The Biosynthesis of Polyunsaturated Fatty Acids in Plants 1 M.I. GURR, Unilever Research Laboratory, Colworth House, Sharnbrook, Bedford, England
are released during the reaction, and to examine the catalytic site of the enzymes. This could be done by obtaining the enzymes in soluble form and purifying them. Our efforts in this direction have been mainly confined to animal desaturases. The work described here typifies another approach, working with rather crude enzyme systems but using a variety of tailormade substrates to probe the specificity of the desaturase reaction. Whole cell cultures or subcellular preparations of the green alga Chlorella vulgaris, were used as our model for higher plants because fatty acid and lipid composition and general lipid metabolism are very similar to plant leaf chloroplasts. However, there are two main differences: Chlorella yields subcellular preparations capable of synthesizing polyunsaturated fatty acids (1,2) which no plant leaf chloroplast preparation has done, and it can directly desaturate exogenous stearic acid (3). Stearic acid or stearoyl-CoA could be directly desaturated to oleic acid in a whole range of animal preparations, but no such reaction could be demonstrated in higher plants. Monoenoic fatty acid synthesis in plants, the so called plant pathway, seemed to be fundamentally different from that in animals. The nearest saturated precursor to oleate was myristic acid, 14:0 (4). The idea of the plant pathway was disproved in two ways. (a) In our laboratory a leaf preparation was incubated anaerobically with labeled
This paper is a review of some of the work being done at the author's laboratory. The phospholipids and glycolipids of the alga, Chlorella vulgaris, have been implicated in fatty acid transformations such as chain elongation and desaturation. Labeling studies with [ 14C] acetate have shown that newly synthesized galactosyl glycerides have mainly saturated fatty acids. Subsequent to de novo synthesis, a series of alterations of fatty acid structure takes place within the same glycolipid molecules. The specific incorporation of [14C]oleic acid into Chlorella phosphatidyl choline provides a convenient model system for studying the lipid dependent desaturatiou of oleic to linoleic acid. The inhibitor of fatty acid desaturation, sterculic acid, only inhibits the conversion of oleate into linoleate if added before the precursor fatty acid has been incorporated into a complex lipid. Studies with isomeric monoenoic fatty acids have suggested that there are two enzymes which catalyze the formation of linoleic from oleic acid. One measures the position of the second double bond from the carboxyl group, the other, from the methyl end of the chain. The latter enzyme probably requires the complex lipid substrate. I NTROD UCTI ON
The objective of our work is to show how, in desaturation, the two hydrogen atoms C14:1
FIG.1. The direct desaturation and plant pathways for oleic acid biosynthesis. X is a hypothetical acyl carrier such as CoA or ACP; E, chain elongation steps; D, desaturation and T, transacylation steps.
FIG. 2. Fatty acid transformations in monogalactosyl dig]yceride (MGDG) subsequent to de novo
sy n thes"ls f rom [2- 14C]acetate. The ordinate represents the radioactivity of each fatty acid as a percentage of the total radioactivity in MGDG.
