Ecology and Metabolism of Plant Lipids - American Chemical Society

Chapter 4

Fatty Acids in Plants: A Model System 1

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Downloaded by UNIV OF CALIFORNIA SANTA BARBARA on August 26, 2015 | http://pubs.acs.org Publication Date: December 24, 1987 | doi: 10.1021/bk-1987-0325.ch004

Glenn Fuller and P. K. Stumpf 1

Western Regional Research Center, Agricultural Research Service, U.S. Department of Agriculture, Albany, CA 94710 Department of Biochemistry and Biophysics, University of California, Davis, CA 95616

2

Fatty acids in plants are synthesized from acetate in the chloroplasts of leaf cells and the plastids of seeds. In leaf cells there are active desaturase enzymes which convert the fatty acids to linoleic and linolenic acids in large amounts. The predominant acyl lipids are triglycerides in seeds, phospholipids and glycolipids in leaves. Lemna minor was used to examine lipid synthesis in a plant which can grow under both light and dark conditions when provided proper nutrition. In interactions of plants with t h e i r environment, changes i n the membrane l i p i d s of the host are often i n evidence, e s p e c i a l l y chemical changes i n the composition of f a t t y acids. The biosynthetic pathways to these f a t t y acids are modified and i t i s these changes which are the subject of much current study. De novo synthesis of saturated f a t t y acids i n both procaryotes and eucaryotes i s w e l l understood. Subsequent desaturation reactions and pathways by which acyl glycerides are formed are not as w e l l defined. I n t h i s chapter we w i l l discuss f a t t y acid and t r i g l y c e r i d e biosynthesis and w i l l present data on using duckweed, a small aquatic weed, to study some of the reactions of l i p i d s i n plants. Fatty Acid Synthesis Synthesis of f a t t y acids i n plants has been the p r i n c i p a l subject of a number of recent reviews (1, 2, 3, 4 ) . The properties of a c y l lipids are determined by t h e i r f a t t y acid composition and d i s t r i b u t i o n . For instance, Van der Waals bonding i s stronger f o r saturated f a t t y acids than f o r cis-unsaturated acids having the same carbon number (5); thus, the saturated acids and t h e i r a c y l l i p i d s have higher melting points. There i s some evidence that lower melting points or phase t r a n s i t i o n temperatures i n membrane l i p i d s c o r r e l a t e with c h i l l resistance (6). The melting range, oxidation s t a b i l i t y , and Theological properties of t r i g l y c e r i d e s

This chapter not subject to U.S. copyright. Published 1987, American Chemical Society

In Ecology and Metabolism of Plant Lipids; Fuller, G., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 1987.


4. FULLER AND STUMPF

45

Fatty Acids in Plants

are a l l influenced by t h e i r composition and are c r i t i c a l t o t h e i r commercial uses. Each plant c e l l manufactures i t s own f a t t y acids, since there i s no l i p i d transport i n plants. The synthesis begins with acetate which i s formed from pyruvate, formed from phosphoglycerate i n the Calvin cycle i n leaf tissue or v i a degradation of sugars i n seeds or f r u i t s . De novo synthesis of f a t t y acids takes place i n the chloroplasts of vegetative tissues or i n p l a s t i d s of other plant tissues. Acetate i s f i r s t e s t e r i f i e d to the -SH function of coenzyme A (CoA) v i a an enzyme, a c e t y l CoA synthetase. Coenzyme A i s an adenosine d e r i v a t i v e attached to a 4*-phosphopantetheine moiety, a chemical subunit which i s ubiquitous i n the metabolism of f a t t y acids. Formation of Acetyl and Malonyl CoA Acetyl CoA synthetase CH3COOH+ATP+C0A

• CH COCoA+AMP+PPi 3

(1)

Acetyl CoA Carboxylase CH3COC0A+ATP+CO2

•

HOOCCH COCoA+ADP+Pi 2

(2)

