequivocally that developing pea embryos contain two distinct isoforms of debranching enzyme capable of hydrolyzing both amylopectin and pullulan. At least one of these forms is plastidial, and is present in the embryo from an early stage of development and probably throughout the period of starch synthesis. Both native gel analysis and subcellular fractionation experiments indicate that the plastidial pullulanase contributes a minor fraction of the total pullulanase activity. The remaining pullulanase activity is extraplastidial, and appears to increase relative to the plastidial activity as development proceeds. The extraplastidial activity forms three or more bands on native gels. Purified extraplastidial pullulanase appears as two bands on SDS-polyacrylamide gels, and it is possible that both of these bands are pullulanases. Purified pullulan-hydrolyzing activity from mature mung beans also migrated as two bands of approximately 100 kD on SDS-PAGE, and production of monoclonal antisera revealed that the two bands were immunologically very closely related (Morinaga et al., 1997). However, our results do not exclude the possibility that products of a single gene account for all of the extraplastidial activity.

The isoforms of pullulanase present in developing embryos are probably also responsible for the pullulanase activity in germinating embryos. The appearance of pullulanase bands on native gels is the same in late development, in dry seed, after imbibition, and in germination. Measurement of pullulanase activity by the reducing-sugar method indicates that there is little difference in activity per embryo between these developmental stages. However, these results and those used to quantify the proportion of pullulanase in the plastid must be treated with some caution. Native gels cannot be regarded as quantitative, and the assay for pullulanase is susceptible to interference by enzymes that can hydrolyze maltotriose and maltotetraose. Pullulan is not an endogenous substrate for plant enzymes.

The presence of plastidial and extraplastidial isoforms of pullulanase (limit-dextrinase, R-enzyme) has been reported in leaves of three species, but not in other plant organs. Cell-fractionation studies showed that approximately one-half of the activity of spinach (Okita et al., 1979), sugar beet (Li et al., 1992), and broad bean (Ghiena et al., 1993) leaves is chloroplastic. Studies of other organs have for the most part identified only one activity, for example in germinating barley (Sissons et al., 1992; MacGregor et al., 1994); developing and germinating rice endosperm (Iwaki and Fuwa, 1981; Nakamura et al., 1996a); maize endosperm (Doehlert and Knutson, 1991); and broad bean seeds (Gordon et al., 1975).

In the most detailed study to date, a single form of pullulanase was purified from developing rice endosperm and the cDNA encoding it was isolated and sequenced. The cDNA encoded a putative transit peptide, indicating that the isoform is plastidial. Southern-blot analysis using the cDNA as a probe suggested that there may be only a single pullulanase gene in the rice genome (Nakamura et al., 1996a). In maize three forms of pullulanase were separated by hydroxyapatite chromatography of the partially purified enzyme from developing endosperm (Pan and Nelson, 1984). However, the number of different gene products responsible for these activities and their intracellular locations were not investigated.

The properties of the extraplastidial pullulanase purified from developing pea embryos are broadly similar to those of pullulanases from other plant sources. The purified protein(s) is approximately 100 kD, a value very similar to those reported for most of the pullulan-hydrolyzing enzymes characterized so far (Nakamura, 1996). The pea enzyme hydrolyzes pullulan more readily than amylopectin and displays little or no detectable activity with glycogen. This order of substrate preference is the same as that of the enzymes from, for example, spinach chloroplasts (Okita and Preiss, 1980; Ludwig et al., 1984), sugar beet leaves (Li et al., 1992), mature broad bean seeds (Gordon et al., 1975), and developing rice endosperm (Nakamura et al., 1996a). The properties of our pea enzyme are also very similar to those of a limit-dextrinase purified by ammonium sulfate and acetone fractionation and cyclohexa-amylose chromatography from imbibed pea seeds (Yellowlees, 1980). It seems likely that this limit-dextrinase and the extraplastidial pullulan-hydrolyzing activity from developing embryos are the same enzyme.

