Enolase Isozymes from Developing Oilseeds: A Survey

Enolase Isozymes from Developing Oilseeds: A Survey!)

Biology Department, Queen's University, Kingston, Ontario K7L3N6, Canada Received September 29, 1986 . Accepted October 31, 1986

Summary Enolase activity was used to demonstrate the occurrence of both plastid and cytosolic glycolysis in the developing storage organs of several oil-rich seeds. Plastids isolated from these tissues and purified by rate-zonal sedimentation on discontinuous sucrose gradients contained substantial amounts of enolase activity. The enolase activity associated with isolated plastids could be separated from that found in the cytosol by ion-filtration and ion exchange chromatography and could be attributed toa distinct isozyme. Hence, plastid glycolysis in developing oilseeds appears to be universal.

Key words: Ricinus, enolase, glycolysis, isozymes, oilseeds, plastids.

Introduction

Endosperm cells of developing castor oil seeds contain two discreet, spatially separate glycolytic pathways, one in the cytosol and the other located within the plastids (reviewed by Dennis and Miernyk, 1982). Cytosolic glycolysis is a component of the classical respiratory catabolism of hexoses, whereas plastid glycolysis provides pyruvate for fatty acid biosynthesis. It has been shown that each of the enzymes of the plastid glycolytic pathway is an isozyme that can be separated from the activity that is found in the cytosol (DeLuca and Dennis, 1978; Simcox and Dennis, 1978; Garland and Dennis, 1980 a; Miernyk and Dennis, 1982). For several enzymes in Ricinus, the glycolytic isozymes have been purified and characterized (Miernyk and Dennis, 1984; Ireland et aI., 1980; Garland and Dennis, 1980 b). We have previously reported the occurrence of two isozymes of enolase from developing castor oil seeds, and shown one to be located in the plastid and one in the cytosol (Miernyk and Dennis, 1982). Both isozymes have been purified and characterized (Miernyk and Dennis, 1984). Previous data from this laboratory have indicated that plastid and cytosolic glycolysis is present in many plant cells (Ireland et aI., 1979; Ireland and Dennis, 1980). However, the presence of glycolytic activities in plastids of certain tissues has been questioned (e.g. Stitt and ap Rees, 1979). In this report, we show that the presence of a plastid isozyme of enolase can be used to determine the extent of plastid glycolysis in tissues other than Ricinus. 1) Supported by the Natural Sciences and Engineering Research Council of Canada. 2) Present address: Seed Biosynthesis Research Unit, The Northern Regional Research Center, USDA/ARS, 1815 N. University Street, Peoria, IL 60604, U.S.A.

3) Author to whom reprint requests should be addressed.

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Materials and Methods Watermelon, cucumber, and sunflower plants were field grown, the immature fruits harvested, and developing seeds removed. All other plants were glasshouse-grown as previously described (Miernyk and Dennis, 1982). Developing seeds were selected at approximately midpoint in storage lipid accumulation (e.g. 21 days after pollination for corn, 30 days for castor oil seeds, 34 days for cotton). Enolase (2-phospho-D-glycerate hydro-lyase, EC4.2.1.11), and organelle marker enzymes were assayed by previously established procedures, as were absorbance, refractive index, and conductivity (Miernyk and Dennis, 1982; Bortman et aI., 1981; Simcox et al., 1977). One unit of activity is equal to one /Lmol min - 1. Methods of homogenization and centrifugation have been described previously (Miernyk and Dennis, 1982). Following clarification of homogenates, low molecular weight compounds were removed by centrifugation of aliquots through columns of Sephadex G-25 (2.0 ml bedvolume), previously equilibrated with homogenization buffer, and mounted in clinical centrifuge tubes. Isolation of plastid-enriched fractions by rate-zonal sedimentation was according to the method of Simcox et ai. (1977). Purification of plastids by rate-zonal sedimentation on discontinuous sucrose gradients was as described by Dennis and Green (1975). Separation of the enolase isozymes by ion-filtration chromatography (Kirkegaard et aI., 1972) using Sephadex A-25 has been described previously (Miernyk and Dennis, 1982). In some instances, smaller preparations were analysed by ion-exchange chromatography, essentially as described by Garland and Dennis (1980 a) but using the column equilibration and elution conditions of Miernyk and Dennis (1982). The single exception was with developing soybean seeds where greater resolution was obtained with an equilibration pH of 6.9 rather than the usual 6.7. Substrates, cofactors, buffers, and auxilIary enzymes were from the Sigma Chemical Company (St. Louis) or Boehringer-Mannheim (Montreal). Imidazole was purified as previously described (Miernyk and Dennis, 1982). Chromatographic matrices were from Pharmacia.

