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Subcellular distribution of acetyl-coenzyme A carboxylase in mesophyll cells of barley and sorghum leaves
ARCHIVES
OF BIOCHEMISTRY
Vol. 235, No. 2, December,
AND
BIOPHYSICS
pp. 555-561,
1984
Subcellular Distribution of Acetyl-Coenzyme A Carboxylase Mesophyll Cells of Barley and Sorghum Leaves’ BASIL Department
J. NIKOLAII, of Biochemistry Received
EVE SYRKIN and Biophysics, April
WURTELE: University
27, 1984, and in revised
AND
PAUL
of CalZfornia,
Davis,
form
8, 1984
August
in
K. STUMPF4 California
95616
The subcellular distribution of acetyl-CoA carboxylase [acetyl-CoA-carbon dioxide ligase (ADP-forming), EC 6.4.1.21 was determined in mesophyll protoplasts isolation from barley, a C3 plant, and sorghum, a C4 plant. In both species, all of the mesophyll acetyl-CoA carboxylase was demonstrated to be chloroplastic. In barley leaves and mesophyll protoplasts, a single biotinyl protein of 60,000 Da was identified by a modified Western-blotting procedure. The subcellular distribution of this biotinyl protein was identical to that found for acetyl-CoA carboxylase. These results are discussed in relation to the compartmentation of reactions requiring malonyl-CoA as a substrate. (c 1984 Academic Pruss. Inc.
Acetyl-CoA carboxylase catalyzes the rate-limiting reaction in the biosynthesis of fatty acids in Escherichia coli, yeasts, and various mammalian tissues (1). In these organisms, malonyl-CoA, the product of the reaction catalyzed by acetylCoA carboxylase, is utilized only in the biosynthesis of fatty acids, and a correlation between acetyl-CoA carboxylase activity and fatty acid biosynthesis has been demonstrated. With the exception of the work from Hahlbrock’s laboratory (2, 3), previous studies of plant acetyl-CoA carboxylase have concentrated on the involvement of this enzyme in fatty acid biosynthesis (47). However, malonyl-CoA is also an intermediate in the biosynthesis of cuticular waxes (8), flavonoids (9), anthocyanins (9), stilbenoids (lo), malonyl-ACC5 (ll), an-
throquinones (12), and malonic acid (13). Compartmentation of the biosynthesis of some of these plant metabolites among the different cell types of leaves has been suggested (8, 14). In higher plants, fatty acid biosynthesis is localized in the plastids. This has been demonstrated for several plastid types, including chloroplasts (15) and proplastids (16). To establish the relationship between acetyl-CoA carboxylase and fatty acid biosynthesis in leaves, the distribution of acetyl-CoA carboxylase needs to be known among the different leaf compartments which metabolize malonyl-CoA. In a previous publication, we reported the distribution of acetyl-CoA carboxylase among the different cell types of leaves (1’7). In this publication, we present the subcellular distribution of this enzyme and identify
1 Supported in part by NSF Grants PCM79-039’76 and PCM81-04497. a Present address: Department of Cellular, Viral and Molecular Biology, University of Utah, Salt Lake City, Utah 84132. 3 Present address: NPI, 417 Wakara Way, Salt Lake City, Utah 84108. * To whom correspondence should be addressed.
6 Abbreviations used: ACC, l-aminocyclopropane1-carboxylic acid; ACP, acyl-carrier protein; BSA, bovine serum albumin; PAGE, polyacrylamide gel electrophoresis; PEP, phosphoenolpyruvate; PVP, polyvinylpyrrolydone; RuBP, ribulose 1,5-bisphosphate; SDS, sodium dodecyl sulfate; MES, I-morpholineethanesulfonic acid; Hepes, 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid. 555
0003-9861/84 Copyright All rights
$3.00
0 1984 by Academic Press. Inc. of reproduction in any form reserved.
556
NIKOLAU,
its biotinyl subunit in mesophyll a C3 and a C4 plant. MATERIALS
AND
WURTELE,
cells of
METHODS
Acetyl-CoA was synthesized by the method of Stadtman (18) as described previously (1’7). Streptavidin was a kind gift from Dr. E. 0. Stapley, Merck Institute. Iodination of streptavidin with Na’? was carried out by the method of Fracker and Speck (19). Biochemicals were purchased from Sigma, except Cellulysin and Macerase, which were obtained from Calbiochem. The radiochemicals, NaHi4C03 (54 Ci/mol) and Na’=I (carrier-free), were purchased from Amersham. Barley (H&m &are, var. CM67) and sorghum (Sorghum X Sudangrass hybrid, var. WAC Forage 99) seeds were soaked overnight in aerated water, and were germinated in conditions described previously (17). Barley and sorghum seedlings were harvested 7 and 5 days after planting, respectively. Mesophyll protoplasts were prepared from barley leaves essentially as described by Day et al. (20). Leaves were abraded with Carborundum and a painter’s brush, and washed with distilled water. Approximately 2 g leaves was cut transversely into 0.5- to l.O-mm strips. Leaf strips were vacuum infiltrated with 20 ml 2% (w/v) Cellulysin, 0.5% (w/v) Macerase, 0.6 M sorbitol, 0.2 mM CaCl,, 0.2 mM KHaPO,, 1 mM MgClp, 10 mM Mes-NaOH, pH 5.5, three times, and were digested for 2$ h at 28°C with shaking at 60 cycles/min. Following digestion, protoplasts were collected by filtering the digestion medium through an 80-&m-mesh nylon net. The leaf fragments were washed three times with about 5 ml 0.6 M sorbitol, 0.2 mM CaCl,, 0.2 mM KH2POI, 1 mM MgCl*, 5 mM Hepes-NaOH, pH 7.8 (isolation buffer) and the washings were pooled. All procedures from this point on were carried out at 4°C. Protoplasts were pelleted by centrifugation at 3009 for 3 min in a swinging-bucket rotor. Protoplasts were resuspended in a 0.6 M sucrose solution (identical to the isolation buffer, except sucrose replaced sorbitol), overlayed with isolation buffer, and centrifuged at 3008 for 5 min. The intact protoplasts were collected from the sucrose-sorbitol interface with a Pasteur pipet, washed with isolation buffer, and pelleted at 3008 for 3 min. Protoplasts were suspended in 0.3 M sorbitol, 10 m&i 2-mercaptoethanol, 1 mM EDTA, 10 mM Tris-HCl, pH 8.0, and were gently lysed by two passages through a go-pm-mesh nylon net. Sorghum mesophyll protoplasts were obtained following digestion of leaves in 1.5% (w/v) Cellulysin, 0.6 M mannitol, 30 mM Mes-NaOH, pH 5.6, at 28°C for 2 h, as described by Wurtele et al. (21). Isolated protoplasts were suspended in 0.6 M mannitol, 0.1% (W/V) BSA, 1% (w/v) PVP-40, 10 mM Z-mercaptoethanol, 1 mM EDTA, 10 mM Tris-HCl, pH 8.0. Lysis of
AND
STUMPF