The d r i v i n g force f o r the reaction i s the conversion of ATP to AMP. Acetyl CoA i s then carboxylated by the enzyme acetyl-CoA carboxylase to form malonyl CoA. The carboxylase i s a c t u a l l y a multienzyme complex i n which one of the proteins i s attached to b i o t i n , the c a r r i e r of the carboxyl group. Again, the d r i v i n g force f o r the reaction i s hydrolysis of ATP. The carboxyl group added i n t h i s reaction i s l o s t i n the l a t e r condensation reactions. Although t h i s seems i n e f f i c i e n t , the condensation process i s a c t u a l l y driven by the change i n free energy r e s u l t i n g from decarboxylation. The Fatty Acid Synthase (FAS) Sequence. The subsequent reactions of f a t t y acid synthesis are shown i n Figure 1. The next sequence of reactions i s the transfer of both the acetate and malonate moieties from CoA to acyl c a r r i e r protein (ACP). ACP i s a protein on which the 4*-phosphopantetheine group already referred to i s attached to the hydroxyl of a serine which i s located approximately i n the middle of the chain (amino acid number 36 of 77) i n the ACP of E. c o l i . The ACP of both eucaryotic and procaryotic organisms has considerable homology. The ACP of E. c o l i can be used to carry acyl groups through the FAS reactions with both animal and plant FAS enzymes. Transfer of acyl groups to ACP i s e s s e n t i a l f o r the reactions to take place i n plants. The enzymes responsible are acetyl-CoA:ACP transacylase and malonyl-CoA: ACP transacylase, also c a l l e d acetyl and malonyl transferases. There i s evidence that i n eucaryotic organisms other than plants the f a t t y acid synthase enzymes are associated i n a complex i n which several functions e x i s t i n a s i n g l e protein. The synthase i n animal l i v e r has been shown to be a complex of two i d e n t i c a l



46

ECOLOGY AND METABOLISM OF PLANT LIPIDS

Figure 1. Fatty Acid Biosynthesis.





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protein subunits oriented head to t a i l i n which the various functions and the ACP are held together (7). S i m i l a r l y i n yeast the synthase i s a complex of s i x a and s i x B u n i t s . Each u n i t contains several of the s p e c i f i c functions of the system. In E. c o l i and other procaryotes the various enzymes of the FAS appear to be non-associated proteins i n the cytosol. The FAS system i n higher plants i s s i m i l a r to that of the procaryotes i n that i n d i v i d u a l functions can be i d e n t i f i e d with s p e c i f i c proteins found i n the tissue p l a s t i d s (8). A l l of the ACP i n plant leaf c e l l s i s found i n the chloroplasts (9). After conversion to acetyl-ACP and malonyl-ACP, two carbons of the malonyl-ACP are introduced v i a the condensing enzyme, B-ketoacyl-ACP synthetase. Loss of the malonyl carboxyl drives the reaction and i n the f i r s t step of the sequence acetoacetyl-ACP i s formed. The B-ketoacyl-ACP i s then reduced to B-hydroxyacyl-ACP by NADPH and the enzyme B-ketoacyl-ACP reductase. The hydroxy acid i s dehydrated to form a trans-2.3-enoyl-ACP which can be reduced by NADH or NADPH to the saturated ACP derivative (butyrate i n the f i r s t series of steps). Condensation with malonyl-ACP i s then repeated and the cycle continues to produce acyl-ACP derivatives with two a d d i t i o n a l carbon atoms u n t i l palmitoyl-ACP r e s u l t s . A second B-ketoacyl-ACP synthetase accomplishes addition of two more malonyl carbon atoms to allow the formation of stearoyl-ACP. The C -»C B-ketoacyl-ACP synthetase has been shown to be a separate enzyme since i t i s more e a s i l y i n h i b i t e d by arsenite and i s less s e n s i t i v e to the a n t i b i o t i c , cerulenin, than the B-ketoacyl-ACP synthetase forming C to C keto acids. 16

lg

Desaturation. Although the sequence produces stearate, there i s very l i t t l e s t e a r i c acid found i n plant l i p i d s . An active desaturase, the A stearoyl ACP desaturase, also found i n the chlorop l a s t or p l a s t i d s , forms a c i s double bond between Cg and C ^ Q of the carbon chain. This system requires both NADPH and ferredoxin. Stearoyl ACP i s the physiological substrate. The major product i s oleoyl-ACP, which may then be hydrolyzed or converted to oleoyl-CoA. Animals also produce o l e i c acid as a major product. The animal and plant desaturase systems diverge at t h i s point. Animal systems desaturate oleate only between the Cg double bond and the carboxyl group, while plants desaturate between the C ^ Q carbon and the co-methyl. The plant desaturase systems giving r i s e to linoleic acid and a - l i n o l e n i c acid are not yet well characterized. This i s because the desaturases are membrane-bound and have not been i s o l a t e d . Current indications are that phosphatidyl choline with o l e i c acid at the 2-position i s the substrate f o r the 18:1-»18:2 reaction (2). There i s evidence (10) that the 18:2-»18:3 reaction takes place i n the chloroplast and that the substrate i s monogalactosyldiacylglyceride (MGDG). 9