The Isoamylase of Developing and Germinating Embryos

In addition to pullulanases, native gel analysis shows that developing pea embryos contain a second type of debranching enzyme that is capable of hydrolyzing amylopectin but not pullulan. Most or all of this activity is plastidial and is present during development. The following characteristics of partially purified preparations of this enzyme suggest strongly that it is an isoamylase. First, the enzyme does not release reducing sugars from pullulan or Glc from maltoheptaose, showing that it is not pullulanase, α-amylase, α-glucosidase, or disproportionating enzyme. Second, the enzyme liberates reducing sugars from amylopectin, β-limit-dextrin, and glycogen, all of which are substrates for isoamylases (Nakamura, 1996; Manners, 1997). β-Limit-dextrin is not a substrate for β-amylase. Third, the action of the enzyme on amylopectin produces a pattern of Glc chains similar to that produced by bacterial isoamylase, and very different from that produced by α-amylase. The enzyme preparation from pea produced more short chains than did bacterial isoamylase (Fig. 5). Further purification of the enzyme will be required to determine whether this is a real difference between the two isoamylases, or if it was caused by contaminating activities in the pea preparation.

Dry, imbibed, and germinating pea seeds contain an isoamylase activity that is indistinguishable in behavior on native gels, substrate specificity, and pH optimum from the isoamylase from developing embryos.

There is only limited information on the properties of isoamylase in the two plant organs from which it has been reported previously. A single form of the enzyme purified from potato tuber hydrolyzed glycogen and amylopectin at an almost equal rate (Ishizaki et al., 1983). One of two forms of the enzyme separated by anion-exchange chromatography from developing maize kernels hydrolyzed amylopectin about four times faster than phytoglycogen, and the other form was not characterized (Doehlert and Knutson, 1991). Activity of isoamylase in developing maize endosperm appears to be mainly or exclusively plastidial (Yu et al., 1998).

Possible Roles of Debranching Enzymes in Starch Metabolism in the Pea Embryo

The most commonly described role for starch-debranching enzymes is in the degradation of starch. Limit-dextrinases are so called because their substrate in vivo is held to be limit-dextrins generated during hydrolytic starch degradation (Manners, 1997). Their presence in the acellular endosperm of germinating cereals is certainly consistent with such a role, and we assume that debranching enzymes also participate in starch degradation in the cells of the germinating pea embryo. It has been proposed on the basis of electron micrographs that the plastid membranes surrounding the large starch granules break down during the germination of legume embryos (Bain and Mercer, 1966; Harris, 1976), which would allow cytosolic rather than plastidial isoforms of debranching enzyme to participate in starch degradation. The extraplastidial pullulanase may therefore be responsible for hydrolysis of the α1,6-linkages of starch during germination.

A role for the plastidial isoform of pullulanase and for isoamylase in the germinating embryo remains to be discovered. A situation that may be similar to that of debranching enzymes has been described for isoforms of starch phosphorylase in the germinating pea embryo. An isoform of phosphorylase revealed by immunofluorescence microscopy to be cytosolic contributes most of the activity of phosphorylase, and a plastidial isoform is confined to the small plastids, where it presumably cannot participate in starch degradation (Steup, 1988; van Berkel et al., 1991).

The role of debranching enzymes in the developing embryo is unclear. It might be argued that the extraplastidial pullulanase is required for starch degradation during germination, and that its presence during development of seeds simply represents accumulation in advance of this role. The fact that limit-dextrinases of some developing and mature cereal endosperms are reported to be present in a “latent” form, requiring reduction to achieve full activity (Yamada, 1981; Toguri, 1991; Sissons et al., 1993; MacGregor et al., 1994), lends support to this view. However, the presence of plastidial isoamylase and pullulanase from an early stage of development of the pea embryo suggests to us that these enzymes are likely to have a role in starch metabolism during its synthesis in the developing embryo.

The demonstration that both pullulanase and isoamylase occur in the plastids of developing pea embryos as well as in potato tubers and maize endosperm, and the dramatic effect of mutations affecting these enzymes on starch synthesis in organisms as distantly related as C. reinhardtii and cereals, are consistent with a fundamental role for debranching enzymes in starch synthesis. The precise roles of the two kinds of enzyme remain to be discovered.

ACKNOWLEDGMENTS

We are grateful to Sam Zeeman, Takayuki Umemoto, Cathie Martin, Rod Casey, Kay Denyer, and Professor David Manners for their helpful comments during the course of this work and about the manuscript, and to Cliff Hedley and Trevor Wang for the gift of pea seeds.

Footnotes

1

This research was supported by a competitive strategic grant from the Biotechnology and Biological Sciences Research Council (UK) to the John Innes Centre, by an International Collaboration Project funded by the European Union (contract no. CI1*/0417/00), by funding from the Chinese National Natural Foundation of Science to Z.-P.Z., and by an Erasmus Award (European Union) to U.R.

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