Eur. J Biochem IZ9, (1983)

Transcription

1 Eur. J Biochem IZ9, (1983) (K) FEBS 1983 Specificities and Selectivities of Glycerol-3-Phosphate Acyltransferase and Monoacylglycerol-3-Phosphate Acyltransferase from Pea and Spinach Chloroplasts Margrit FRENTZEN, Ernst HEINZ, Thomas A. McKEON, and Paul K. STUMPF Botanisches Institut der Universitat zu Koln; and Department of Biochemistry and Biophysics, University of California, Davis (Received July 30/September 20, 1982) In addition to acyl-coa, purified glycerol-3-phosphate acyltransferases from pea and spinach chloroplasts can also use acyl-(acyl-carrier protein), acyl-acp, as a substrate for glycerol 3-phosphate acylation. The enzyme fractions showed absolute specificity for glycerol 3-phosphate as acyl acceptor. Dihydroxyacetone phosphate was ineffective. Glycerol 3-phosphate was almost exclusively acylated at the C-1 position. If mixtures of palmitoyl- ACP, stearoyl-acp and oleoyl-acp were offered, the oleoyl group was preferred. These data fully agree with previous experiments on these enzymes carried out with various acyl-coa thioesters. Kinetic data determined with different acyl-acps as substrates are consistent with the observed fatty acid selectivity for the oleoyl group. Double labelling experiments with mixtures of oleoyl-acp and oleoyl-coa demonstrated a preference for ACPthioesters. Monoacylglycerol-3-phosphate acyltransferase, localized in the envelope of chloroplasts, can also utilize acyl- ACP as substrate. Envelope fractions of spinach as well as of pea showed a high specificity for the palmitoyl group when ACP-thioesters or CoA-thioesters were offered and directed this acyl group to the C-2 position of the glycerol backbone. Results from competition experiments with ['4C]palmitoyl-ACP and [3H]palmitoyl-CoA indicate that the membrane-bound acyltransferase preferably uses ACP-thioesters for the acylation of 1 -acylglycerol 3-phosphate. According to fatty acid selectivities and specificities the main product of the recombined acyltransferase systems in chloroplasts of 16: 3 plants as well as 18: 3 plants is a phosphatidic acid with the oleoyl group at C-I and the palmitoyl group at C-2 whereas molecules with the oleoyl group at both positions are not synthesized. Under appropriate conditions the phosphatidic acid formed by the soluble acyltransferase and spinach envelope was rapidly converted to monogalactosyl diacylglycerol in the presence of UDP-galactose. In analogous assays with acyltransferase and envelope from pea only a low proportion of labelled phosphatidic acid was converted via diacylglycerol to monogalactosyl diacylglycerol. In both systems monogalactosyl diacylglycerol synthesized in the presence of C16:o-thioesters and C18:l-thioesters carried C18:1 at C-1 and CI~:O at C-2 in agreement with the fatty acid selectivity of the acyltransferase systems. Glycerolipids of leaves mostly contain Clb- and Cle-acyl groups which are specifically distributed between the C-I and C-2 position of the glycerol backbone. Short-time labelling experiments in vivo with so-called 16: 3 plants revealed that the lipids of chloroplasts mainly carry the oleoyl group at C-1 and the palmitoyl group at C-2 [I]. During similar experiments with 18:3 plants, this molecular species was hardly labelled in galactolipids which instead carried identical mixtures of Cis-acyl groups at both positions [I]. Chloroplasts contain a soluble glycerol-3-phosphate acyltransferase [2,3] and an envelope-bound monoacylglycerol- 3-phosphate acyltransferase [3], which catalyze the first and Ahhrcviutions. ACP, acyl-carrier protein ; Mes, 4-morpholine ethanesulphonic acid ; Mops, 4-morpholine/propanesulphonic acid ; in the shorthand for fatty acids the first number denotes the number of carbon atoms and the number following the colon denotes the number of double bonds. Enzymcs. Acyl-CoA : sn-glycerol-3-phosphate 0-acetyltransferase (EC ). acyl-coa: I-acyl-sn-glycerol-3-phosphate 0-acyltransferase (EC ). phosphatidate phosphatase (EC ), UDP-galactose: 1,2-diacylglycerol 3-0-galactosyltransferase (EC ). second acylation reaction in the biosynthesis of diacylglycerol 3-phosphate. This observation poses the question whether these enzymes are involved in the control of the abovementioned fatty acid distribution in chloroplast lipids of 18:3 and 16:3 plants. Some properties of the glycerol-3-phosphate acyltransferase have been described [4]. With acyl-coa thioesters as acyl-donors purified acyltransferase fractions of chloroplasts from both types of plants as represented by pea (1 8 : 3 plant) and spinach (16: 3 plant) specifically catalyze the acylation of the C-1 position of glycerol 3-phosphate and preferably insert the oleoyl group. Preliminary experiments revealed that the enzyme fractions accept acyl-acp as well as acyl-coa thioesters. The acyl-acps are the direct products of fatty acid biosynthesis in chloroplasts [5]. In the present paper we show that the first acyltransferase displays similar specificities with the alternative acyl-donors, and that in addition ACP-thioesters rather than CoA-thioesters were used for the acylation reaction. Furthermore the substrate specificities of the monoacylglycerol-3-phosphate acyltransferase from the 18:3 plant, pea, and the 16:3-plant, spinach, were investigated and found to be identical.