Formation of Acyl L i p i d s Synthesis of polar and non-polar l i p i d s i n plants as well as animals i s v i a the g l y c e r o l phosphate or Kennedy pathway (5). In t h i s pathway 3-sn-glycerol phosphate, formed by reduction of dihydroxyacetone phosphate, Is, acyJ.atecL by. acyl CoA i n the 1- and

American uhemical Society Library 1155 16th St., N.W.

In Ecology and Metabolism ofD.C. Plant Lipids; Washington, 20036Fuller, G., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 1987.



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2-positions. The transferases accomplishing t h i s reaction appear to be d i f f e r e n t since there i s a preference f o r saturated acids i n the 1-position and shorter chain or less saturated acids i n the 2-position. The d i a c y l phosphatidic acid produced may then be hydrolyzed to d i a c y l g l y c e r o l and further acylated to t r i g l y c e r i d e or i t may undergo reactions to form phospholipids or g l y c o l i p i d s . Formation of phospholipids i s accomplished by reaction of d i a c y l g l y c e r o l or phosphatidic acid with c y t i d i n e diphosphate (CDP) or CDP derivatives of ethanolamine or choline. Galactosyl glycerides are formed by reaction of d i a c y l g l y c e r o l with u r i d i n e diphosphate (UDP) derivatives of galactose. A plant model system In our work we were seeking a plant system with which some of the s p e c i f i c reactions, e.g., desaturation and acyl l i p i d formation, could be studied. Lemnaceae (duckweeds) are small aquatic plants which grow r a p i d l y , reproduce vegetatively and have a r e l a t i v e l y simple morphology (11). They grow autotrophically i n inorganic medium, heterotrophically i n the dark with a carbon source, or photoheterotrophically. Because of t h e i r small s i z e and simple growth requirements they can r e a d i l y be maintained i n axenic cultures, and hence, are an excellent model system f o r study of plant metabolism. Although Lemna minor and other duckweed species have been used extensively f o r biochemical studies, r e l a t i v e l y l i t t l e has been reported concerning f a t t y acid metabolism i n Lemna. We proposed to determine differences, i f any, i n l i p i d metabolism under photoautotrophic, photoheterotrophic and t o t a l l y heterotrophic growth conditions. Methods and procedures Lemna minor i n n o n - s t e r i l e culture was treated with Clorox s o l u t i o n d i l u t e d 9:1 with water. A f t e r four minutes i n t h i s solution the fronds were washed twice with s t e r i l e water and placed i n a s t e r i l e solution of modified Hillman M medium (11) to which 1% sucrose had been added. A f t e r growth i n t h i s s t e r i l e culture medium, plants were repeatedly transferred throughout the experiments to 1) inorganic Hillman M medium; 2) Hillman M medium supplemented with 1% sucrose; and 3) Hillman M medium with 1% sucrose supplemented with 600 mg/liter tryptone and 100 mg/liter yeast extract. Lemna was grown i n s t e r i l e f l a s k s (2800-ml Fernbach f l a s k s and 125 ml Erlenmeyer f l a s k s ) under continuous l i g h t at ca. 180 ixmol m~ sec.~*. The f l a s k s were stationary and maintained at ambient temperature of the laboratory. Cultures i n sucrose and sucrose-tryptone medium were also grown i n the dark. Light-grown plants were harvested a f t e r two weeks i n the culture f l a s k , dark-grown cultures a f t e r 6 weeks. Fronds were ground and extracted with 3:2 hexaneisopropanol and a portion of the l i p i d s converted by methanol-H2S04 e s t e r i f i c a t i o n to esters for gas chromatography (12). Other portions of the extracted l i p i d s were separated by 3-directional TLC (13) to determine d i s t r i b u t i o n of l i p i d s . Some 0.4g samples (wet wt.) of Lemna minor fronds were incubated i n 1.5 ml of medium under the same conditions as they were grown ( l i g h t , dark, d i f f e r e n t 2


4.

