Identification, Purification, and Characterization of Monoacylglycerol Acyltransferase from Developing Peanut Cotyledons*

Ajay W. Tumaney, Sunil Shekar, and Ram RajasekharanDagger

From the Department of Biochemistry, Indian Institute of Science, Bangalore 560 012, India

Received for publication, January 2, 2001, and in revised form, January 17, 2001



    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Biosynthesis of diacylglycerols in plants occurs mainly through the acylation of lysophosphatidic acid in the microsomal membranes. Here we describe the first identification of diacylglycerol biosynthetic activity in the soluble fraction of developing oilseeds. This activity was NaF-insensitive and acyl-CoA-dependent. Diacylglycerol formation was catalyzed by monoacylglycerol (MAG) acyltransferase (EC 2.3.1.22) that transferred an acyl moiety from acyl-CoA to MAG. The enzyme was purified by successive chromatographic separations on octyl-Sepharose, blue-Sepharose, Superdex-75, and palmitoyl-CoA-agarose to apparent homogeneity from developing peanut (Arachis hypogaea) cotyledons. The enzyme was purified to 6,608-fold with the final specific activity of 15.86 nmol min-1 mg-1. The purified enzyme was electrophoretically homogeneous, and its molecular mass was 43,000 daltons. The purified MAG acyltransferase was specific for MAG and did not utilize any other acyl acceptor such as glycerol, glycerol-3-phosphate, lysophosphatidic acid, and lysophosphatidylcholine. The Km values for 1-palmitoylglycerol and 1-oleoylglycerol were 16.39 and 5.65 µM, respectively. The Km values for 2-monoacylglycerols were 2- to 4-fold higher than that of the corresponding 1-monoacylglycerol. The apparent Km values for palmitoyl-, stearoyl-, and oleoyl-CoAs were 17.54, 25.66, and 9.35 µM, respectively. Fatty acids, phospholipids, and sphingosine at low concentrations stimulated the enzyme activity. The identification of MAG acyltransferase in oilseeds suggests the presence of a regulatory link between signal transduction and synthesis of complex lipids in plants.



    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Biosynthesis of diacylglycerol is shown to occur by the sequential acylation of glycerol-3-phosphate (1-3). The first enzyme in this pathway, G3P1 acyltransferase, catalyzes the formation of lysophosphatidic acid (LPA). LPA can also be synthesized by acylation followed by the reduction of dihydroxyacetone phosphate that is catalyzed by dihydroxyacetone phosphate acyltransferase (4) and the NADPH-dependent acyl-dihydroxyacetone phosphate reductase (5), respectively. LPA is shown to induce a wide range of activities in animal systems (6-8). LPA can be metabolized through dephosphorylation by a soluble LPA phosphatase to form monoacylglycerol (MAG) or acylated to phosphatidic acid (PA) by LPA acyltransferase. LPA phosphatase has been identified (9), purified (10), and cloned (11) from animal systems. PA is the precursor for diacylglycerol (DAG) and anionic phospholipids. PA phosphatase catalyzes the dephosphorylation of PA to form DAG, which is the immediate precursor for triacylglycerol, phosphatidylcholine, and phosphatidylethanolamine. DAG is also an important signal molecule that activates protein kinase C (12). DAG can also be derived directly from phospholipids by the action of phospholipase C (13).

Alternatively, DAG can be synthesized by the esterification of MAG by acyl-CoA:MAG acyltransferase (EC 2.3.1.22). This enzyme has been proposed to be important for fat absorption in human small intestine (14-16). Another acyltransferase, acyl-CoA-independent MAG acyltransferase, has been purified to homogeneity from rat intestinal mucosa (17), whereas acyl-CoA-dependent MAG acyltransferase has not been purified from any source. All the acyltransferases in these pathways are membrane-bound, and most of them use acyl-CoA as a primary acyl donor (1-3).

We identified DAG biosynthetic activity from the soluble fraction of developing peanut (Arachis hypogaea) cotyledons, and the enzyme involved was purified to apparent homogeneity by successive column chromatographic procedures and characterized. This is the first report of the purification of acyl-CoA-dependent MAG acyltransferase.