the protoplasts was achieved by three passages through a 20-km-mesh nylon net. Aliquots of the ruptured protoplasts were layered onto linear sucrose density gradients [30% to 60% (w/w) sucrose] buffered with 10 mM Tris-HCl, pH 8.0, containing 1 mM EDTA and 10 mM 2-mercaptoethanol. Gradients were centrifuged for 20 min at a maximum speed of 23,000 rpm in a SW27 rotor (113,OOOg), and were fractionated into l-ml fractions. Gradient fractions and ruptured protoplasts were analyzed for acetyl-CoA carboxylase (6), NADPmalate dehydrogenase (22), catalase (23), and fumarase (24) activities. RuBP carboxylase activity was determined according to Wishnick and Lane (25), following 10 min activation at 30°C in the assay buffer without RuBP. PEP carboxylase was assayed as for RuBP carboxylase, but without prior activation and replacing RuBP with PEP. Chlorophyll was assayed by the method of Arnon (26), and protein by a dye-binding method (27). Polyacrylamide gel electrophoresis in the presence of SDS was carried out by the method of Laemmli (28) in gels composed of 10% (w/v) acrylamide/ 0.27% (w/v) bisacrylamide. Protoplast and sucrose density gradient fractions were adjusted, prior to electrophoresis, to 1% (w/v) SDS, 5% (v/v) 2-mercaptoethanol, and were heated for 5 min at 100°C. An extract of barley leaf was prepared for SDSPAGE by the procedure previously described (17). Biotinyl proteins were detected by a modified Western blotting procedure. Following SDS-PAGE, proteins were transferred from the gel to nitrocellulose paper electrophoretically, essentially as described by Towbin et al. (29), and the nitrocellulose paper was soaked overnight in a solution of 3% (w/ v) BSA in 10 mM Tris-HCl, pH 7.4, 0.9% (w/v) NaCl. To detect biotinyl proteins, the nitrocellulose paper was then soaked in the same solution containing 8.0 X lo6 cpm of [‘251]streptavidin (about 8 X 10’ cpm/ mg protein) for 2 h. Unbound [izI]streptavidin was removed by washing the paper three times with 10 mM Tris-HCl, pH 7.4, 0.9% (w/v) NaCl. After airdrying, the nitrocellulose paper was exposed to Kodak X-Omat G film at -7O”C, using a DuPont Cronex Lightning Plus intensifying screen. Exposed film was developed as recommended by the manufacturer with Kodak GBX developer. RESULTS
AND
DISCUSSION
Chloroplasts of leaf mesophyll cells are able to synthesize malonyl-CoA and utilize it for fatty acid biosynthesis (5-7, 30). To investigate whether other subcellular compartments of mesophyll cells are also able to synthesize malonyl-CoA, the subcellular distribution of acetyl-CoA carboxylase was determined in mesophyll
SUBCELLULAR
LOCALIZATION
OF
ACETYL-CoA
557
CARBOXYLASE
protoplasts isolated from barley and sorghum. The yields of protoplasts were between 1 X lo6 and 5 X lo6 protoplasts per gram of barley leaves, and between 2 X lo5 and 8 X lo5 protoplasts per gram of sorghum leaves. Figure 1 shows the distribution of enzymes and markers in a typical sucrose density gradient of ruptured barley mesophyll protoplasts. Two clearly defined regions in the gradient (fractions 5-8 and 12-18) contained chlorophyll. Eighty percent of the chlorophyll recovered from the gradient was found in the higher density band (peak density = 1.219 g/cm3) (Table I). This band also contained 81% of the recovered RuBP carboxylase activity, an enzyme characteristic of intact chloroplasts. The chlorophyll band at the lower density (peak density = 1.177 g/cm3) contained 20% of the chlorophyll recovered from the gradient and minimal RuBP carboxylase activity, consistent with the identification of this band as broken chloroplasts. Eight percent of the RuBP carboxylase activity was recovered in the first three fractions of the gradient. Approximately 80% of the chloroplasts retained their intactness during the course of this experiment, as judged from the distribution of chlorophyll and RuBP carboxylase activity in the gradient fractions. The peak density of the mitochondria, as indicated by fumarase activity, was located at 1.166 g/cm3 (fraction 6). Fu-
FIG. 1. Distribution of enzymes and markers in a linear sucrose density gradient following centrifugation of ruptured mesophyll protoplasts isolated from leaves of barley. Enzyme activities are expressed as nmol min? ml-‘, except for catalase and fumarase, which are in pmol min-’ ml-‘.
TABLE
I
DISTRIBUTIONANDRECOVERIESOFENZYMESANDMARKERSFOLLOWINGSUCROSEDENSITYGRADIENT CENTRIFUGATIONOFRUPTURED MESOPHYLL PROTOPLASTS FROMBARLEY Distribution
Enzyme
or marker
Protein Chl Catalase Fumarase PEP carboxylase RuBP carboxylase Acetyl-CoA earboxylase
Loading on gradient 3562
pg
448 m 1972 4.4 76.6 849 17.3
rmol/min amol/min nmol/min nmol/min nmol/min
Total recovery (%) 103 87 61 102 119 83 100
cytoso1 (fractions l-3) 24 0 64 66 89 8 11
within
gradient
Broken chloroplasts (fractions 5-8) 8 18 13 22 3 6 3
(%) Intact chloroplasts (fractions 12-18) 65 79 12 0 3 81 86
558
NIKOLAU,
WURTELE,
marase activity at the top of the gradient was due to the release of this enzyme from broken mitochondria. Catalase, used as a marker for peroxisomes, was located at the top of the gradient. Because of the short centrifugation time (20 min), peroxisomes did not attain their equilibrium density. PEP carboxylase activity was utilized as a cytoplasmic marker; 90% of the recovered activity was found at the top of the gradient. Contamination of the intact chloroplasts with mitochondrial, peroxisomal, and cytoplasmic enzymes was minimal, as judged by the distribution of fumarase, catalase, and PEP carboxylase activities, respectively (Table I). The distribution of acetyl-CoA carboxylase activity within the gradient was identical to that of RuBP carboxylase; namely, 86% of the recovered activity was associated with the intact chloroplasts and 11% was at the top of the gradient. The small amount of RuBP carboxylase and acetyl-CoA carboxylase activities associated with broken chloroplasts can be attributed to entrapment of the stromal content of chloroplasts by membrane vesicles. The recoveries of the enzyme activities and markers from the gradient ranged between 61 and 119% of that applied to the gradient. Thus, losses dur-
WL
M
cytoso I r 1235678910
AND
ing the course of the experiment were minimal. The distributions of acetyl-CoA carboxylase and RuBP carboxylase activities were found to be identical in six additional experiments with barley mesophyll protoplast gradients. In each case, the ratio of these two activities recovered in the intact chloroplast band was consistent with the proportion of intact chloroplasts, as judged by the chlorophyll distribution. The biotinyl subunit of acetyl-CoA carboxylase was detected by a modified Western blotting procedure. In contrast to many other plant species (17), barley mesophyll protoplasts and leaves contained a single biotinyl protein of molecular weight 60,000 (Fig. 2, lanes M and WL). The distribution of this 60-kDa biotinprotein band was determined in the fractions from the sucrose density gradient of ruptured mesophyll protoplasts of barley. The profile of this band in the gradient correlated closely to the profile of acetylCoA carboxylase activity. The majority of the biotin-protein, as judged by the intensity of the bands, was in fractions 12 to 15, corresponding to the region of the gradient containing intact chloroplasts (compare Figs. 1 and 2). Lesser amounts
Broken Chloroplasts I
STUMPF
Intact I
Chloroplasts
I Ii
I 12 13
14
15
I6
I7
I8
-Mol
Wt(XIo-3)
-
60
-
25
FIG. 2. Western blot indicating the presence of biotinyl proteins in barley leaves (WL), isolated barley mesophyll protoplasts (M), and biotinyl protein distribution in fractions from a linear sucrose density gradient following centrifugation of ruptured barley mesophyll protoplasts (l-17).