PLANT UNSATURATED ACID SYNTHESIS
5 o < n.16:1 18:2
FIG. 3. Differences in the specific activity of monoenoic acids in different MGDG species after incubation with [2-14C]acetate for 2 hr. [1,1] represents species containing two monoenes (two double bonds per molecule); [1,2] represents species containing one monoene and one diene (three double bonds per molecule). acetate. Labeled stearic acid accumulated and, on transferring the incubation to aerobic conditions, the radioactivity in stearic acid declined, giving place to an equal amount of oleic acid (5). (b) Bloch (6) demonstrated that although spinach chloroplasts would not d e s a t u r a t e stearic acid or stearoyl-CoA, stearoyl-ACP was converted into oleic acid. Thus, plants can perform direct desaturation of stearic acid, and there is no fundamental difference between plant and animal pathways to monoenoic fatty acids. Why then, is endogenous stearic acid itself not desaturated? Bloch's experiments suggest that the substrate must be bound to ACP. We believe that a stearoyl-CoA:ACP acyl transferase is lacking in higher plants. This gives rise to two metabolic states of stearic acid. One is synthesized in situ from acetate and is desaturated to oleic acid without becoming mixed with the other pool which is available to exogenously added stearate. Chlorella has proved to be a useful tool because the transferase appears to be inducible by transferring heterotrophically grown cells to autotrophic conditions (Fig. 1) (1). I N C O R P O R A T I O N OF F A T T Y ACIDS INTO LIPIDS A N D D E S A T U R A T I O N
One cannot study fatty acid synthesis and desaturation without studying the related area of complex lipid synthesis since a cell does not synthesize fatty acids as an end in itself. Because nonesterified fatty acids (NEFA) are extremely inhibitory to most enzymes and must of necessity be maintained at a low con.centration, only minute levels are usually found in cells. The object of fatty acid synthesis is to
FIG. 4. Time course of incorporation of [1-14C] oleate into PC and its s.ubsequent desaturation to linoleate. PC, phosphatidyl choline; NEFA, nonesterified fatty acid. The ordinate represents radioactivity as a percentage of the maximum label incorporated into PC (about 70% of the added label). provide the lipophilic moiety of a complex lipid whether that lipid be used as an energy storage compound or a membrane component. This must be kept in mind during the study of fatty acid biosynthesis and desaturation. In the incorporation of labeled acetate into Chlorella lipids (7), all lipids become labeled, but there is a group of three lipids, p h o s p h a t i d y l glycerol (PG), phosphatidyl choline (PC) and monogalactosyl diglyceride (MGDG) which incorporate radioactivity very rapidly. Once the maximum uptake of label has occurred, the total amount of radioactivity in these lipids remains constant for a considerable time. Initially most of this label is present in the saturated acids 16:0 and 18:0. As the incubation proceeds, even though the total amount of label in a given lipid remains constant, the label in 16:0 and 18:0 falls while the label in the corresponding monoenes first increases and then declines, and so on through to the trienoic acids. Figure 2 illustrates this point using the MGDG and the C 1 6 acids as an example. Another way of looking at this phenomenon is to fractionate the MGDG into individual molecular species (8). When this is done, the specific activities of individual fatty acids vary considerably from one species to another. For example the specific activity of monoenoic acids in species containing two double bonds per molecule is more than six times greater than in species containing three double bonds per molecule after a 2 hr incubation (Fig. 3). This situation can arise if a series of alterations of fatty acid structure takes place within the MGDG molecule subsequent to de novo synthesis from saturated fatty acids. During the course of the incubation these fatty acids are desaturated and the label passes steadily from species of lower degree of desaturation to those more highly unsaturated. LIPIDS, VOL. 6, NO. 4
* ' 1 8 : 1 - PC kiN 2
E~ E; E: E:
*'18:1 - PC t lair
"18:1.'18:2 after desaturation
FIG. 7. The distribution of radioactivity among +~ 18:1- NEFA
FIG. 5. The conversion of endogenous [ 14C] oleoyl-PC into [14C] linoleoyl-PC. The diagrams represent radiochemical gas liquid chromatography traces. For the sake of simplicity, mass traces are not shown. In another type of experiment, 14Clabeled oleic acid rather than acetate was used as precursor (2). Unlike acetate, which is incorporated into most of the phospholipids and glycolipids of Chlorella, about 70% of the oleic acid is incorporated extremely rapidly into phosphatidyl choline (Fig. 4). Desaturation occurs at a slower rate than incorporation, and when it begins after an initial lag, much of the oleic acid has already been incorporated into PC. The linoleic acid which is formed by desaturation is located exclusively in the PC fraction. Labeled linoleic
FIG. 6. The time courses for the desaturation of oleoyl-PC and oleoyl-CoA. LIPIDS, VOL. 6, NO. 4