    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Materials-- [1-14C]Palmitoyl-CoA (54 mCi mmol-1), [9,10-3H(N)]trioleoylglycerol (10 Ci mmol-1), [glycerol-U-14C]PA (100 mCi mmol-1), [[2-palmitoyl-9,10-3H]phosphatidylcholine (92.3 Ci mmol-1) and [2- 3H]G3P (12 Ci mmol-1), were obtained from Perkin Elmer Biosystems. Prep grade Superdex 75 (26/60) and Superdex 200 (10/30) FPLC columns, octyl-Sepharose 4 Fast Flow matrix, and gel filtration molecular mass standards were purchased from Amersham Pharmacia Biotech. Protein assay reagents were obtained from Pierce. Thin layer chromatography plates were from Merck. All other reagents were obtained from Sigma. MAG was purified by preparative silica-TLC and quantified colorimetrically (18, 19). Field grown-developing peanut (Arachis hypogaea L.) cotyledons were harvested at 20-25 days after flowering and used either fresh or stored at -80 °C until further use.

Lipid Extraction-- Lipids were extracted from 10 g of frozen immature seeds by grinding the tissue in liquid nitrogen to a fine powder in mortar and pestle (20). The powder was extracted with 20 ml of boiling isopropyl alcohol. The mixture was then centrifuged briefly, supernatant was removed, and the extraction repeated twice. The pooled isopropyl alcohol extracts were brought to dryness on a rotary evaporator. The tissue residue was then reextracted twice with 38 ml of chloroform/methanol/10% acetic acid (1:2:0.8, v/v). After centrifugation, the supernatant was added to the isopropyl alcohol extract, and 20 ml each of chloroform and water were added to the mixture. The biphasic system was mixed and centrifuged. The lower chloroform phase was removed and dried in a rotary evaporator. The lipid residue was dissolved in chloroform/methanol (1:1, v/v) and stored at -20 °C.

Preparation of Subcellular Fractions-- Differential centrifugation was used to fractionate intracellular components (21). Either fresh or frozen immature seeds (100 g) were ground in a prechilled mortar and pestle with 10 g of acid-washed sand and 250 ml of buffer consisting of 50 mM Tris-HCl, pH 8.0, 1 mM EDTA, 10 mM KCl, 1 mM MgCl2, 1 mM beta -mercaptoethanol, 0.1 mM phenylmethylsulfonyl fluoride, 1 µg ml-1 leupeptin, and 0.25 M sucrose. The extract was passed through two layers of cheesecloth and centrifuged at 3,000 × g for 10 min. The supernatant was centrifuged at 18,000 × g for 15 min. The 18,000 × g supernatant was further centrifuged at 150,000 × g for 2.5 h. The pellets were resuspended in small volume of buffer containing 20 mM Tris-HCl, pH 7.0, and 1 mM beta -mercaptoethanol. All these operations were performed at 4 °C. All the fractions were assayed for acyltransferase activities. Protein concentrations were determined by the bicinchoninic acid method (22) using bovine serum albumin as the standard. For the purification of MAG acyltransferase, the homogenate was centrifuged directly at 18,000 × g for 30 min, and the supernatant was then centrifuged at 150,000 × g for 2.5 h. The 150,000 × g supernatant (soluble fraction) was used as the source for MAG acyltransferase.

Enzyme Assays-- The assay mixtures consisted of 50 mM Tris-HCl, pH 7.0, 20 µM [1-14C]palmitoyl-CoA (100,000 dpm), 15-45 µg enzyme, and 50 µM MAG (1-oleoyl) in a total volume of 100 µl. The incubation was carried out at 30 °C for 10 min and stopped by the addition of 400 µl of CHCl3/CH3OH (1:2, v/v). Following lipid extraction by the modified method of Bligh and Dyer (23), the lower chloroform-soluble materials were separated by TLC on 250 µm silica gel G plates either using petroleum ether/diethyl ether/acetic acid (70:30:1, v/v) or chloroform/methanol/water (98:2:0.5, v/v) as the solvent system (24). The lipids were visualized with iodine vapor, and the spots of DAG scraped off for determination of radioactivity by liquid scintillation counting.