2 630 MATERIALS AND METHODS Chemicals [I -'4C]Palmitoyl-ACP (57 Ci/mol), [l-'4c]stearoyl-acp (56.5 Ci/mol) and [l-'4c]oleoyl-acp (56 Ci/mol) were made and purified as described by Rock and Garwin [6]. The buffer was exchanged on Sephadex G-15 to 10mM Mes-NaOH buffer ph 5.9 containing 0.1 % bovine serum albumin. [9,10-3H]Palmitoyl-CoA (330 Ci/mol) and [9,10-3H]oleoyl-CoA (450 Ci/mol) were prepared enzymatically with a Pseudomonas acyl-coa synthetase purchased from Sigma. The reaction mixture contained 0.1 M Mops-NaOH buffer ph 7.4, 50 pm CoA, 5 mm ATP, 9 mm MgClz, 1 mm dithiothreitol, 0.05 'i: Tween, 42 pm [9,10-3H]oleic acid (450 Ci/mol) or 60 pm [9,10-3H]palmitic acid (330 Ci/mol) and about 0.2 nkat acyl-coa synthetase in a total volume of 1 ml. Incubations were carried out at 37 C for 20 min. Free fatty acids were extracted by light petroleum after acidifying the reaction mixture with acetic acid. The resulting [3H]acyl-CoA samples contained less than 5 of free fatty acids. Biochemicals and auxiliary enzymes were obtained from Boehringer, Serva and Sigma, while radioactive substances were purchased from New England or Amersham. Plants Pisum sativum L. cv. Kleine Rheinlanderin was grown in the greenhouse, and Spinacia olerucea L. cv. Mona Lisa in the garden. Chloroplast Envelope Purified chloroplasts of pea and spinach leaves were prepared [7] and separated into envelope and thylakoid fractions according to the method of Joyard and Douce [8]. Purification of Glycerol-3- Phosphate Acyltrunsferase The purification of acyltransferase from pea leaves was carried out as described before [4] while the purification from spinach leaves was modified in the following way. The concentrated Sephadex G-100 eluate was dialyzed over night against 50 mm sodium acetate buffer ph 6 containing 5 mm dithiothreitol. Protein precipitated during dialysis was removed by centrifugation and the resulting supernatant solution was applied to a column of CM-Sephadex (2.5 x29cm) equilibrated in dialysis buffer. The column was rinsed with this buffer and enzymic activity eluted with a linear gradient mixed from 200 ml 0.1 M NaCl and 200 ml 0.3 M NaCl in the acetate buffer. Peak fractions were pooled, concentrated by ammonium sulfate precipitation and further purified by electrofocussing using Ampholine ph 4-6 as previously described [4]. The ph of the peak fractions was adjusted to 7 by Tris base, and acyltransferase samples were stored at -20 "C in 50 glycerol. Storage under these conditions led to no detectable decrease in activity over a 6-month period. Enzyme Assays Glycerol-3-phosphate acyltransferase activity was determined in standard assays [4]. Assays with acyl-acp were conducted in a similar way. The reaction mixture of 80 p1 contained 0.25 M Mops-NaOH buffer ph 7.4, 1 mm glycerol 3-phosphate, pg bovine serum albumin, purified acyltransferase fractions of spinach or pea ( pg protein) and varying amounts of ['4C]acyl-ACP as given in the text. Monoacylglycerol-3-phosphate acyltransferase was measured by adding purified envelope fractions of pea and spinach chloroplasts (20-80 pg protein) to the reaction mixture used to measure glycerol-3-phosphate acyltransferase. In some cases purified acyltransferase and glycerol 3-phosphate were substituted by 25 mm 1 -oleoylglycerol3-phosphate, dispersed in SO mm Mops-NaOH buffer at ph 7.4. The lipid Substrate was made by incubating glycerol-3-phosphate acyltransferase with oleoyl-coa under standard assay conditions. For biosynthesis of monogalactosyl diacylglycerol in vifro, the incubation with both acyltransferases was carried out for 15 min, then the ph of the reaction mixture was adjusted to 8.2 followed by an additional incubation in the presence of 0.35 mm unlabelled UDP-galactose. Phosphatidate phosphatase activity was measured according to Joyard and Douce [9]. Lip id Analysis Assays were stopped and lipids extracted as described for glycerol-3-phosphate acyltransferase tests [4]. Lysophosphatidic acid, phosphatidic acid and free Fatty acids were separated by thin-layer chromatography in chloroform/pyridine/formic acid (50 : 30 : 7) [ 10 J while monogalactosyl diacylglycerol was separated from other lipids in chloroform/methanol/water (65 : 25 : 4). Positional analyses of monoacylglycerol 3-phosphate were carried out as described before but the monoacylglycerol isomers were separated on boric-acid-impregnated plates in chloroform/acetone (85 : 25) [Ill to achieve a good separation of 2-acylglycerol and free fatty acids. Positional analyses of other