diglyceride species derived from PC. In the upper diagram, the species were derived from cells grown in the presence of [1-14C]oleic acid under dark anaerobic conditions. In the lower diagram, the species were derived from cells grown in the presence of [1-14C]oleic acid under light aerobic conditions. Under these conditions there was an overall conversion of oleic into linoleic acid of 46%. The PC was purified from the lipid extract by DEAE cellulose chromatography, hydrolyzed by phospholipase C and the resulting diglycerides separated by argentation thin layer chromatography in 4% ethanol in chloroform. O, origin; SF, solvent front. The nomenclature for the different species is explained in the legend of Figure 3. acid is not detected in the NEFA fraction until well after 30 rain when a major part of the label has already been incorporated into lipid and desaturated. Experiments such as these suggested that the desaturation of oleic acid was taking place on the lipid. In other words, the lipid is the substrate, or a "carrier molecule" for the desaturase. To test this hypothesis, two types of experiments were performed (2). In the first, PC labeled with t4C-oleic acid was synthesized by growing Chlorella in the presence of the labeled acid, in the dark, under an atmosphere of nitrogen, so that no desaturation could take place. After purification, the labeled lipid was sonicated in a buffer to give an almost water clear micellar dispersion which was incubated with a Chlorella chloroplast preparation. Linoleoyl-phosphatidyl choline was formed in a yield of 3%. The linoleate could not have come from oleic acid released from the lipid by phospholipase action, because the chloroplast fraction contains no activating enzyme and c a n n o t desaturate NEFA. Although this approach gave us the information that oleoylPC is a substrate for the desaturase, it is nevertheless technically very difficult to do this kind of experiment because of problems involved in solubilizing the substrate. In addition, PC micelles, in the required concentration, were inhibitory to the enzyme. Another approach
PLANT UNSATURATED ACID SYNTHESIS
E C12:1-~-~ ~ C18:1 -
C 2-~-~.~-~.~ C12:0.~.~ ..~ C 18:0 9 18:1C,"
t b 18:1-E
T / ~lnhibition
P k ~
18:2 FIG. 8. Hypothetical pathways for the formation of linoleic acid in Chlorella vulgaris. The fatty acids are assumed to be incorporated into phosphatidyl choline by the acylation of lysophosphatidyl choline catalyzed by specific acyl transferases. was more successful (2). Chlorella PC was labeled with oleic acid in the dark under N 2 as described. A chloroplast fraction was then prepared and all but a very small amount of unincorporated oleic acid washed out. The result was a desaturase preparation labeled with phosphatidyl choline in situ, but unable to desaturate through lack of oxygen and light. When these preparations were brought into the light and aerated, 10% of oleoyl-PC was transformed into linoleoyl-PC during a 2 hr incubation (Fig. 5). As a final check on this reaction the rate of desaturation of lipid-bound oleate was compared with that of oleoyl-CoA. As Fig. 6 shows, the oleoyl-PC is the better substrate. Analysis of the molecular species of PC labeled with 14C_olei c acid alone indicated that four of the seven different PC species which contain oleic acid had incorporated the label (Fig. 7). After exposure of cells labeled with oleoyl-PC to desaturating conditions there was a movement of label towards more highly unsaturated species. As in the acetate experiment, the specific activities of oleate and linoleate differed widely in different species. No single species appeared to be involved exclusively in the desaturation reaction, although the species containing two oleic acid molecules has the highest turnover and may be quantitatively the most important species as far as linoleate production is concerned (9). The current concept of the coupling of oleate desaturation with PC biosynthesis is summarized in Figure 8. One possibility is that the oleoyl-PC acts as an acyl donor to the enzyme, the acid is desaturated to linoleate as an acyl enzyme intermediate, followed by transfer of the acyl group back to PC. The whole reaction
C18:0 FIG. 9. Different proposals for the site of inhibition of desaturation by sterculic acid. See legend of Figure 1 for explanation of symbols. would be highly compartmentalized, ensuring that the linoleate remains associated with PC. Alternatively, the fatty acid may remain bound in ester linkage with PC throughout the reaction (2). At present, there is no conclusive evidence distinguishing between these two mechanisms. TWO POSSIBLE M O D E S OF D E S A T U R A T I O N
So far, there is less evidence that stearic acid desaturation is as intimately linked with phospholipid or glycolipid synthesis as is oleic acid desaturation. Indeed, nearly all the evidence we have accumulated suggests that the mechanisms for desaturating stearate and oleate are quite different. One of the ways of probing enzyme mechanisms and alternative metabolic routes is by the use of inhibitors. A compound which specifically inhibits desaturation is the cyclopropene acid, sterculic acid:
CH3(CH2)TC = C - (CH2)7COOH Neither the site nor the mechanism of action of sterculic acid is yet known. In relation TABLE I Effect of Sterculic Acid on the Desaturation of Oleic Acid Dark-N2, hr 16
TABLE II Formation of Methylene Interrupted Dienes: Specificity With Regard to Double Bond Position of the Monoene Substrate ~9 "~A9,12-diene 15:1 16:1 17:1 18:1 19:1