SUBCELLULAR
LOCALIZATION
5
10
OF
15
Froctm
FIG. 3. Distribution linear sucrose density gation of ruptured sorghum leaves.
of enzymes and protein in a gradient following centrifumesophyll protoplasts from
of this band were seen in the fractions from the top of the gradient; the presence of biotin-protein in these fractions is attributed to release of acetyl-CoA carboxylase from chloroplasts ruptured during the course of the experiment (Table I). There was also an increase in the intensity of the 60-kDa band in fraction ‘7, corresponding to acetyl-CoA carboxylase entrapped in membrane vesicles of broken chloroplasts. To extend our data to a C4 plant, the subcellular distribution of acetyl-CoA carboxylase activity was examined in meTABLE DISTRIBUTION
AND RECOVERIES CENTRIFUGATION
OF ENZYMES OF RUPTURED
ACETYL-CoA
sophyll protoplasts of sorghum. Figure 3 shows the distribution of enzymes and protein in a linear sucrose density gradient of lysed sorghum mesophyll protoplasts. Two distinct green bands were visible on visual inspection of the gradient prior to fractionation. However, the presence of neutral red dye at the top of the gradient precluded the determination of the chlorophyll distribution. The recovery of the markers used in this gradient was from 80% for NADPmalate dehydrogenase to over 100% for the recovery of protein (Table II). The position of intact chloroplasts, indicated by NADP-malate dehydrogenase activity, was at a peak density of 1.192 g/cm3 and included gradient fractions 8 to 15. Coincident with this activity peak was that of acetyl-CoA carboxylase. Seventy one percent of the recovered acetyl-CoA carboxylase activity and 70% of NADP-malate dehydrogenase activity was found in fractions 8 to 15 (Table II). The remainder of the activities were at the top of the gradient, and in a band of peak density of 1.166 g/cm3, which coincided with one of the green bands distinguishable by visual inspection. We attribute this band to broken chloroplasts in which these enzymes have become entrapped. Minimal contamination of intact and broken plastid fractions by cytoplasmic enzymes was indicated by the distribution of PEP carboxylase activity, 93% of which was found at the top of the gradient. Although previous studies have revealed the presence of acetyl-CoA carboxylase in chloroplasts (5-7, 31) and proplastids (16, II
AND PROTEIN FOLLOWING MESOPHYLL PROTOPLASTS Distribution
Enzyme
or marker
Protein PEP carboxylase NADP-malate dehydrogenase Acetyl-CoA carboxylase
Loading gradient 313 35.7 47.8 1.37
on
ia nmol/min nmol/min nmol/min
559
CARBOXYLASE
Total recovery (%)
Cytosol (fractions
120 82 85 99
43 93 18 20
l-3)
SUCROSE DENSITY FROM SORGHUM within
Broken chloroplasts (fractions 4-7) 17 2 12 9
gradient
GRADIENT
(96) Intact chloroplasts (fractions S-15) 40 6 70 71
560
NIKOLAU,
WURTELE,
31, 32, 43), the data presented in this publication demonstrate that acetyl-CoA carboxylase in mesophyll cells of barley and sorghum is primarily, if not solely, localized in the chloroplasts. Thus, in mesophyll cells of these species, all the reactions of fatty acid biosynthesis from acetate may occur exclusively within the chloroplasts (15, 33, 34). The absence of extrachloroplastic acetyl-CoA carboxylase in mesophyll cells means that biosynthetic reactions in these cells requiring malonyl-CoA as a substrate need to be located within the chloroplast or that malonyl-CoA can be translocated out of the chloroplasts. Many reactions requiring malonyl-CoA may be compartmented in cell types other than mesophyll cells. Indeed, acetyl-CoA carboxylase has been demonstrated in epidermal cells of C3 and C, plants and in the bundle sheath cells of Cq plants (6, 17). Furthermore, chalcone synthase, the enzyme requiring malonyl-CoA as a substrate for the biosynthesis of flavonoids, occurs exclusively in the epidermis of leaves (35). Another example of compartmentation of reactions requiring malonylCoA is the biosynthesis of very-long-chain fatty acids (chain lengths of &,-C,), the major building block of the cuticle of leaves (8) and the storage wax in jojoba seeds (36). The synthesis of very-longchain fatty acids requires acetyl-CoA carboxylase to supply malonyl-CoA for two sets of reactions: de novo synthesis of Cl8 fatty acids in the plastids, and for their elongation to C&& fatty acids in an extraplastidic compartment (8, 36). The synthesis of very-long-chain fatty acids in leaves takes place in the epidermal tissues (8). The subcellular distribution of acetyl-CoA carboxylase in nonmesophyll cells may be different from that found in this study. Most leaves contain multiple biotinyl proteins (17), which may be due to isoenzymes of acetyl-CoA carboxylase localized in different subcellular compartments to supply malonyl-CoA for a number of biosynthetic pathways. Multiple biotinyl proteins present in developing jojoba seeds (unpublished data) may also be representative of isoenzymes of acetyl-
AND
STUMPF
CoA carboxylase required to supply malonyl-CoA for de novo synthesis and elongation of fatty acids to very-long-chain fatty acids (37, 38). Using our modified Western blotting procedure, the 60-kDa biotinyl subunit of acetyl-CoA carboxylase was identified as the only biotin-containing peptide present in barley leaves and in mesophyll cells of sorghum (1’7). The size of the biotinyl protein found in this study agrees closely with the size of the biotinyl subunit of the purified maize leaf acetyl-CoA carboxylase (39). Biotinyl proteins of 60-kDa have also been found in leek and pea (17). Previously reported plant biotinyl proteins of 21 (40, 41), 240 (3), and 210 kDa (42) were not detected in this study. The cell type which was examined in this study, namely mesophyll cells from barley and sorghum leaves, contained a single biotinyl protein. However, most other cells and tissues examined contained multiple biotinyl proteins [(17), and unpublished data]. Analysis of the subcellular distribution of the biotinyl proteins and of acetyl-CoA carboxylase in these tissues may help to identify the function of the 51-, 34-, and 30-kDa biotin-containing proteins which have been detected in leaves of maize, sorghum, pea, and leek (17). ACKNOWLEDGMENTS We are grateful to Dr. E. 0. Stapley of Merck, Sharp and Dohme Research Lab, Rahway, New Jersey, for the kind gift of streptavidin. We particularly appreciate the encouragement and helpful suggestions of Dr. E. E. Conn, whose grant supported one of us (ESW). We thank Ms. Billie Gabriel for the preparation of the manuscript.