glycerolipids have been described before [12]. The molecular species of lipids were separated by thinlayer chromatography on AgN03-impregnated silica gel G plates. Phosphatidic acid was methylated with diazomethane and analyzed by thin-layer chromatography in chloroform/ methanol (96:4). Fatty acid esters were prepared by transmethylation and separated by argentation-chromatography on AgN03-impregnated plates [4] or by reversed-phase partition chromatography on siliconized plates [13]. RESULTS AND DISCUSSION Purification of Glycerol-3- Phosphate Acyltrunsferases The purification procedure of glycerol-3-phosphate acyltransferase from spinach leaves yielded enzyme fractions with specific activities of 11 nkat/mg protein under standard assay conditions. This represents a 45-fold-higher purification when compared to previous data [4] and the activities are of the same order as those determined for the purified enzyme fractions from pea leaves (29-53 nkat/mg protein). The enzyme fractions were not pure when analyzed by polyacrylamide gel electrophoresis. However, they did not contain interfering enzymic activities such as phosphatases, acyl-coa thioesterase or monoacylglycerol-3-phosphate acyltransferase although a low acyl-acp thioesterase activity was detected. In glycerol-3-phosphate acyltransferase tests with ['4C]oleoyl- ACP as acyl donor [14, IS], at most 7 'i: of the acyl component of oleoyl-acp was found in the fraction of free fatty acids (Fig. 1 b).

3 ~ ~ 63 1 h Fig. 1. Acyl-acceptor spec@ciiy of' glycerol-3-phosphate acyliransjerase. Reaction mixtures contained 2 pm labelled oleoyl-acp and 1 mm of (a) dihydroxyacetone phosphate, (b) glycerol 3-phosphate. Assays with enzyme fractions of pea instead of spinach gave similar results. Incubations lasted for 5 min. (1 = start, 2 = lysophosphatidic acid, 3 = free fatty acids) Table 1. Positional analjsis oj monoacyirlycero1 3-phosphate,formed by the enriched ~Iyrerol-3-pho.sphaie acjkmsferase,fractions,fiorn glycerol 3-phosphate and ucyl-acp Palmitoyl-ACP, oleoyl-acp and an equimolar mixture of both were each offered at 5 pm. Incubations lasted for 1-5 min. I-Acylglycerol 3-phosphate formed is given as percentage of total amount of I-acyl and 2-acyl isomers formed Acyl-ACP Cih citr C1h:oiCis:l (1/1) Speciji'citie.s of the Acylation Reaction with Acyl-ACP Glycerol-3-phosphate acyltransferases of pea and spinach chloroplasts possess a specificity for the acyl acceptor. As depicted in Fig. 1, dihydroxyacetone phosphate did not serve as acceptor substrate (Fig. 1 a) whereas glycerol 3-phosphate was rapidly acylated to monoacylglycerol 3-phosphate (Fig. 1 b). The analysis of the isomeric composition of lysophosphatidic acid formed during incubations with either palmitoyl- ACP or oleoyl-acp or mixtures of both are given in Table 1. In contrast to similar experiments with glycerol-3-phosphate acyltransferases of Escherichia coli [I61 the same positional distributions were obtained at different incubation times ranging from 1-5 min. These results reveal high positional specificity of the purified enzyme fractions since labelled acyl groups were almost exclusively transferred to the C-1 position of the glycerol backbone. During the above-described incubations only slightly higher acylation rates were observed with oleoyl-acp as compared to palmitoyl-acp which indicate that the enzyme fractions do not have a pronounced fatty acid specificity when LO % C18., in mixtures of C16:o-/C - ACP Fig. 2. FatijJ acid selectivity of glycerol-3-phosphate acyirrunsfhrci.st.. Figures represent the percentage of CIS. in 1 -acylglycerol 3-phosphate formed during 2-min incubations with 4.5 pm ACP mixtures of varying palmitoyl-acp and oleoyl- ACP thioester proportions the acyl-acps were offered separately. If, however, mixtures of these ACP-thioesters were offered, the analysis of the htty acid composition of the corresponding reaction products revealed a fatty acid selectivity (Fig.2 and 3). (The term specificity is used to denote comparisons of acylation rates measured in the presence of single acyl-donors while the term selectivity is used when the acylation rates are measured in the presence of thioester mixture.) As depicted in Fig. 2 the acyltransferases preferably used oleoyl groups even with excess palmitoyl-acp in the substrate mixture. This preference for oleoyl groups was not only found in comparison to Clb:o (Fig. 3a) but also to C18:o (Fig. 3 b). The fatty acid composition of 1 -acylglycerophosphate formed by the spinach enzyme from equimolar