Results

The methodology developed for the purification of plastids from developing Ricinus endosperm could also be used for the purification of plastids from all other developing oilseeds that were examined, with the exception of soybeans. A typical gradient profile (from developing sunflower cotyledons) is presented in Fig. 1. It was possible to prepare a plastid-enriched fraction from developing soybean cotyledons by rate-zonal sedimentation, but the organelles appeared to aggregate, preventing further purification by sedimentation on a discontinuous sucrose gradient. In most instances, the enolase activity in total homogenates could be resolved into two discreet peaks by ion-exchange or ion-filtration chromatography. A typical profile (from developing watermelon cotyledons) is presented in Fig. 2 A. Fractions enriched in plastid or cytosolic components were analyzed by the same procedures (Figs. 2 B, C). In all instances, the plastid isozymes eluted from the columns at lower conductivities than the cytosolic isozymes. With homogenates from developing cotton (Fig. 3) and flax cotyledons, three peaks of enolase activity were resolved. In both instances the plastid fractions were enriched in a single isozyme which eluted at the lowest conductivity on the ion-exchange columns. The cytosol-enriched fractions contained the isozyme which bound most tightly.

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Fig. 1: Separation of organelles from developing sunflower cotyledons by rate-zonal sedimentation on a discontinuous sucrose gradient. Developing embryos (21 g) were homogenized and a plastid-enriched pellet was prepared and resuspended as described in Materials and Methods. This was layered onto a gradient consisting of 6 ml of 60 %, 16 ml of 45 % and 13 ml of 35 % sucrose, and centrifuged at 100,000g for 20 min, using a Beckman SW-28 rotor. In order to calculate actual enzyme activities in /Lmol min - 1 fraction -1, ordinate values should be multiplied by: 0.22 for alcohol dehydrogenase (ADH) (0); 0.30 for citrate synthase (CS) (e); 19.40 for triose-phosphate isomerase (TPI) (0) and 1.03 for enolase (A). The activity of triose-phosphate isomerase in fractions 30 through 36 was 73 % of that applied to the gradient.

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Fig. 2: Ion-exchange chromatography on DEAE-Sephacel of a clarified homogenate (A); a purified plastid fraction (B); and a cytosol-enriched fraction (C), prepared from developing watermelon embryos and assayed for enolase activity. In each case approximately 1.8 g of tissue was used. The column equilibration buffer was 20mM imidazole, pH 6.7, containing 2 mM MgCb. Actual enzyme activities in JLmol min - I fraction -1 can be obtained by multiplying ordinate values by: 0.29 (A); 0.38 (B); or 0.27 (C). Recoveries following chromatography were 111 % (A); 99% (B); and 94% (C).

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Fig. 3: Ion-filtration chromatography using Sephadex A-25 of: a clarified homogenate (A); a purified plastid fraction (B); and a cytosol-enriched fraction (C), prepared from developing cotton embryos and assayed for enolase activity. In each case, approx. 10.5g of 34-day old embryos were used. Column equilibration buffer was 20 mM imidazole, pH 6.7, containing 10 mM MgCb. Actual enzyme activities in /Lmol min ~ 1 fraction ~ 1 can be obtained by multiplying ordinate values by: 1.01 (A); 0.51 (B); or 1.43 (C). Recoveries following chromatography were: 105 % (A); 101 % (B) and 89 % (C).

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Table 1: Localization of the enolase isozymes from developing oilseeds. Species

Organ

Ricinus communis L. cv. Baker 296 endosperm (castor oil seed) Brassica napus L. cotyledons (rape) Heliantbus annus L. cv. giant Russian cotyledons (sunflower) Citrullus vulgaris Schrad. cotyledons (watermelon) Cucumis sativus L. cotydedons (cucumber) Glycine max. L. cotyledons (soybean) Linum usitatissimum L. cotyledons (flax) Gossypium birsutum L. cv. Deltapine 16 cotyledons (cotton) Zea maize L. cv. W64A embryo b) Schroeder et al., 1974. ") Norton and Harris, 1983.