o)6 co7 co8 (.o9 col0
16:1 16:1 17:1 18:1 19:1
to Chlorella, sterculic acid, at a level twice that of the substrate concentration, inhibits 72% of the desaturation of stearate to oleate, whereas the desaturation of oleate to linoleate is hardly affected (10). Alternatively, a dose 100 times that of the substrate is needed to inhibit oleate desaturation by the same a m o u n t , and thus the mechanism of this inhibition appears to be of a different nature f r o m the stearate-oleate inhibition. While the f o r m a t i o n of oleic acid f r o m stearic acid is dramatically inhibited by sterculic acid, the inhibition is decreased if labeled acetate is the precursor of oleate. A similar p h e n o m e n o n is seen in rat (11) or hen (12) liver. Reiser and J o h n s o n believe that the action of sterculic acid is directly on the stearate desaturase enzyme. When this is inhibited by sterculic acid an alternative pathway, according to Reiser, is able to bypass stearate desaturation. This bypass involves desaturation at the C12 level (Fig. 9). In our opinion, the inhibition is at the level of an acyl transferase (possibly involved in transferring stearic acid to stearoyl-ACP), which prevents labeled stearic acid f r o m reaching the desaturase c o m p l e x (Fig. 9) (13). Stearate arising f r o m acetate, however, is coupled directly to the desaturase c o m p l e x and so bypasses the blocked acyl transferase. A n y small inhibition of oleate f o r m a t i o n f r o m acetate is probably of a less specific kind. E x p e r i m e n t s were made to clarify some of these points. TABLE III
LIPIDS, VOL. 6, NO. 4
A7 15:1 (,O8 A7 18:1 coil A l l 18:1 ~,o7 A8 18:1 t.o10 A10 18:1 t.~8 A12 18:1 606 ---> A9,12-18:2 by A9 desaturase
If some desaturations depend on the prior i n c o r p o r a t i o n of the substrate fatty acid into a c o m p l e x lipid, is it possible that sterculic acid is incorporated preferentially into lipid, effectively blocking the desaturation? Sterculic acid is incorporated at a rate which is equal to, or slightly greater than, oleic acid and very m u c h greater than stearic acid. This might account, therefore, for the relatively large quantities required to inhibit oleate desaturation by a c o m p e t i t i v e effect for sites on the lipids, but could not explain the more specific effect on stearate desaturation. In a n o t h e r experiment the following operations were carried out: (a) A control incubation consisted of incubating 14C-oleate in the dark under N 2 for 17 hr then transferring the incubation to the light in air for a further 5 hr. (b) The same e x p e r i m e n t was done anaerobically for 16 hr. Sterculic acid was then added and incubated a further hour, followed by a light-aerobic phase of 5 hr. (c) Finally, sequence (a) was repeated exactly except that sterculic acid was present f r o m the very beginning (Table I). Only in (c) was there significant inhibition. These results are interpreted to mean that sterculic acid has no effect on oleate desaturation when added after all the labeled precursor has been incorporated into lipids, as it certainly has after 16 hr. SPECI F I C I T Y OF DESATURASES WITH R E G A R D TO DOUBLE BOND POSITION
As far as the desaturase which introduces the first double bond is concerned, all saturated fatty acids f r o m 12:0 to 19:0 yield a m o n o e n o i c acid with the double b o n d in the 9,10 position (14). Since the double b o n d is always in the same position regardless of chain Inhibition, % length, we conclude that the active site of the e n z y m e measures the site for desaturation from 8 the carboxyl end of the molecule. (In Chlorella 53 the situation is complicated by the existance of
Effect of Sterculic Acid (CPE) on the Formation of a Dienoic Acid From A9-18:1 or A12-18:1 Monoenes Desaturation, %
exogenous- C18.0 FIG. 10. Interdependent pathways of fatty acid synthesis, desaturation and complex lipid biosynthesis in Chlorella vulgaris. an enzyme which produces a cis-7-monoene, but for simplicity this discussion will be confined to the cis-9 series). The question now arises, does the enzyme which introduces the second double bond measure from the carboxyl end of the chain, from the methyl end, or solely with reference to the first double bond? To investigate this, three series of monoenoic fatty acids were synthesized. The members of a given series differed in chain length, but series (a) had the double bond in the carboxyl-9 position, series (b) in the 60-9 position, while in series (c) the bond was neither co- nor carboxyl-9 (D. Howling et al., in preparation.) The results of these studies, shown in Table I, are as follows: The chain length of the monoene is not critical within the range CIs-C19. Outside this range desaturation is severely restricted. Within this range all A9 and w9 monoenes were desaturated to yield a methylene-interrupted diene. Monoenes with no double bonds in these positions were not substrates, with the exception of A12-18:1, which was converted into linoleic acid. The yield of linoleic acid from A9-18:1 (609, oleic TABLE IV Tritium-Carbon Ratios of Lipid Components of Chlorella After Incubation With [ I-I4c] Oleic and [9,103-H2]Oleic Acidsa 3H/14C Phospholipids Glycolipids Triglyceride NEFA Substrate Diglyceride Water soluble aMean of 10 values +s.d. 0.5.