mixtures of C~~:O-ACP/C~~:~-ACP or C18:o- ACP/Clg:l-ACP consisted of about 90 "/, of oleic acid in both cases (Fig. 3a, b). The enzyme fraction from pea (isoelectric point 6.3 [4]) catalysed the acylation reactions somewhat less selectively (Fig.2 and 3). Offering an equimolar mixture of palmitoyl-acp and stearoyl-acp, a slight discrimination against CI~:~ is evident (Fig. 3c). The results presented here are in full agreement with previous experiments carried out with various acyl-coa thioesters [4]. Since acyl-acp thioesters in contrast to acyl-coa compounds do not form micelles, the observed preference for oleic acid from mixtures of thioesters cannot be ascribed to differences in the physical state of the substrates. Therefore, this selectivity appears to be a characteristic property of the glycerol-3-phosphate acyltransferases from chloroplasts. Kinetic Data of Glycerol-3-Phosphate Acyltransferase Typical Michaelis-Menten kinetics were obtained using acyl-acp as substrate. The kinetic data determined for glycerol-3-phosphate acyltransferase of spinach are consistent with the fatty acid selectivity shown in Fig. 2 and 3. As summarized in Table 2, kinetic constants for both acyl- ACP and glycerol 3-phosphate vary in relation to the nature of

4 ~~~~ ~~~ ~- ~ ~ ~~~~ ~ 632.;!.; Q, J A h L, z Cn.c_ O L a n* 2-l 75A jlll -0 l n.- 5a 2 c COLD -7 Em *. Fig. 3. Fati), ucid.w/ec~iii~iti~ of ~l~~~cerol-3-pho.~/~hafe acj,lrransferuse. Thin-layer radioscans of Fatty acid methyl esters obtained by methanolysis from various acyl-acp mixtures and from 1 -acylglycerol 3-phosphate formed from these mixtures by purified enzyme fractions of Spincrcia and Plsunr. Incubations lasted for 3 min and the reaction mixtures contained 0.25 M Mops-NaOH buffer ph 7.4, 30 pg bovine serum albumin, 6 pm [I4C]acyl- ACP, acyltransferase fractions (Spinacia, 0.3 pg protein; Pisum, 0.7 pg protein) and glycerol 3-phosphate concentrations of (a, b) 1 mm, and (c) 20 mm or? m'd Table 2. Kinetic constants jkr various substrates qf gl~~cerol-3-phosphate acy Itransf erase Initial velocities were determined in I-min assays as described under Materials and Methods at various concentrations of the one substrate while the other substrate was hold at a fixed level of about 10 K,. In spinach, in contrast to oleoyl-acp, V values of the saturated acyl-acps vary with the amount of bovine serum albumin added to the reaction mixture, while the K,, values were not influenced. The values given in the table were determined in assays containing 36 pg albumin Enzyme Acyl donor K, K, V source acyl-acp glycerol 3-P PM nkat/mg protein Spinacia palmitoyl-acp stearoyl- ACP oleoyl-acp Pisum palmitoyl-acp oleoyl-acp Table 3. Dependenre of fatty acid selectivity om gl.vcero1 3-phosphut~ ('oncentrations Fatty acid compositions of 1-acylglycerol 3-phosphate formed by acyltransferases of Pisum and Spinacia (0.3 pg protein) from 5.5 pm C18:I- ACP/C16,,,-ACP mixture ( CI~:I/C~~:~ = 2: 23) and glycerol 3-phosphate at two concentrations (1 mm and 0.3 mm) during 2 min Glycerol-3-P Enzyme source Incorporation into lysophosphatidic acid of o/ mm /" C18 1 Clh P isum Sp inaciu Pisum Spinacia the acyl group of the thioester. Therefore the selectivity of the C-I acylation does not only depend on the concentrations of the various acyl-acp thioesters but also on the concentration of glycerol 3-phosphate. The effect of glycerol 3-phosphate concentrations on fatty acid selectivity can be demonstrated at nonsaturating levels of CI~:~-ACP (Table 3). Palmitoyl-ACP and oleoyl-acp were offered in a mixture so that the concentration of each substrate was about 1.5-times its K,, whereas the two different glycerol 3-phosphate concentrations of 1 mm and 0.3 mm provide V conditions for C18:1-ACP, but not for C16:o-ACP (Table 2). In accordance with the kinetic data the incorporation of the oleoyl group into 1 -acylglycerol 3-phosphate was markedly increased by decreasing the concentration of glycerol 3-phosphate. Glycerol 3-phosphate concentrations up to 15 mm hardly influenced the preferred acylation with the oleoyl group if the acyltransferase was provided with saturating levels of CIS, ACP but not C16:o-ACP. Offering equimolar mixtures of C16,o-ACP and Cls,l-ACP at concentrations in the range of 6 pm, the corresponding reaction products consisted of about 90 oleic acid independent of the glycerol 3-phosphate concentrations. Preference of ACP-thioesters or CoA-thioesters by Glycerol-3-Phosphate Acyltran.ferase Since purified gl ycerol-3-phosphate acyltransferase fractions of chloroplasts can use acyl-acp as well as acyl-coa for