FULLER AND STUMPF

49

Fatty Acids in Plants 14

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media) with one uCi of 1- C acetate (sodium a c e t a t e - l - C , 0.133 •uCi/uD. Fatty acid d i s t r i b u t i o n was measured using a radioactive detector and a mass detector on a gas chromatograph (DEGS column) or by c o l l e c t i o n of fractions from an HPLC column and l i p i d d i s t r i b u t i o n by autoradiography of the TLC plates. Results


Table I reports the d i s t r i b u t i o n of f a t t y acids i n Lemna minor growr Table I.

Fatty Acids i n Lemna Minor

Fatty Acid (%)

Medium Emergence before 16:0 Inorganic ( l i g h t ) Sucrose ( l i g h t ) Tryptone ( l i g h t ) Sucrose (dark) Tryptone (dark)

3

12 2 9 4 1

16:0 47 27 25 37 27

Unknowns emerging before 18:0 18:0 7 tr tr 2 tr

3 3 tr 3 2

18:1 tr 6 2 14 16

18:2

18.3

7 21 27 23 28

24 38 37 17 25

Determined by GC of f a t t y acid methyl esters. i n d i f f e r e n t media i n l i g h t and dark. Growth i n inorganic medium was less than h a l f the rate of photoheterotrophic growth i n sucrose and sucrose- tryptone. (Doubling time ca. 5 days vs. 36-48 hrs.) This i s r e f l e c t e d i n a slower de novo synthesis of f a t t y acid and a high accumulation of 16:0 and lower f a t t y acids. Among the C^g acids there were high levels of 18:3 acid i n the autotrophic culture, perhaps indicating that the chloroplasts were very active. Sucrose and sucrose-tryptone i n the l i g h t gave somewhat more normal r a t i o s of 18-carbon acids with l i n o l e n i c acid s t i l l predominating. The plants grown i n the dark had lower than normal 18:3 acids, probably r e f l e c t i n g the absence of normal chloroplast organelles. Labelled acetate incorporation. Lemna fronds grown i n l i g h t and dark i n sucrose medium were incubated with 1 uCi of 1-^C acetate f o r two, four and s i x hours. The most pronounced differences were at s i x hours; r e s u l t s are shown i n Table I I . With light-grown plants, incorporation of acetate was rapid and r e l a t i v e C a c t i v i t y of the various acids c l o s e l y approached the composition r a t i o s of these acids. In the dark, however, l a b e l l e d 18:2 acid was low and 18:3 was p r a c t i c a l l y not formed i n s i x hours, i n d i c a t i n g low 18:2 desaturase a c t i v i t y and extremely low 18:3 14


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Table I I .

C-Acetate Incorporation into Fatty Acids of Lemna Grown i n Sucrose Medium 6-Hour Incubation Light-grown

a

Dark--grown

a

16:0 &


16:0 &

% Total Fatty A c i d s % of

14

c C Activity

b

18:1

18:2

18:3

18:1

18:2

18:3

33

21

38

33

24

33

20

23

28

43

15

3

Plants were incubated under the same l i g h t and dark conditions under which they were grown. k By GC of methyl esters from Lemna grown on sucrose medium. By c o l l e c t i o n and counting of f r a c t i o n s from HPLC separations of f a t t y acid isopropylidenehydrazides. activity. Thus, f o r rapid incorporation of acetate an active photosynthesizing system appears necessary. L i p i d s were separated into classes by t h r e e - d i r e c t i o n a l TLC (see Table I I I ) . The incorporation of acetate into the l i p i d s i n l i g h t was rapid. In the dark, C - a c e t a t e was not quickly incorporated into the galactol i p i d s . This f a c t correlates with the slow desaturation to a - l i n o lenate and i s not s u r p r i s i n g since up to 90% of the f a t t y acid i n galactolipids i s linolenate. 14

Conclusions Lemna minor was shown to be a representative plant when grown autotrophically and photoheterotrophically. L i p i d and f a t t y acid patterns were i n the range expected for plants. Growing the plants i n the dark d i d not stop f a t t y acid biosynthesis but changed the pattern i n that the amount of 18:3 acid was decreased. These r e s u l t s suggest a number of experiments to modify action of c r i t i c a l enzymes, e s p e c i a l l y the desaturases and makes Lemna a useful system f o r study of the reactions catalyzed by such enzymes. In addition to i t s a b i l i t y to grow h e t e r o t r o p h i c a l l y i n the dark, Lemna minor i s e a s i l y grown i n s t e r i l e culture, a useful aspect when studies could be influenced by microbial-plant interactions. Moreover, large quantities of an active growing tissue under completely controlled conditions can be made a v a i l a b l e to the investigator.