Purification of MAG Acyltransferase-- All operations were conducted at 4 °C except the FPLC purification step, which was conducted at ambient temperature. Buffer A contained 50 mM Tris-HCl, pH 8.0, 1 mM EDTA, 100 mM KCl, 1 mM MgCl2, 1 mM beta -mercaptoethanol, and 0.1 mM phenylmethylsulfonyl fluoride.

Octyl-Sepharose Chromatography-- Solid ammonium sulfate was added to bring the soluble fraction to 1 M followed by centrifugation to make the solution clear and then loaded onto an octyl-Sepharose column (4.4 × 15 cm) that had been pre-equilibrated with 1 M ammonium sulfate in Buffer A with a flow rate of 1 ml min-1. The column was washed with the same buffer to remove unbound proteins until the effluent had very low absorbency at 280 nm. The enzyme was eluted with 200 ml of a linear-reversed gradient from 1 to 0 M ammonium sulfate in Buffer A, and fractions of 5 ml were collected. The active fractions were pooled and dialyzed against Buffer A.

Blue-Sepharose Chromatography-- Active fractions from the octyl-Sepharose were combined, dialyzed against Buffer A, and applied onto a blue-Sepharose (cibacron blue A) column. The column was eluted with a 0-1 M NaCl gradient.

Size Exclusion Chromatography-- The fractions eluted at 0.35-0.4 M NaCl from the blue-Sepharose were concentrated using a Centricon (30-kDa cut-off) concentrator and filtered. The filtrate was applied onto a preparative Superdex 75 FPLC column fitted with Bio-Rad BioLogic low-pressure chromatography system. The column was eluted with the same buffer at a flow rate of 5 ml min-1.

Palmitoyl-CoA-agarose Chromatography-- A 2.5-ml sample of palmitoyl-CoA-agarose was pre-equilibrated with Buffer A at room temperature, and the active fractions from the previous column was mixed with matrix for 90 min at 4 °C. The mixture was then poured into a column, washed with Buffer A, and eluted with Buffer A containing 0.25, 0.5, and 1 M NaCl, respectively. The MAG acyltransferase activity was eluted at 0.5 M NaCl.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Subcellular Distribution of DAG Biosynthesis in Immature Peanuts-- The subcellular distribution of DAG biosynthesis in fresh peanut cotyledons was studied by isolating the intracellular components by differential centrifugation. DAG biosynthetic capacity was found low in the membrane fraction because of the presence of active triacylglycerol biosynthesis (data not shown). However, DAG formation was higher in 150,000 × g supernatant as compared with the corresponding microsomal pellet, which is in agreement with the earlier report on developing rapeseed (25). The pattern of distribution remained the same even in the frozen tissue, but total DAG biosynthetic activity decreased 21% after freezing and thawing. These results suggested that the additional DAG biosynthetic machinery could exist in the soluble fraction.

Marker Enzyme Activities from Peanut Soluble Fraction-- There was a significant amount of DAG formation in the soluble fraction, and this could either be caused by the presence of soluble enzymes or the presence of nonsedimentable cellular membrane fragments generated during isolation procedures. Marker enzyme activities were measured to assess the extent of contamination in the soluble fraction with the membranes. The activities of succinate dehydrogenase (26), NADH cytochrome C reductase (27), and vanadium sensitive ATPase (28) were measured as the marker enzymes for mitochondrial, microsomal, and plasma membranes, respectively. About 6% (1.2 µmol min-1 mg-1), 18% (1.3 µmol min-1 mg-1) and 11% (0.55 µmol min-1 mg-1) activities of these enzymes were detected in the soluble fraction, respectively. These results indicated that the membrane contamination in the soluble fraction was not significant.

Incorporation of [14C]Palmitoyl-CoA into Diacylglycerol-- Effect of [14C]palmitoyl-CoA incorporation into DAG was measured both in the soluble and the membrane fractions of immature peanuts (Fig. 1). Maximum activity was observed at 10 µM in the soluble fraction and at 20 µM palmitoyl-CoA in the membrane fraction. Exogenous MAG did not alter the rate and the pattern of incorporation into DAG. To determine the amounts of MAG in the immature peanut seeds, the total lipid was extracted as described under "Experimental Procedures." The amounts of MAG in the immature seeds and the soluble fraction were calculated to be about 134 nmol/g fresh weight and 2.6-3.1 nmol/mg protein, respectively.