5 A / /" Y a L 7.2 [Cle., -CoAIIpM) - a L 7.2 L x._ IC,, -CoAI(pM) { ' L 6 a [C,e 1 -COA/CI,,~-ACPI Equimolar mixture IpM) CoA C,, from ACP U Fig. 4. Prefbrence of ACP-thioesters or CoA-thioesters by glvcerol-3-phosphate ucyltransferuse. Labelled 1 -acylglycerol 3-phosphate formed by the purified enzyme fraction of spinach chloroplasts (0.3 pg protein) from 1 mm glycerol 3-phosphate and different concentrations of (a) [3H]oleoyl- CoA in the absence and presence of 2 pm oleoyl-acp; (b) [3H]oleoyl-CoA in the presence of 2 pm ['4C]oleoyl-ACP; (c) an equimolar mixture of [3H]oieoyl-CoA and ['4C]oleoyi-ACP. The incubations lasted for 1 min the acylation reaction [4] we next determined whether the enzyme fractions display a preference for ACP-thioesters or CoA-thioesters. To this end double labelling experiments with ['4C]oleoyl-ACP and [3H]oleoyl-CoA were conducted. As shown in Fig.4a the acylation rates depending on oleoyl-coa concentrations were strongly inhibited in the presence of 2 pm oleoyl-acp. On the other hand the addition of oleoyl-coa to reaction mixtures containing oleoyl-acp did not influence the acylation rate with ACP-thioesters (Fig. 4 b). Increasing concentrations of oleoyl-coa at a constant level of oleoyl-acp resulted in an increasing formation of tritium-labelled lysophosphatidic acid (Fig. 4b). But even at a 3.5-fold excess of oleoyl-coa in the thioester mixture the resulting monoacylglycerol 3-phosphate carried 62 % oleoyl groups from acyl-acp (Fig. 4 b). Similar results were obtained with the enzyme fractions from pea chloroplasts. The preference for ACP-thioesters became even more obvious if an equimolar mixture of oleoyl-coa and oleoyl-acp was offered at different concentrations (Fig. 4c). During these incubations glycerol 3-phosphate was almost exclusively acylated by acyl groups of ACP-thioesters, since the reaction products contained about 93 "/, ''C-labelled acyl groups independent of the thioester concentrations offered (Fig. 4c). These experiments clearly demonstrate that in the presence of acyl-acp and acyl-coa, ACP-thioesters were preferably or even exclusively used as substrate for the acylation of glycerol 3-phosphate. In view of the results presented above a knowledge of the concentrations of thioesters in chloroplasts is of interest. The concentration of total ACP in spinach chloroplasts has been estimated to be about 8 pm [17]. Similar values can be calculated from the incorporation rates of ['4C]acetate into acyl- ACP while the labelling rate of acyl-coa was one tenth only (P. G. Roughan, personal communication). These determinations indicate that in plastids an excess of ACP-thioesters compared to acyl-coa is available for the acylation reaction. Furthermore the acyl-coa synthetase has recently been localized in the outer envelope membranes of spinach chloroplasts whereas the acyl-coa thioesterase is a membranebound enzyme of the inner envelopes [ 181. These data and the results of the double-labelling experiments (Fig. 4) suggest that acyl-acp rather than acyl-coa serves as acyl-donor for the biosynthesis of 1 -acylglycerol 3-phosphate in chloroplasts (Fig. 7). The acyl-acp fraction labelled by ['4C]acetate during incubations of isolated chloroplasts consisted of 7 % C14:0, 28 % C16:0, 20 "/, and 36 "/, C18:1 (P. G. Roughan, personal communication), whereas the fatty acid analysis of acyl-acp thioesters isolated from spinach leaves gave even lower amounts of saturated acyl groups [19]. From these ACP-mixtures the glycerol-3-phosphate acyltransferase will preferably direct oleoyl groups to the C-1 position of glycerol 3-phosphate (Fig. 2), almost independent of the glycerol 3-phosphate concentration in chloroplasts, since the mixture only contains saturating levels Of C18:1-ACP. Specificities of' Monoucylglycerol-3- Phosphate Acyltransfivme Monoacylglycerol-3-phosphate acyltransferase, which is firmly bound to the envelope membranes of chloroplasts, catalyzed the acylation of lysophosphatidic acid to phosphatidic acid. This enzyme has been usually measured with acyl-coa as acyl donor [3]. The addition of isolated envelope membranes to a glycerol- 3-phosphate acyltransferase assay with labelled ACP-thioesters as acyl donors resulted in the formation of phosphatidic acid which was labelled at both positions (Table 4). This labelling of phosphatidic acid demonstrates that not only the soluble glycerol-3-phosphate acyltransferase but also the membrane-bound monoacylglycerol-3-phosphate