Table I I I .

L i p i d Class

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14C-Acetate Incorporation into L i p i d s of Lemna a Grown i n Sucrose Medium

LiRht-Rrown 14 C Lipids Total Lipids

Dark-srown Total L i p i d s

C Lipids

4-4-4-44-4-

++-++ +++ 4-++ +

Phospholipids PC PE PG PA PI

4-4-4-44-4+4+ 4-

4-4-44-4444-

4-+++ 4-4-

4-

+ +

+

Glycolipids MGDG DGDG

4-44-4-

Sulfolipid SQDG Netural L i p i d s FFA TG MG & DG SE

4444-

4 4-

4-4444-

After 6-hour incubation i n dark or l i g h t . From autoradiogram of TLC plate and charring of t o t a l l i p i d plate. Abbrevations PC = Phosphatidylcholine; PE = Phosphatidylethamolamine, PG = Phosphatidylglycerol; PA = Phosphatidic acid; PI Phosphatidylinositol, MGDG - Monogalatosyldiglyceride; DGDG = Digalactosyldiglyceride; SQDG = Sulfoquinovosyldiglyceride; TG = T r i g l y c e r i d e ; MG = Monoglyceride; DG = Diglyceride; SE = S t e r o l Ester; FFA = Free f a t t y acid. Q u a l i t a t i v e inspection of charred spots on TLC plate autoradiogram, 4- = very l i g h t , 4-4-4-4- very heavy.


and of

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Acknowledgments The authors wish to thank Elaine Tobin f o r suggesting modifications to Lemna media and Raymond Pacovsky for preparation of Figure 1. Thanks are also due to Gisela Travis and Emily Grey f o r typing t h i s and several other manuscripts i n the book.


Literature Cited 1. Stumpf, P. K. in "Fatty Acid Metabolism and Its Regulation", Numa, S., Ed., Elsevier, New York, 1984, Chap. 6. 2. Roughan, P. G. and Slack, C. R. 1982, Ann. Rev. of Plant Physiol. 33:97-132. 3. Jeffcoat, R., 1977, Biochem. Soc. Transactions 5:811-818. 4. Hitchcock, C. and Nichols, B. W., "Plant Lipid Biochemistry", Academic Press, London, 1971. 5. Gurr, M. I. and James, A. T., "Lipid Biochemistry. An Introduction", 3d edition, Chapman and Hall, London, 1980. 6. Bishop, D. G. in "Biosynthesis and Function of Plant Lipids", Thomson, W. W.; Mudd, J. B. and Gibbs, M. Eds., Amer. Soc. of Plant Physiologists, Rockville, MD, 1983, pp. 86-89. 7. Wakil, S. J., Stoops, J. K. and Joshi, V. C., 1983, Ann. Rev. Biochem. 52: 537-579. 8. Stumpf, P. K., Shimata, T., Eastwell, K., Murphy, D. J., Liedvogel, G., Ohlrogge, J. B. and Kuhn, D. B. in "Biochemistry and Metabolism of Plant Lipids", Wintermans, J. F. G. M. and Kuiper, P. J. C. Eds., Elsevier Biomedical Press i,B.V. (1982), pp. 3-11. 9. Ohlrogge, J. B., Kuhn, D. N. and Stumpf, P. K. (1979) Proc. Nat. Acad. Sci. U.S.A. 76: 1194-1198. 10. Murphy, D. J., Harwood, J. L., Lee, K. A., Roberto, F., Stumpf, P. K. and St. John, J. B. (1985), Phytochemistry 24: 1923-1929. 11. Posner, H. in Methods in Developmental Biology, Witt, F. H. and Wessels, N. K., Eds., Crowell, New York, 1967, pp. 301-317. 12. American Oil Chemists Sociey. Official and Tentative Methods. 3d ed. 1984. Method No. Ce2-66. 13. Kramer, J. K. G., Fouchard, R. C. and Farnworth, E. R., 1983, Lipids, 18: 896-899. RECEIVED

September 26, 1986


Ecology and Metabolism of Plant Lipids - American Chemical Society

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