REFERENCES 1. LANE, M. D., Moss, J., AND POLAKIS, S. E. (1974) Cm-r. Top. Cell. Reg. 8, 139-195. 2. EBEL, J., AND HAHLBROCK, K. (1977) Eur. J. Biochem 75,201-209. 3. EGIN-B~HLER, B., LOYAL, R., AND EBEL, J. (1980) Eur. J. Biochem 203,90-100. 4. REITZEL, L., AND NIELSEN, N. C. (1976) Eur. J. B&&em 65,131-138. 5. THOMSON, L. W., AND ZALIK, S. (1981) Plant PhysioL 67, 655-661.
SUBCELLULAR
LOCALIZATION
6. NIKOLAU, B. J., HAWKE, J. C., AND SLACK, C. R. (1981) Arch Biochem Biophys. 211,605-612. 7. EASTWELL, K. C., AND STUMPF, P. K. (1983) Plant Physiol. 72, 50-55. 8. KOLATPUKUDY, P. E., CROTEAU, R., AND BUCKNE, J. S. (1976) in Chemistry and Biochemistry of Natural Waxes (Kolattukudy, P. E., ed.), pp. 289-347, Elsevier/North-Holland, New York/ Amsterdam. 9. HAHLBROCK, K. (1981) in The Biochemistry of Plant; A Comprehensive Treatise (Stumpf, P. K., and Conn, E. E., eds.), Vol. 7, pp. 425456, Academic Press, New York. 10. GORHAM, J. (1980) in Progress in Phytochemistry (Reinhold, L., Harborne, J. B., and Swain, T., eds.), Vol. 6, pp. 203-252, Pergamon Press, Oxford. 11. AMRHEIM, N., AND KIONKA, C. (1983) Plant Physiol 72, s-37. 12. PACKTER, N. M. (1980) in The Biochemistry of Plants; A Comprehensive Treatise (Stumpf, P. K. and Conn, E. E., eds.), Vol. 4, pp. 535570, Academic Press, New York. 13. STUMPF, D. K., AND BURRIS, R. H. (1981) Plant Physiol. 68,992-995. 14. WIERMANN, R. (1981) in The Biochemistry of Plants; A Comprehensive Treatise (Stumpf, P. K., and Conn, E. E., eds.), Vol. 7, pp. 85116, Academic Press, New York. 15. OHLROGGE, J. B., KUHN, D. N., AND STUMPF, P. K. (1979) Proc. Natl Acad Sci USA 76, 1194-1198. 16. VICK, B., AND BEEVERS, H. (1978) Plant Physiol. 62,173-178. 17. NIKOLAU, B. J., WURTELE, E. S., AND STUMPF, P. K. (1984) Plant Physiol 75, 895-901. 18. STADTMAN, E. R. (1957) in Methods in Enzymology (Colowick, S. P., and Kaplan, N. O., eds.), Vol. 3, pp. 931-941, Academic Press, New York. 19. FRACKER, P. J., AND SPECK, J. C. (1978) B&hem Biophys. Res. Commun 80, 849-857. 20. DAY, D. A., JENKINS, C. L. D., AND HATCH, M. D. (1981) Au&. J. Plant Physiol 8,21-29. 21. WURTELE, E. S., THAYER, S. S., AND CONN, E. E. (1982) Plant Physiol 70.1732-1737. 22. HATCH, M. D., AND SLACK, C. R. (1969) B&hem. Biophys. Res. Commun 34,589-593.
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ACETYL-CoA
CARBOXYLASE
561
23. LUCK, H. (1965) in Methods of Enzymatic Analysis (Bergmeyer, H-U., ed.), pp. 885-894, Academic Press, New York. 24. HILL, R. L. AND BRADSHAW, R. A. (1969) in Methods in Enzymology (Lowenstein, J. M., ed.), Vol. 13, pp. 91-99, Academic Press, New York. 25. WISHNICK, M., AND LANE, M. D. (1971) in Methods in Enzymology (San Pietro, A., ed.), Vol. 23, pp. 570-599, Academic Press, New York. 26. ARNON, D. I. (1949) Plant Physiol. 24,1-15. 27. BRADFORD, M. M. (1976) Anal. Biochem 72,248254. 28. LAEMMLI, II. K. (1970) Nature (London) 227,680685. 29. TOWBIN, H., STAEHELIN, T., AND GORDON, J. (1979) Proc. Nat1 Acad Sci USA 76,4350-4354. 30. STUMPF, P. K. (1980) in The Biochemistry of Plants; A Comprehensive Treatise (Stumpf, P. K., and Conn, E. E., eds.), Vol. 4, pp. 177204, Academic Press, New York. 31. MOHAN, S. B., AND KEKWICK, R. G. 0. (1980) B&hem. J. 187, 667-676. 32. BURDEN, T. S., AND CANVIN, D. T. (1975) Canad J. Bot. 53, 1371-1376. 33. KUHN, D. N., KNAUF, M., AND STUMPF, P. K. (1981) Arch Biochem Biophys. 209, 441-450. 34. SHIMAKATA, T., AND STUMPF, P. K. (1982) Plant Physiol. 69.1257-1262. 35. HRAZDINA, G., MARX, G. A., AND HOCH, H. C. (1982) Plant Physiol. 70,745-748. 36. YERMANOS, D. M. (1975) J. Amer. CM Chem. Sot. 52,115-117. 37. OHLROGGE, J. B., POLLARD, M. R., AND STUMPF, P. K. (1978) Lipids 13,203-210. 38. POLLARD, M. R., MCKEON, T., GUPTA, L. M., AND STIJMPF, P. K. (1979) Lipids 14, 651-662. 39. NIKOLAU, B. J., AND HAWKE, J. C. (1984) Arch Biochem. Biophys. 228,86-96. 40. BROCK, K., AND KANNANGARA, C. G. (1976) Carlsberg Res. Commun 41,121-129. 41. KANNANGARA, C. G., AND JENSEN, C. J. (1975) Eur. J. B&hem 54,25-30. 42. EGIN-B~HLER, B., AND EBEL, J. (1983) Eur. J. B&hem. 133,335-339. 43. FINLAYSON, S. A., AND DENNIS, D. T. (1983) Arch. B&hem+ Biophys. 225, 576-585.