Mature Seed Total Distribution (%) Lipid Content Enzyme % Dry Weight Activity Plastid Cytosol (/Lmol min -I gfw-I) 56

8.37

33

67

50")

3.40

17

83

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35.88

29

71

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9.79

11

89

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3.61

20

80

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2.35

14

86

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3.74

27

73

19

5.26

37

63

4.90

32

68

25

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C) Tan and Morrison, 1979.

The proportions of enolase aCtIVlty localized in each subcellular compartment (Table 1) were estimated from ion-exchange or ion-filtration chromatographs. Recoveries from the columns were between 80 and 115 %, and it was assumed that there was no preferential loss of either isozyme. The proportion of plastid enolase based upon chromatographic separation was in good agreement with that calculated from organelle isolation after correction for the breakage of organelles during homogenization. Discussion Multiple forms of the glycolytic enzymes have been reported in bacteria, eucaryotic microbes, plants, and animals. In microbial systems, expression of specific glycolytic isozymes may be related to external carbon nutrition (Hess et aI., 1968; Entian et aI., 1984). In mammalian systems, glycolytic isozyme expression can be tissuespecific (Vandeberg et aI., 1975; Marangos et aI., 1982; Tanaka et aI., 1967) or related to development state (Decker and Mohrenweiser, 1981; Ureta, 1982). In contrast, the differential expression of the plant glycolytic isozymes is related to the functions of different subcellular locations within the cell (Dennis and Miernyk, 1982). Enolases from all sources so far examined occur as dimers (Wold, 1971). Three types of enolase subunits are found in mammalian tissues (Marangos et al., 1978; Shimizu et al., 1983). In neural tissues there are three isozymes with subunit compositions of aa, (3(3, and 'Y'Y. A a'Y-heterodimer also appears to form in vivo (Shimizu et aI., 1983). Three enolase isozymes are also found in yeast extracts (Entian et aI., 1983). However, Holland et al. (1981) showed that Saccharomyces cerevisiae cells contain

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only two enolase structural genes per haploid genome. The third isozyme is a heterodimer and is composed of one isozyme-1 and one isozyme-2 subunit (McAlister and Holland, 1982). Whether the heterodimer occurs in vivo is controversial. However, the presence of isozymes is not essential since a deletion mutant constructed in vitro, that had a single homodimeric enolase, grew normally on glucose or ethanol (McAlister and Holland, 1982). Two enolase isozymes were found in most of the developing oilseeds that were surveyed, one of which was located in the plastids and the other in the cytosol. In cotton and flax preparations three isozymes were found. The subcellular location of the isozymes of intermediate charge was not resolved. It is not known whether the three isozymes are encoded by discreet structural genes (Gottlieb, 1982), are the result of post-transcriptional or post-translational processing, or if heterodimers of plastid and cytosolic subunits can be formed. Gottlieb and associates have reported that heterodimer formation between plastid and cytosolic subunits is uncommon (e.g. Weeden and Gottlieb, 1982), although it has recently been observed with plastid and cytosolic triose-phosphate isomerase isozymes (Pichersky et aI., 1984). Developmental studies suggest that, in the case of cotton, the isozyme of intermediate charge is a heterodimer of one cytosolic and one plastid subunit that is formed during tissue disruption A. Miernyk, unpublished). Structural studies with homogenous subunits will be required for a definitive answer to this question. The high activities of plastid enolase observed in preparations from developing oilseeds probably reflects the high demand for pyruvate for fatty acid synthesis in these oil-rich tissues. However, there is no correlation between the amount of storage lipid accumulated and the level of plastid enolase activity (Table 1), but in all cases the level of plastid enolase activity is in excess of that required for the in vivo rate of storage lipid accumulation (e.g. Simcox et al., 1977). The de novo synthesis of fatty acids in plant cells occurs entirely within the plastids (Ohlrogge et aI., 1979), and it is logical that the two-carbon precursors of fatty acids should be provided by plastid glycolysis (Dennis and Miernyk, 1982) and the plastid pyruvate dehydrogenase complex (Reid et aI., 1977; Camp and Randall, 1985). Fatty acids are also required as structural components of membrane lipids and, hence, plastid glycolysis would be expected to occur even in tissues which do not accumulate storage lipids. It was recently observed that plastids from corn endosperm contain all of the glycolytic enzymes (Miernyk, Echieverria, and Shannon, unpublished) even though this tissue does not store lipids. The plastid enolase occurs as a distinct isozyme in this tissue. The above results suggest that plastid glycolysis occurs in all plant tissues at some stage of development. The plastid isozyme of enolase can be used as a probe to determine the extent of glycolysis in this organelle.

a.

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