8.2 8.9 8.9 10.0 10.3 11.3
acid) was at least twice that from any other substrate. The interpretation of these results is that there are two enzymes capable of producing a diene from a monoene. Each enzyme will only insert the second double bond to produce a methylene-interrupted fatty acid. One enzyme inserts the second double bond by measuring from the carboxyl group. This is the enzyme responsible for producing cis, eism9,1 2-dienes from A9 monoenes. The other has methyl end control and is the enzyme which produces cis,cis-co,6,9-dienes from 609 monoenes. Since oleic acid is both a A9 and 609 monoene, it is capable of being desataurated by both enzymes. Exactly the same results have been obtained with a higher plant, the castor bean. Since the carboxyl end is locked up in the lipid while the fatty acid chains project and their ends are available for interaction with the enzyme, it is probable that the lipid-linked desaturase is the methyl end controlled enzyme. As mentioned earlier, A12-18:1 is anomalous in that it is the only monoene which does not have a double bond in the 609 or A9 position, yet produces a diene when incubated with Chlorella (Table II). The diene in this case is A9,12-18:1, linoleic acid, and the second double bond is in the A9 position. The desaturation of this substrate was probably catalyzed by the stearate desaturase, especially as the same reaction is catalyzed by animal preparations which cannot form linoleate from oleate. To check this we made use of the fact that low concentrations of sterculic acid, which have little effect on oleate desaturation, severely inhibit stearate desaturation. We incubated a mixture of tritium-labeled oleic acid with carbon-labeled A12-18:1 with and without 1 mM sterculic acid. The results (Table III) indicate that, while the desaturation of oleic acid was inhibited only 8%, the desaturation of A12-18:1 was inhibited 53%. This is consistent with the view that linoleic acid is formed from LIPIDS, VOL. 6, NO. 4
A1 2-18:1 by the stearate desaturase. Although these two substrates behaved differently as far as desaturation was concerned, their distribution among Chlorella lipids at the end of the incubation was rather similar. ISOTOPE DISCRIMINATION
When we compared the incorporation of [ 3 H ] A 9 - 1 8 : l and [ 1 4 C ] A 1 2 - 1 8 : l , we alsoran a parallel incubation using a mixture of tritiated and 14C_labele d A9-18: 1. The incorporation of these two different forms of oleic acid into the different lipids of Chlorella was studied by measuring the tritium-carbon ratio in each individual lipid. The conditions were such that no 18:2 formation took place. The results are shown in Table IV. The tritium-carbon ratio in the phospholipids, glycolipids and triglycerides is lower than that of the substrate, but higher in the case of the diglycerides and water soluble radioactivity, presumably unincorporated fatty acid or thiol esters. This indicates a preferential incorporation of 14C-substrate into the complex lipids, leaving the residual NEFA enriched in the tritium-labeled substrate. A similar phenomenon has been observed in the case of the incorporation of stearic acids into hen liver lipids. The explanation is not clear at present, but is probably due to differences in the strengths of binding of the acyl chain to the transferase enzyme. The presence of two methylene groups containing pairs of tritium atoms instead of hydrogen atoms may significantly alter the binding energy of the enzyme and substrate. The enzyme reaction is sufficiently sensitive to these small changes for the effect to be reflected in differences in incorporation of the two substrates. In conclusion, these results are summarized from many types of experiments, and a coherent picture which can be fitted into an overall scheme of fatty acid biosynthesis in plants is given (Fig. 10). Polyunsaturated C18 fatty acids arise by sequential desaturation of oleic acid, which in turn arises, as we have shown, in plants as well as animals by direct desaturation of stearic acid. Normally stearic acid is the end-product of the fatty acid synthetase. Exogenously added stearic acid must be transferred to the central enzyme complex in order to be desaturated. It is this transfer step that may be the target for sterculic acid inhibition of stearate desaturation. The enzyme or enzymes which perform this desaturation fix the substrate at the carboxyl group and the active site is so placed in relation to this binding site that the double LIPIDS, VOL. 6, NO. 4