OF BIOCHEMISTRY
Vol. 235, No. 2, December,
AND
BIOPHYSICS
pp. 555-561,
1984
Subcellular Distribution of Acetyl-Coenzyme A Carboxylase Mesophyll Cells of Barley and Sorghum Leaves’ BASIL Department
J. NIKOLAII, of Biochemistry Received
EVE SYRKIN and Biophysics, April
WURTELE: University
27, 1984, and in revised
AND
PAUL
of CalZfornia,
Davis,
form
8, 1984
August
in
K. STUMPF4 California
95616
The subcellular distribution of acetyl-CoA carboxylase [acetyl-CoA-carbon dioxide ligase (ADP-forming), EC 6.4.1.21 was determined in mesophyll protoplasts isolation from barley, a C3 plant, and sorghum, a C4 plant. In both species, all of the mesophyll acetyl-CoA carboxylase was demonstrated to be chloroplastic. In barley leaves and mesophyll protoplasts, a single biotinyl protein of 60,000 Da was identified by a modified Western-blotting procedure. The subcellular distribution of this biotinyl protein was identical to that found for acetyl-CoA carboxylase. These results are discussed in relation to the compartmentation of reactions requiring malonyl-CoA as a substrate. (c 1984 Academic Pruss. Inc.
Acetyl-CoA carboxylase catalyzes the rate-limiting reaction in the biosynthesis of fatty acids in Escherichia coli, yeasts, and various mammalian tissues (1). In these organisms, malonyl-CoA, the product of the reaction catalyzed by acetylCoA carboxylase, is utilized only in the biosynthesis of fatty acids, and a correlation between acetyl-CoA carboxylase activity and fatty acid biosynthesis has been demonstrated. With the exception of the work from Hahlbrock’s laboratory (2, 3), previous studies of plant acetyl-CoA carboxylase have concentrated on the involvement of this enzyme in fatty acid biosynthesis (47). However, malonyl-CoA is also an intermediate in the biosynthesis of cuticular waxes (8), flavonoids (9), anthocyanins (9), stilbenoids (lo), malonyl-ACC5 (ll), an-
throquinones (12), and malonic acid (13). Compartmentation of the biosynthesis of some of these plant metabolites among the different cell types of leaves has been suggested (8, 14). In higher plants, fatty acid biosynthesis is localized in the plastids. This has been demonstrated for several plastid types, including chloroplasts (15) and proplastids (16). To establish the relationship between acetyl-CoA carboxylase and fatty acid biosynthesis in leaves, the distribution of acetyl-CoA carboxylase needs to be known among the different leaf compartments which metabolize malonyl-CoA. In a previous publication, we reported the distribution of acetyl-CoA carboxylase among the different cell types of leaves (1’7). In this publication, we present the subcellular distribution of this enzyme and identify
1 Supported in part by NSF Grants PCM79-039’76 and PCM81-04497. a Present address: Department of Cellular, Viral and Molecular Biology, University of Utah, Salt Lake City, Utah 84132. 3 Present address: NPI, 417 Wakara Way, Salt Lake City, Utah 84108. * To whom correspondence should be addressed.
6 Abbreviations used: ACC, l-aminocyclopropane1-carboxylic acid; ACP, acyl-carrier protein; BSA, bovine serum albumin; PAGE, polyacrylamide gel electrophoresis; PEP, phosphoenolpyruvate; PVP, polyvinylpyrrolydone; RuBP, ribulose 1,5-bisphosphate; SDS, sodium dodecyl sulfate; MES, I-morpholineethanesulfonic acid; Hepes, 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid. 555
0003-9861/84 Copyright All rights
$3.00
0 1984 by Academic Press. Inc. of reproduction in any form reserved.
556
NIKOLAU,
its biotinyl subunit in mesophyll a C3 and a C4 plant. MATERIALS
AND
WURTELE,
cells of
METHODS
Acetyl-CoA was synthesized by the method of Stadtman (18) as described previously (1’7). Streptavidin was a kind gift from Dr. E. 0. Stapley, Merck Institute. Iodination of streptavidin with Na’? was carried out by the method of Fracker and Speck (19). Biochemicals were purchased from Sigma, except Cellulysin and Macerase, which were obtained from Calbiochem. The radiochemicals, NaHi4C03 (54 Ci/mol) and Na’=I (carrier-free), were purchased from Amersham. Barley (H&m &are, var. CM67) and sorghum (Sorghum X Sudangrass hybrid, var. WAC Forage 99) seeds were soaked overnight in aerated water, and were germinated in conditions described previously (17). Barley and sorghum seedlings were harvested 7 and 5 days after planting, respectively. Mesophyll protoplasts were prepared from barley leaves essentially as described by Day et al. (20). Leaves were abraded with Carborundum and a painter’s brush, and washed with distilled water. Approximately 2 g leaves was cut transversely into 0.5- to l.O-mm strips. Leaf strips were vacuum infiltrated with 20 ml 2% (w/v) Cellulysin, 0.5% (w/v) Macerase, 0.6 M sorbitol, 0.2 mM CaCl,, 0.2 mM KHaPO,, 1 mM MgClp, 10 mM Mes-NaOH, pH 5.5, three times, and were digested for 2$ h at 28°C with shaking at 60 cycles/min. Following digestion, protoplasts were collected by filtering the digestion medium through an 80-&m-mesh nylon net. The leaf fragments were washed three times with about 5 ml 0.6 M sorbitol, 0.2 mM CaCl,, 0.2 mM KH2POI, 1 mM MgCl*, 5 mM Hepes-NaOH, pH 7.8 (isolation buffer) and the washings were pooled. All procedures from this point on were carried out at 4°C. Protoplasts were pelleted by centrifugation at 3009 for 3 min in a swinging-bucket rotor. Protoplasts were resuspended in a 0.6 M sucrose solution (identical to the isolation buffer, except sucrose replaced sorbitol), overlayed with isolation buffer, and centrifuged at 3008 for 5 min. The intact protoplasts were collected from the sucrose-sorbitol interface with a Pasteur pipet, washed with isolation buffer, and pelleted at 3008 for 3 min. Protoplasts were suspended in 0.3 M sorbitol, 10 m&i 2-mercaptoethanol, 1 mM EDTA, 10 mM Tris-HCl, pH 8.0, and were gently lysed by two passages through a go-pm-mesh nylon net. Sorghum mesophyll protoplasts were obtained following digestion of leaves in 1.5% (w/v) Cellulysin, 0.6 M mannitol, 30 mM Mes-NaOH, pH 5.6, at 28°C for 2 h, as described by Wurtele et al. (21). Isolated protoplasts were suspended in 0.6 M mannitol, 0.1% (W/V) BSA, 1% (w/v) PVP-40, 10 mM Z-mercaptoethanol, 1 mM EDTA, 10 mM Tris-HCl, pH 8.0. Lysis of
AND
STUMPF