View larger version (14K):
[in this window]
[in a new window]
 
Fig. 1.   Effect of palmitoyl-CoA on diacylglycerol biosynthesis. Incorporation of [14C]palmitoyl-CoA into DAG was carried out in the presence of 50 µM MAG in soluble (open circle ) and membrane () fractions. Each point is the average of two independent experiments.

Formation of DAG by the Acylation of MAG-- DAG can be synthesized either by the dephosphorylation of PA or by the direct acylation of MAG. To find out the contribution of each step to the total DAG pool, peanut soluble fraction was treated with various concentrations of NaF to inhibit phosphatase activity, and these were used for the incorporation of [14C]palmitoyl-CoA into DAG. Initially, we studied the effect of various concentrations of NaF on the dephosphorylation of PA in the membrane fraction and found that there was no formation of DAG at 20 mM NaF (Fig. 2A). As shown in Fig. 2B, there was only a 25-38% decrease in the incorporation of [14C]palmitoyl-CoA into DAG in the soluble fraction in the presence of 20 mM NaF suggesting that the DAG formation was independent of PA dephosphorylation activity. NaF-treated membrane fraction showed a profound inhibition of DAG formation indicating the presence of NaF sensitive PA dephosphorylation activity (Fig. 2C). These results suggest that the soluble fraction has NaF-insensitive and PA dephosphorylation-independent DAG biosynthesizing activity in the soluble fraction of immature peanuts.



View larger version (13K):
[in this window]
[in a new window]
 
Fig. 2.   Generation of diacylglycerol. A, effect of NaF on the dephosphorylation of [14C]PA to DAG was determined in the soluble (open circle ) and membrane () fractions. The incorporation of [14C]palmitoyl-CoA in the presence (open circle ) or absence () of 20 mM NaF was performed with soluble (B) and membrane (C) fractions into DAG in the absence of exogenous acyl acceptor. Each point is an average of three determinations.

Evaluation of Different Plant Tissues for MAG Acyltransferase Activity-- Incorporation of [14C]palmitoyl-CoA into DAG was studied in the soluble and the particulate fractions from the tissues of leaf and hypocotyl of peanut and immature seed and leaf tissue of castor (Ricinus communis L.). The MAG acyltransferase activity was not detected in the soluble fractions of leaf and hypocotyl, but the enzyme activity was in the membrane fractions (data not shown). In peanut leaf, the activity was ~48-fold lower than that of immature seed. These data suggest that MAG acyltransferase was also present in other plant tissues but at low levels.

Purification of MAG Acyltransferase from Peanut Cotyledons-- MAG acyltransferase activity was found to be high in the soluble fraction, and the rate of synthesis of diacylglycerol in the soluble fraction was 24 pmol min-1 mg-1. Solid ammonium sulfate was added to bring the soluble fraction to 1 M and then loaded onto an octyl-Sepharose column. The column was eluted with a 1-0 M linear-reversed gradient of ammonium sulfate (Fig. 3A). This step was the most effective resulting in a 221-fold purification of acyltransferase and yielding a 2.6-fold increase in the total activity. The active fractions from the octyl-Sepharose were loaded onto a blue-Sepharose column and eluted with a linear NaCl gradient. The activity was eluted between 0.35 and 0.4 M NaCl (Fig. 3B). The recovery of MAG acyltransferase activity from the blue-Sepharose column was nearly 77% of that applied. The pooled active fractions were applied to a preparative Superdex 75 column. The MAG acyltransferase activity was eluted as a single peak from fraction 27 to 31 (Fig. 3C). The active fractions were pooled and applied to a palmitoyl-CoA agarose column as the final step. An overall purification of ~6,608-fold was obtained, and the specific activity of acyltransferase was 15.86 nmol min-1 mg-1 (Table I). The purified enzyme was resolved on a 12% SDS-polyacrylamide gel, which showed a single band with a molecular mass of 43 kDa (Fig. 4). The native molecular mass of the purified enzyme was found to be 43 kDa by Superdex 200 column chromatography (data not shown).