bond is inserted at the 9 and 10 positions. We believe that the stearic acid is in a bent conformation on the enzyme surface to bring the two D-hydrogens in the correct configuration for desaturation. It is interesting to speculate that at some time in evolution the A7 desaturase arose by deletion of a single amino acid from the relevant part of the desaturase polypeptide chain, which we calculate would bring the active site in the correct position to produce a A7 monoene. There are two enzymes which produce the dienoic acids. One controls the double bond position from the carboxyl group, the other controls this position from the methyl end. This may produce not only some linoleic acid from oleate but also 7,10-diene from A7-16:1 (significantly, A7-18:1 is not a substrate). Linoleate, therefore, arises from the combined effects of two enzymes. At least one of these enzymes acts on lipid-bound oleic acid and this may be the one which locates the double bond from the methyl end of the chain. Two monoenes are potential precursors of linoleic acid when incubated with Chlorella, and by studying their desaturation, differences between two enzymes, i.e., the monoeneforming and diene-forming desaturases, can be pinpointed. Oleic acid, (A9-18:1,) is desaturated in the lipid-bound form by the methylend diene-forming enzyme whereas A 12-18: 1 is desaturated by the monoene-forming enzyme. The systems for forming unsaturated fatty acids in Chlorella are very complex, despite the simple fatty acid composition of the organism. Transacylases ar e very important and inseparable from the desaturase systems because some desaturations require lipid-bound substrates. We are aware that many of our observations are open to one or even several other interpretations. Many loose ends and even fundamental points remain to be clarified. Does sterculic acid block the transacylase as we have suggested or does it in fact directly inhibit the desaturase? What are the roles of the lipidlinked and nonlipid-linked systems? Do the same considerations apply to formation of trienoic acids? We hope that current and future researches will supply some of the answers. ACKNOWLEDGMENTS This paper represents the combined researches of T. James, B. Nichols, L. Morris, D. Howling, P. Harris, E. Hammond, P. Robinson and many students. REFERENCES 1. Harris, R.V., and A.T. James, Biochim. Biophys. Acta 106:456 (1965).
PLANT UNSATURATED ACID SYNTHESIS 2. Gurr, M.I., M.P. Robinson and A.T. James, Eur. J. Biochem. 9:70 (1969). 3. Harris, R.V., P. Harris and A.T. James, Biochim. Biophys. Acta 106:465 (1965). 4. Stumpf, P.K., and A.T. James, Ibid. 70:20 (1963). 5. Harris, R.V., A.T. James and P. Harris in "Biochemistry of Chloroplasts" Vol. 2, Edited by T.W. Goodwin, Academic Press, London (1967). 6. Nagai, J., and K. Bloch, J. Biol. Chem. 241:1925 (1966). 7. Nichols, B.W., Lipids 3:354 (1968). 8. Nichols, B.W., and R. Moorhouse, Ibid. 4:311 (1969).
9. Gurr, M.I., and P. Brawn, Eur. J. Biochem. 17:19 (1970). 10. James, A.T., P. Harris and J. Bezard, Ibid. 3:318 (1968). 11. Raju, P.K., and R. Reiser, Biochim. Biophys. Acta 176:48 (1969). 12. Johnson, A.R., J.A. Pearson, F.S. Shenstone and A.C. Fogarty, Nature 214:1244 (1967). 13. James, A.T., Chem. Brit. 4:484 (1968). 14. Howling, D., L.J. Morris and A.T. James, Biochim. Biophys. Acta 152:224 (1968). [ R e c e i v e d O c t o b e r 1, 1 9 7 0 ]