acyltransferase can use ACP-thioesters as acyl donors. The second acyltransferase did not accept glycerol 3-phosphate as acyl-acceptor since no lysophosphatidic acid and hardly any phosphatidic acid was formed when glycerophosphate acyltransferase was omitted from the reaction mixture. In contrast to phosphatidic acid phosphatase, a further enzyme firmly bound to envelope membranes [9], the second acyltransferase can use not only substrate synthesized in situ, but also exogenously added substrate. In the presence of acyl-acp, 1 -oleoylglycerol 3-phosphate dispersed in buffer was rapidly converted to phosphatidic acid by envelope membranes. In order to investigate the fatty acid selectivity of the second acyltransferase from pea and spinach chloroplasts, palmitoyl-

6 ~~ ~~ ~~ ~ ~~~ ~ ~~ ~~~~~ ~ ~~~ 634 Table 4. Analysis of futty acids at the C-I and C-2 position of lysophosphatidic acid andphosphatidic acid formed from glycerol3-phosphate and an equimolar mixture qfpalmitoyl-.4 CP and oleoyl-acp by the reconstituted acyltransferase systems of pea and spinach ch1oroplast.s Assay conditions : 7.5 pm [I4C]acyl-ACP, 1 mm glycerol 3-phosphate, glycerol 3-phosphate acyltransferase of pea or spinach (0.6 pg protein) and envelope of pea (80 pg protein) or spinach (20 pg protein), incubation time 1.5 min Enzyme source Product analysed Fatty acid incoporated total C-1 position (2-2 position Cin I c16 n Cin i Ci6 n Cln 1 Cih n Pi sum lysophosphatidic acid phosphatidic acid Spinacia lysophosphatidic acid phosphatidic acid C II Fig. 5. Reaction products.formed hy the reconstituted acyltransj&me.s.vstem of chloroplasts from lahelled ACP-thioesters and glycerol 3-phosphatc,. 5-min assays consisted of 0.25 M Mops-NaOH buffer ph 7.4, 1 mm glycerol 3-phosphate, 30 pg bovine serum albumin, purified acyltransferase from pea (0.7 pg protein), envelope fraction from pea chloroplasts (80 pg protein) and 7.5 pm (a) oleoyl-acp, (b) palmitoyl- ACP, (c) equimolar mixture of both thioesters. (1 = start, 2 = lysophosphatidic acid, 3 = phosphatidic acid, 4 = front) ACP and oleoyl-acp were offered individually as well as in an equimolar mixture. The radioscans of Fig. 5 reveal that palmitoyl-acp is a much more effective substrate for the second step of diacylglycerol 3-phosphate biosynthesis than oleoyl-acp. Furthermore, the fatty acid composition at the C-2 position of phosphatidic acid formed during incubations with the thioester mixture (Fig. 5) consisted of 97 % palmitic acid. Similar results were obtained with acyl-coa instead of acyl-acp as substrate and no differences with respect to fatty acid selectivity could be detected using envelope fractions of spinach or pea chloroplasts as enzyme source (Table 4). These experiments revealed that monoacylglycerol-3-phosphate acyltransferases from chloroplasts of 16 : 3 plants as well as 18 : 3 plants possess a high preference for palmitoyl groups. When mixtures of palmitoyl-thioesters and oleoylthioesters were offered, C16:,,-acyl groups were exclusively directed to the C-2 position of the glycerol backbone. Since phosphatidic acid formation was severely reduced when other thioesters were offered rather than palmitoyl thioester, the second acyltransferase does not only show fatty acid selectivity but in contrast to the first acyltransferase, even a pronounced fatty acid specificity (Fig. 5). On the other hand monoacylglycerol-3-phosphate acyltransferase did not display a specificity with respect to the acyl group in the acceptor lysophosphatidic acid, since l-oleoylglycerol 3-phosphate as well as 1 -palmitoylglycerol 3-phosphate were used as substrate for the C-2 acylation. To investigate whether monoacylglycerol-3-phosphate acyltransferases prefer CoA-thioesters or ACP-thioesters as acyl donor, envelope fractions of spinach chloroplasts were incubated in the presence of 25 pm I-oleoylglycerol3-phosphate with 1.6 pm ['4C]palmitoyl-ACP and different concentrations of [3H]palmitoyl-CoA (0.8 pm, 1.6 pm, 3.2 pm). The resulting phosphatidic acid carried at least 90% of the incorporated radioactivity at the C-2 position and consisted of 92 % of [14C]palmitate transferred from the ACP-thioester when acyl-acp was offered in a concentration twice that of acyl-coa. This percentage was reduced to 89 % and 85 % when the CoA concentration increased up to twice that of ACP. These results indicate that the membrane-bound acyltransferase preferably uses ACP-thioesters for the acylation, especially if the low amount of acyl-coa present in these organelles is taken into consideration. In conclusion acyl-acp rather than acyl-coa is the physiological acyl donor for glycerolipid biosynthesis in chloroplasts (Fig. 7). The first step in the reaction sequence is the acylation of the C-I position of glycerol 3-phosphate by the soluble acyltransferase while the membrane-bound acyltransferase catalyzes the acylation of the C-2 position. The main product of the combined acyltransferase system in chloroplasts of 16:3 plants as well as 18:3 plants is a phosphatidic acid with oleoyl groups at the C-1 position and palmitoyl groups at the C-2 position of the glycerol backbone