the protoplasts was achieved by three passages through a 20-km-mesh nylon net. Aliquots of the ruptured protoplasts were layered onto linear sucrose density gradients [30% to 60% (w/w) sucrose] buffered with 10 mM Tris-HCl, pH 8.0, containing 1 mM EDTA and 10 mM 2-mercaptoethanol. Gradients were centrifuged for 20 min at a maximum speed of 23,000 rpm in a SW27 rotor (113,OOOg), and were fractionated into l-ml fractions. Gradient fractions and ruptured protoplasts were analyzed for acetyl-CoA carboxylase (6), NADPmalate dehydrogenase (22), catalase (23), and fumarase (24) activities. RuBP carboxylase activity was determined according to Wishnick and Lane (25), following 10 min activation at 30°C in the assay buffer without RuBP. PEP carboxylase was assayed as for RuBP carboxylase, but without prior activation and replacing RuBP with PEP. Chlorophyll was assayed by the method of Arnon (26), and protein by a dye-binding method (27). Polyacrylamide gel electrophoresis in the presence of SDS was carried out by the method of Laemmli (28) in gels composed of 10% (w/v) acrylamide/ 0.27% (w/v) bisacrylamide. Protoplast and sucrose density gradient fractions were adjusted, prior to electrophoresis, to 1% (w/v) SDS, 5% (v/v) 2-mercaptoethanol, and were heated for 5 min at 100°C. An extract of barley leaf was prepared for SDSPAGE by the procedure previously described (17). Biotinyl proteins were detected by a modified Western blotting procedure. Following SDS-PAGE, proteins were transferred from the gel to nitrocellulose paper electrophoretically, essentially as described by Towbin et al. (29), and the nitrocellulose paper was soaked overnight in a solution of 3% (w/ v) BSA in 10 mM Tris-HCl, pH 7.4, 0.9% (w/v) NaCl. To detect biotinyl proteins, the nitrocellulose paper was then soaked in the same solution containing 8.0 X lo6 cpm of [‘251]streptavidin (about 8 X 10’ cpm/ mg protein) for 2 h. Unbound [izI]streptavidin was removed by washing the paper three times with 10 mM Tris-HCl, pH 7.4, 0.9% (w/v) NaCl. After airdrying, the nitrocellulose paper was exposed to Kodak X-Omat G film at -7O”C, using a DuPont Cronex Lightning Plus intensifying screen. Exposed film was developed as recommended by the manufacturer with Kodak GBX developer. RESULTS
AND
DISCUSSION
Chloroplasts of leaf mesophyll cells are able to synthesize malonyl-CoA and utilize it for fatty acid biosynthesis (5-7, 30). To investigate whether other subcellular compartments of mesophyll cells are also able to synthesize malonyl-CoA, the subcellular distribution of acetyl-CoA carboxylase was determined in mesophyll
SUBCELLULAR
LOCALIZATION
OF
ACETYL-CoA
557
CARBOXYLASE
protoplasts isolated from barley and sorghum. The yields of protoplasts were between 1 X lo6 and 5 X lo6 protoplasts per gram of barley leaves, and between 2 X lo5 and 8 X lo5 protoplasts per gram of sorghum leaves. Figure 1 shows the distribution of enzymes and markers in a typical sucrose density gradient of ruptured barley mesophyll protoplasts. Two clearly defined regions in the gradient (fractions 5-8 and 12-18) contained chlorophyll. Eighty percent of the chlorophyll recovered from the gradient was found in the higher density band (peak density = 1.219 g/cm3) (Table I). This band also contained 81% of the recovered RuBP carboxylase activity, an enzyme characteristic of intact chloroplasts. The chlorophyll band at the lower density (peak density = 1.177 g/cm3) contained 20% of the chlorophyll recovered from the gradient and minimal RuBP carboxylase activity, consistent with the identification of this band as broken chloroplasts. Eight percent of the RuBP carboxylase activity was recovered in the first three fractions of the gradient. Approximately 80% of the chloroplasts retained their intactness during the course of this experiment, as judged from the distribution of chlorophyll and RuBP carboxylase activity in the gradient fractions. The peak density of the mitochondria, as indicated by fumarase activity, was located at 1.166 g/cm3 (fraction 6). Fu-
FIG. 1. Distribution of enzymes and markers in a linear sucrose density gradient following centrifugation of ruptured mesophyll protoplasts isolated from leaves of barley. Enzyme activities are expressed as nmol min? ml-‘, except for catalase and fumarase, which are in pmol min-’ ml-‘.
TABLE
I
DISTRIBUTIONANDRECOVERIESOFENZYMESANDMARKERSFOLLOWINGSUCROSEDENSITYGRADIENT CENTRIFUGATIONOFRUPTURED MESOPHYLL PROTOPLASTS FROMBARLEY Distribution
Enzyme
or marker
Protein Chl Catalase Fumarase PEP carboxylase RuBP carboxylase Acetyl-CoA earboxylase
Loading on gradient 3562
pg
448 m 1972 4.4 76.6 849 17.3
rmol/min amol/min nmol/min nmol/min nmol/min
Total recovery (%) 103 87 61 102 119 83 100
cytoso1 (fractions l-3) 24 0 64 66 89 8 11
within
gradient
Broken chloroplasts (fractions 5-8) 8 18 13 22 3 6 3
(%) Intact chloroplasts (fractions 12-18) 65 79 12 0 3 81 86
558
NIKOLAU,
WURTELE,
marase activity at the top of the gradient was due to the release of this enzyme from broken mitochondria. Catalase, used as a marker for peroxisomes, was located at the top of the gradient. Because of the short centrifugation time (20 min), peroxisomes did not attain their equilibrium density. PEP carboxylase activity was utilized as a cytoplasmic marker; 90% of the recovered activity was found at the top of the gradient. Contamination of the intact chloroplasts with mitochondrial, peroxisomal, and cytoplasmic enzymes was minimal, as judged by the distribution of fumarase, catalase, and PEP carboxylase activities, respectively (Table I). The distribution of acetyl-CoA carboxylase activity within the gradient was identical to that of RuBP carboxylase; namely, 86% of the recovered activity was associated with the intact chloroplasts and 11% was at the top of the gradient. The small amount of RuBP carboxylase and acetyl-CoA carboxylase activities associated with broken chloroplasts can be attributed to entrapment of the stromal content of chloroplasts by membrane vesicles. The recoveries of the enzyme activities and markers from the gradient ranged between 61 and 119% of that applied to the gradient. Thus, losses dur-
WL
M
cytoso I r 1235678910
AND
ing the course of the experiment were minimal. The distributions of acetyl-CoA carboxylase and RuBP carboxylase activities were found to be identical in six additional experiments with barley mesophyll protoplast gradients. In each case, the ratio of these two activities recovered in the intact chloroplast band was consistent with the proportion of intact chloroplasts, as judged by the chlorophyll distribution. The biotinyl subunit of acetyl-CoA carboxylase was detected by a modified Western blotting procedure. In contrast to many other plant species (17), barley mesophyll protoplasts and leaves contained a single biotinyl protein of molecular weight 60,000 (Fig. 2, lanes M and WL). The distribution of this 60-kDa biotinprotein band was determined in the fractions from the sucrose density gradient of ruptured mesophyll protoplasts of barley. The profile of this band in the gradient correlated closely to the profile of acetylCoA carboxylase activity. The majority of the biotin-protein, as judged by the intensity of the bands, was in fractions 12 to 15, corresponding to the region of the gradient containing intact chloroplasts (compare Figs. 1 and 2). Lesser amounts
Broken Chloroplasts I
STUMPF
Intact I
Chloroplasts
I Ii
I 12 13
14
15
I6
I7
I8
-Mol
Wt(XIo-3)
-
60
-
25
FIG. 2. Western blot indicating the presence of biotinyl proteins in barley leaves (WL), isolated barley mesophyll protoplasts (M), and biotinyl protein distribution in fractions from a linear sucrose density gradient following centrifugation of ruptured barley mesophyll protoplasts (l-17).