7 ~~~~~~~~ ~ ~ ~ ~ ~~ ~ ~~~ ~ ~ ~ ~ ~ ~ Table 5. Analysis of fatty acid.7 at the C-I and C-2 position of lysophosphatidic acid, phosphatidic acid and nzonogalac fosyl diacyl~l~~cero1,formrtl by fhr reconstituted enzyme system of spinach chloroplasts Assays contained 0.25 M Mops-NaOH buffer ph 7.4, 1 mm glycerol 3-phosphate, 30 pg bovine serum albumin, purified acyltransferase of spinach (0.6 pg protein), spinach envelope (20 pg protein) and 7.5 pm of an equimolar mixture of C16,0-ACP and C18:1-ACP. After an incubation of 15 min the ph of the reaction mixture was shifted to 8.2 and then incubated for 60 min in the presence of 0.35 mm UDP-galactose 635 Product analysed Fatty acid incorporated total C-1 position C-2 position cl8 1 C*6 0 CIS 1 c16 0 cis 1 c16 0 x ~ ~ ~ ~ ~ ~ ~ Lysophosphatidic acid Phosphatidic acid Monogaldctosyl didcylglycerol (Fig.7). This phosphatidic acid is used as precursor for the biosynthesis of galactolipids (see below) and may be also used for the biosynthesis of other glycerolipids in chloroplasts such as sulpholipid and phosphatidylglycerol since it reflects the fatty acid pattern of these lipids found after short-time labelling experiments with whole leaves [20] or isolated chloroplasts [21]. Biosynthesis of Galactolipids de novo As shown by Joyard and Douce [Y], the envelope membranes of spinach chloroplasts contain a specific alkaline phosphatidic acid phosphatase which dephosphorylates phosphatidic acid to diacylglycerol, the substrate for subsequent galactosylation reactions in the envelope membranes [3,22]. Under appropriate conditions biosynthesis de novo of monogalactosyl diacylglycerol could be demonstrated in reaction mixtures containing purified acyltransferase and envelope of spinach chloroplasts. The reaction products labelled in assays with an equimolar mixture of [14C]palmitoyl-ACP and ['4C]oleoyl-ACP in the presence of glycerol 3-phosphate and UDP-galactose were isolated and rechromatographed on AgNOs-impregnated plates. In each case only one molecular species was formed as the main product. The analysis of fatty acids at the C-I and C-2 position of the glycerol backbone revealed (Table 5) that the precursor phosphatidic acid as well as the product monogalactosyl diacylglycerol are mainly molecular species with oleate at C-I and palmitate at C-2. Analogous experiments with envelope fractions of pea instead of spinach chloroplasts resulted in the same products but gave significant differences in reaction rates. Under conditions giving rapid conversion of phosphatidic acid to monogalactosyl diacylglycerol by envelope fractions of spinach chloroplasts [Y], pea envelopes produced only a small proportion of acyl-labelled monogalactosyl diacylglycerol. But the same membrane preparation of pea chloroplasts rapidly incorporated labelled UDP-galactose into galactoselabelled galactolipids. The slow formation of acyl-labelled monogalactosyl diacylglycerol by the enzyme system of pea can be related to a certain extent to the phosphatidic acid phosphatase activity in 18:3 plants. As depicted in Fig.6, envelope fractions of Pisum showed very low phosphohydrolase activity at conditions optimized for the enzyme of spinach envelope. As observed Fig. 6. Phosphatidic acid phosphatase activity of envelope memhrunes of' spinach and pea chloroplasts. Envelope fractions (50 pg protein) labelled with phosphatidic acid synthesized in situ (0.4 nmol, specific activity lo4 dis. x inin-' x ninol-') were incubated for 20 min at ph 9 in a total volume of 40 p1. (1 = start, 2 = phosphatidic acid, 3 = monoacylglycerol, 4 = diacylglycerol) with the spinach enzyme [Y] the activity of pea enzyme was decreased if the ph of the reaction mixture was lowered. The analysis of monogalactosyl diacylglycerol formed de novo by the reconstituted enzyme system of pea gave a fatty acid pattern similar to that determined for the lipid synthesized by the spinach system. Similar results were obtained by [I4C]acetate incubations with isolated chloroplasts of 16: 3 plants and 18 : 3 plants 123,261. This fatty acid distribution in monogalactosyl diacylglycerol reflects the pattern of monogalactosyl diacylglycerol labelled in vivo in the case of 16 : 3 plants [20] whereas monogalactosyl diacylglycerol from 18: 3 plants carries an identical set of Cl8 fatty acids at both positions [l]. Consequently, according to the fatty acid selectivities of the first acyltransferases and the fatty acid specificities of the second acyltransferases described here, biosynthesis de now of glycerolipids in chloroplasts forms products with predomi-

8 c GLYCEROL 3 -PHOSPHATE C16.0 -CoA - ACP C16 C18:o -COA Ct8:o - ACP C18 -COA C18:1 -ACP LYSOPHOSPHATIDIC ACID %8:1 16:0Ip C16:o{L18:1 PHOSPHATIOIC ACID DIACVLGLYCEROL UONOGALACTOSYL DIACYLGLVCE ROL Fig. 7. Biosynthesis de novo of molecular species of glycerolipids in chloroplasts of 16: 3 plants and 18: 3 plants according to the results presented in thispaper. The selectivities and specificities with respect to acyl groups and coenzymes of the soluble glycerol-3-phosphate acyltransferase and the membrane-bound I-acylglycerol-3-phosphate acyltransferase are symbolized in the scheme nantly C,g-acyl groups at C-1 and C16-acyl groups at C-2 from which the Cl* fatty acids are excluded (Fig. 7). In contrast the main molecular species of galactolipids in 18 : 3 plants which are also present in 16:3 plants i.e. a Cl$Zlg combination cannot be synthesized within the chloroplasts but presumably are derived from outside these organelles. We gratefully acknowledge financial support by the Deutsche For-.scliunRsRemein.scha~t and we thank K. Wrage for expert technical assistance. In addition support from an National Institutes of Health grant GM administered by PKS is also acknowledged. REFERENCES 1. Heinz, E., Siebertz, H. P., Linscheid, M., Joyard, J. & Douce, R. (1979) in Advunces in the Biochemistry and Physiology of Plant Lipids (Appelqvist, L. A. & Liljenberg, C., eds) pp , Elsevier/North-Holland Biomedical Press, Amsterdam. 2. Bertrams, M. & Heinz, E. (1976) Planta (Berl.) 132, Joyard, J. & Douce, R. (1977) Biochim. Biophys. Acta, 486, Bertrams, M. & Heinz, E. (1981) Plant Physiol. 68, Stumpf, P. K. (1977) in MTP International Review uf Science, Biochemistry oflipids II(Goodwin, T. W., ed.) vol. 14, pp , Butterworths, London. 6. Rock, C. O.&Garwin, J. L.(1979)J. Biol. Chem.254, Haas, R., Siebertz, H. P., Wrage, K. & Heinz, E. (1980) Planta (Berl.) 148, Joyard, J. & Douce, R. (1976) Physiol. Veg. 14, Joyard, J. & Douce, R. (1979) FEBS Lett. 102, Hauser, G. & Eichberg, J. (1975) J. Biol. Chem. 250, Thomas, A. E., Scharoun, J. E. & Ralston, H. (1965) J. Am. Oil Chem. Soc. 42, 789 ~ 12. Siebertz, 8. P., Heinz, E., Linscheid, M., Joyard, J. & Douce, R. (1979) Eur. J. Biochem. 101, Malins, D. C. & Mangold, H. K. (1460) J. Am. Oil Chem. Soc. 37, Shine, W. E., Mancha, M. & Stumpf, P. K. (1976) Arch. Biochem. Biophys. 172, Ohlrogge, J. B., Shine, W. E. & Stumpf, P. K. (1978) Arch. Biochem. Biophy,~. 18Y, Rock, C. O., Goelz, S. E. & Cronan, J. E., Jr (1981) J. Bid. Chem. 256, Ohlrogge, J. B., Kuhn, D. N. & Stumpf, P. K. (1979) Proc. Nut1 Acad. Sci. USA, 76, Dome, A. J., Block, M. A., Joyard, J. & Douce, R. (1982) in Biochemistry and Metabolism OfPlunt Lipids (Wintermans, J. F. G. M. & Kuiper, P. J. C., eds) pp , Elsevier/North-Holland Biomedical Press, Amsterdam. 19. Sanchez, J. & Mancha, M. (1980) Phytochemistry, IY, Joyard, J., Douce, R., Siebertz, H. P. & Heinz, E. (1980) Eur. J. Biochem. 108, Sparace, S. A. & Mudd, J. B. (1982) in Biochemistry and Metabolism of Plant Lipids (Wintermans, J. F. G. M. & Kuiper, P. J. C., eds) pp , Elsevier/North-Holland Biomedical Press, Amsterdam. 22. Van Besouw, A.. Wintermans, J. F. G. M. & Bogernann, G. (1981) Biochim. Biophy.s. Acta, 663, Heinz, E. & Roughan, P. G. (1982) in Biochemistry and Metabolism of Plant Lipids (Wintermans, J. F. G. M. & Kuiper, P. J. C., eds) pp , Elsevier/North-Holland Biomedical Press, Amsterdam. 24. McKee, J. W. A. & Hawke, J. C. (1979) Arch. Biochem. Bioph.n. IY7, Roughan, P. G., Holland, R. & Slack, C. R. (1980) Biochem. J. 188, Drapier, D., Dubacq, J. P., Tremolieres, A. & Mazliak, P. (1982) Plant Cell Physiol. 23, M. Frentzen and E. Heinz, Botanisches Institut der Universitat zu Koln, GyrhofstraBe, 15, D-5000 Kdn 41, Federal Republic of Germany T. A. McKeon and P. K. Stumpf, Department of Biochemistry and Biophysics, University of California at Davis, Briggs Hall, Davis, California, USA 95616