SUBCELLULAR
LOCALIZATION
5
10
OF
15
Froctm
FIG. 3. Distribution linear sucrose density gation of ruptured sorghum leaves.
of enzymes and protein in a gradient following centrifumesophyll protoplasts from
of this band were seen in the fractions from the top of the gradient; the presence of biotin-protein in these fractions is attributed to release of acetyl-CoA carboxylase from chloroplasts ruptured during the course of the experiment (Table I). There was also an increase in the intensity of the 60-kDa band in fraction ‘7, corresponding to acetyl-CoA carboxylase entrapped in membrane vesicles of broken chloroplasts. To extend our data to a C4 plant, the subcellular distribution of acetyl-CoA carboxylase activity was examined in meTABLE DISTRIBUTION
AND RECOVERIES CENTRIFUGATION
OF ENZYMES OF RUPTURED
ACETYL-CoA
sophyll protoplasts of sorghum. Figure 3 shows the distribution of enzymes and protein in a linear sucrose density gradient of lysed sorghum mesophyll protoplasts. Two distinct green bands were visible on visual inspection of the gradient prior to fractionation. However, the presence of neutral red dye at the top of the gradient precluded the determination of the chlorophyll distribution. The recovery of the markers used in this gradient was from 80% for NADPmalate dehydrogenase to over 100% for the recovery of protein (Table II). The position of intact chloroplasts, indicated by NADP-malate dehydrogenase activity, was at a peak density of 1.192 g/cm3 and included gradient fractions 8 to 15. Coincident with this activity peak was that of acetyl-CoA carboxylase. Seventy one percent of the recovered acetyl-CoA carboxylase activity and 70% of NADP-malate dehydrogenase activity was found in fractions 8 to 15 (Table II). The remainder of the activities were at the top of the gradient, and in a band of peak density of 1.166 g/cm3, which coincided with one of the green bands distinguishable by visual inspection. We attribute this band to broken chloroplasts in which these enzymes have become entrapped. Minimal contamination of intact and broken plastid fractions by cytoplasmic enzymes was indicated by the distribution of PEP carboxylase activity, 93% of which was found at the top of the gradient. Although previous studies have revealed the presence of acetyl-CoA carboxylase in chloroplasts (5-7, 31) and proplastids (16, II
AND PROTEIN FOLLOWING MESOPHYLL PROTOPLASTS Distribution
Enzyme
or marker
Protein PEP carboxylase NADP-malate dehydrogenase Acetyl-CoA carboxylase
Loading gradient 313 35.7 47.8 1.37
on
ia nmol/min nmol/min nmol/min
559
CARBOXYLASE
Total recovery (%)
Cytosol (fractions
120 82 85 99
43 93 18 20
l-3)
SUCROSE DENSITY FROM SORGHUM within
Broken chloroplasts (fractions 4-7) 17 2 12 9
gradient
GRADIENT
(96) Intact chloroplasts (fractions S-15) 40 6 70 71
560
NIKOLAU,
WURTELE,
31, 32, 43), the data presented in this publication demonstrate that acetyl-CoA carboxylase in mesophyll cells of barley and sorghum is primarily, if not solely, localized in the chloroplasts. Thus, in mesophyll cells of these species, all the reactions of fatty acid biosynthesis from acetate may occur exclusively within the chloroplasts (15, 33, 34). The absence of extrachloroplastic acetyl-CoA carboxylase in mesophyll cells means that biosynthetic reactions in these cells requiring malonyl-CoA as a substrate need to be located within the chloroplast or that malonyl-CoA can be translocated out of the chloroplasts. Many reactions requiring malonyl-CoA may be compartmented in cell types other than mesophyll cells. Indeed, acetyl-CoA carboxylase has been demonstrated in epidermal cells of C3 and C, plants and in the bundle sheath cells of Cq plants (6, 17). Furthermore, chalcone synthase, the enzyme requiring malonyl-CoA as a substrate for the biosynthesis of flavonoids, occurs exclusively in the epidermis of leaves (35). Another example of compartmentation of reactions requiring malonylCoA is the biosynthesis of very-long-chain fatty acids (chain lengths of &,-C,), the major building block of the cuticle of leaves (8) and the storage wax in jojoba seeds (36). The synthesis of very-longchain fatty acids requires acetyl-CoA carboxylase to supply malonyl-CoA for two sets of reactions: de novo synthesis of Cl8 fatty acids in the plastids, and for their elongation to C&& fatty acids in an extraplastidic compartment (8, 36). The synthesis of very-long-chain fatty acids in leaves takes place in the epidermal tissues (8). The subcellular distribution of acetyl-CoA carboxylase in nonmesophyll cells may be different from that found in this study. Most leaves contain multiple biotinyl proteins (17), which may be due to isoenzymes of acetyl-CoA carboxylase localized in different subcellular compartments to supply malonyl-CoA for a number of biosynthetic pathways. Multiple biotinyl proteins present in developing jojoba seeds (unpublished data) may also be representative of isoenzymes of acetyl-
AND
STUMPF
CoA carboxylase required to supply malonyl-CoA for de novo synthesis and elongation of fatty acids to very-long-chain fatty acids (37, 38). Using our modified Western blotting procedure, the 60-kDa biotinyl subunit of acetyl-CoA carboxylase was identified as the only biotin-containing peptide present in barley leaves and in mesophyll cells of sorghum (1’7). The size of the biotinyl protein found in this study agrees closely with the size of the biotinyl subunit of the purified maize leaf acetyl-CoA carboxylase (39). Biotinyl proteins of 60-kDa have also been found in leek and pea (17). Previously reported plant biotinyl proteins of 21 (40, 41), 240 (3), and 210 kDa (42) were not detected in this study. The cell type which was examined in this study, namely mesophyll cells from barley and sorghum leaves, contained a single biotinyl protein. However, most other cells and tissues examined contained multiple biotinyl proteins [(17), and unpublished data]. Analysis of the subcellular distribution of the biotinyl proteins and of acetyl-CoA carboxylase in these tissues may help to identify the function of the 51-, 34-, and 30-kDa biotin-containing proteins which have been detected in leaves of maize, sorghum, pea, and leek (17). ACKNOWLEDGMENTS We are grateful to Dr. E. 0. Stapley of Merck, Sharp and Dohme Research Lab, Rahway, New Jersey, for the kind gift of streptavidin. We particularly appreciate the encouragement and helpful suggestions of Dr. E. E. Conn, whose grant supported one of us (ESW). We thank Ms. Billie Gabriel for the preparation of the manuscript.
REFERENCES 1. LANE, M. D., Moss, J., AND POLAKIS, S. E. (1974) Cm-r. Top. Cell. Reg. 8, 139-195. 2. EBEL, J., AND HAHLBROCK, K. (1977) Eur. J. Biochem 75,201-209. 3. EGIN-B~HLER, B., LOYAL, R., AND EBEL, J. (1980) Eur. J. Biochem 203,90-100. 4. REITZEL, L., AND NIELSEN, N. C. (1976) Eur. J. B&&em 65,131-138. 5. THOMSON, L. W., AND ZALIK, S. (1981) Plant PhysioL 67, 655-661.
SUBCELLULAR
LOCALIZATION
6. NIKOLAU, B. J., HAWKE, J. C., AND SLACK, C. R. (1981) Arch Biochem Biophys. 211,605-612. 7. EASTWELL, K. C., AND STUMPF, P. K. (1983) Plant Physiol. 72, 50-55. 8. KOLATPUKUDY, P. E., CROTEAU, R., AND BUCKNE, J. S. (1976) in Chemistry and Biochemistry of Natural Waxes (Kolattukudy, P. E., ed.), pp. 289-347, Elsevier/North-Holland, New York/ Amsterdam. 9. HAHLBROCK, K. (1981) in The Biochemistry of Plant; A Comprehensive Treatise (Stumpf, P. K., and Conn, E. E., eds.), Vol. 7, pp. 425456, Academic Press, New York. 10. GORHAM, J. (1980) in Progress in Phytochemistry (Reinhold, L., Harborne, J. B., and Swain, T., eds.), Vol. 6, pp. 203-252, Pergamon Press, Oxford. 11. AMRHEIM, N., AND KIONKA, C. (1983) Plant Physiol 72, s-37. 12. PACKTER, N. M. (1980) in The Biochemistry of Plants; A Comprehensive Treatise (Stumpf, P. K. and Conn, E. E., eds.), Vol. 4, pp. 535570, Academic Press, New York. 13. STUMPF, D. K., AND BURRIS, R. H. (1981) Plant Physiol. 68,992-995. 14. WIERMANN, R. (1981) in The Biochemistry of Plants; A Comprehensive Treatise (Stumpf, P. K., and Conn, E. E., eds.), Vol. 7, pp. 85116, Academic Press, New York. 15. OHLROGGE, J. B., KUHN, D. N., AND STUMPF, P. K. (1979) Proc. Natl Acad Sci USA 76, 1194-1198. 16. VICK, B., AND BEEVERS, H. (1978) Plant Physiol. 62,173-178. 17. NIKOLAU, B. J., WURTELE, E. S., AND STUMPF, P. K. (1984) Plant Physiol 75, 895-901. 18. STADTMAN, E. R. (1957) in Methods in Enzymology (Colowick, S. P., and Kaplan, N. O., eds.), Vol. 3, pp. 931-941, Academic Press, New York. 19. FRACKER, P. J., AND SPECK, J. C. (1978) B&hem Biophys. Res. Commun 80, 849-857. 20. DAY, D. A., JENKINS, C. L. D., AND HATCH, M. D. (1981) Au&. J. Plant Physiol 8,21-29. 21. WURTELE, E. S., THAYER, S. S., AND CONN, E. E. (1982) Plant Physiol 70.1732-1737. 22. HATCH, M. D., AND SLACK, C. R. (1969) B&hem. Biophys. Res. Commun 34,589-593.
OF
ACETYL-CoA
CARBOXYLASE
561
23. LUCK, H. (1965) in Methods of Enzymatic Analysis (Bergmeyer, H-U., ed.), pp. 885-894, Academic Press, New York. 24. HILL, R. L. AND BRADSHAW, R. A. (1969) in Methods in Enzymology (Lowenstein, J. M., ed.), Vol. 13, pp. 91-99, Academic Press, New York. 25. WISHNICK, M., AND LANE, M. D. (1971) in Methods in Enzymology (San Pietro, A., ed.), Vol. 23, pp. 570-599, Academic Press, New York. 26. ARNON, D. I. (1949) Plant Physiol. 24,1-15. 27. BRADFORD, M. M. (1976) Anal. Biochem 72,248254. 28. LAEMMLI, II. K. (1970) Nature (London) 227,680685. 29. TOWBIN, H., STAEHELIN, T., AND GORDON, J. (1979) Proc. Nat1 Acad Sci USA 76,4350-4354. 30. STUMPF, P. K. (1980) in The Biochemistry of Plants; A Comprehensive Treatise (Stumpf, P. K., and Conn, E. E., eds.), Vol. 4, pp. 177204, Academic Press, New York. 31. MOHAN, S. B., AND KEKWICK, R. G. 0. (1980) B&hem. J. 187, 667-676. 32. BURDEN, T. S., AND CANVIN, D. T. (1975) Canad J. Bot. 53, 1371-1376. 33. KUHN, D. N., KNAUF, M., AND STUMPF, P. K. (1981) Arch Biochem Biophys. 209, 441-450. 34. SHIMAKATA, T., AND STUMPF, P. K. (1982) Plant Physiol. 69.1257-1262. 35. HRAZDINA, G., MARX, G. A., AND HOCH, H. C. (1982) Plant Physiol. 70,745-748. 36. YERMANOS, D. M. (1975) J. Amer. CM Chem. Sot. 52,115-117. 37. OHLROGGE, J. B., POLLARD, M. R., AND STUMPF, P. K. (1978) Lipids 13,203-210. 38. POLLARD, M. R., MCKEON, T., GUPTA, L. M., AND STIJMPF, P. K. (1979) Lipids 14, 651-662. 39. NIKOLAU, B. J., AND HAWKE, J. C. (1984) Arch Biochem. Biophys. 228,86-96. 40. BROCK, K., AND KANNANGARA, C. G. (1976) Carlsberg Res. Commun 41,121-129. 41. KANNANGARA, C. G., AND JENSEN, C. J. (1975) Eur. J. B&hem 54,25-30. 42. EGIN-B~HLER, B., AND EBEL, J. (1983) Eur. J. B&hem. 133,335-339. 43. FINLAYSON, S. A., AND DENNIS, D. T. (1983) Arch. B&hem+ Biophys. 225, 576-585.