View larger version (24K):
[in this window]
[in a new window]
 
Fig. 3.   Elution profile of MAG acyltransferase with column chromatography. A, the soluble fraction from immature peanuts was loaded onto an octyl-Sepharose column that was pre-equilibrated with 1 M ammonium sulfate. The MAG acyltransferase was eluted from the octyl-Sepharose column in a reversed linear gradient of ammonium sulfate (---), and 5-ml fractions were collected. B, the active fractions from the octyl-Sepharose were pooled and loaded onto a blue-Sepharose column. The MAG acyltransferase was eluted in a linear gradient of NaCl (---), and 1.5-ml fractions were collected. C, active fractions from the blue-Sepharose column were pooled and loaded onto a preparative gel filtration column (Superdex 75), and 5-ml fractions were collected. All the fractions from various column effluents were assayed for MAG acyltransferase activity (open circle ) and protein concentration ().


                              
View this table:
[in this window]
[in a new window]
 
Table I
Purification of MAG acyltransferase from developing peanut cotyledons
The results are from the summary of purification of the MAG acyltransferase. Frozen immature seed (100 g) was used for preparing the soluble fraction. The enzyme activity measurement and the purification steps are described in "Experimental Procedures."



View larger version (95K):
[in this window]
[in a new window]
 
Fig. 4.   SDS-polyacrylamide gel electrophoresis profile of MAG acyltransferase purification. Samples from each purification step were separated by 12% SDS-polyacrylamide gel electrophoresis. Lanes 1-5 correspond to the pooled fractions from steps 1-5 (Table I). Lane Mw represents the standard molecular mass marker.

To confirm that the 43-kDa polypeptide corresponds to the MAG acyltransferase, the purified enzyme was loaded onto a 12% SDS-polyacrylamide gel in the presence of 0.1% SDS without boiling and electrophoresed at 4 °C. The gel was cut into 0.5-cm sections, the protein eluted from the gel with Buffer A and assayed for the enzyme activity (29). The yield of MAG acyltransferase activity from the gel was low (1.5%), and the activity was associated with the area of the gel corresponding to the 43-kDa protein (data not shown). These results indicate that the 43-kDa protein detected on the silver-stained gel (Fig. 4) was indeed the MAG acyltransferase.

The reaction products formed at each step of purification were analyzed on a silica-TLC and autoradiographed (Fig. 5). When the soluble fraction was incubated with [14C]palmitoyl-CoA in the presence of 1-MAG (16:0), formation of 1-acyl-, 1,2-diacyl-, and 1,3-diacyl- and triacylglycerols was observed suggesting the presence of many different acylation activities. The active fractions eluted from the octyl-Sepharose column showed minor amounts of other acylation activities, which diminished in further purification steps as shown in Fig. 5.



View larger version (53K):
[in this window]
[in a new window]
 
Fig. 5.   Autoradiography of the TLC profile of the reaction products formed at each step of purification. The enzyme was assayed using [14C]palmitoyl-CoA and 1-MAG (16:0), and the products formed were chromatographed using petroleum ether/diethyl ether/acetic acid (70:30:1, v/v; A) and chloroform/methanol/acetic acid (98:2:0.5, v/v; B) as the solvent systems. Lane 1 represents heat-inactivated soluble fraction; lanes 2-6 correspond to the active fractions from steps 1-5 (Table I).

Characteristics of Peanut MAG Acyltransferase-- The enzyme activity was linear with respect to time and protein concentrations, and the pH optimum of the MAG acyltransferase was found to be 7.0. The enzyme was specific for MAG and did not utilize any other acyl acceptor such as glycerol, G3P, LPA, or lysophosphatidylcholine. The effect of various detergents on the MAG acyltransferase activity was studied (data not shown). In the presence of 0.3% Triton X-100, the enzyme activity was reduced to 58%. At 20 mM concentration of the zwitterionic detergent (CHAPS), the enzyme lost its activity completely. The activity of MAG acyltransferase was reduced to 50% in the presence of 40 mM octylglucoside (data not shown).

Substrate Dependence of MAG Acyltransferase-- The substrate specificity of MAG acyltransferase was studied by providing monoacylglycerol of varying chain lengths and position of fatty acid as the substrate. The MAG acyltransferase activity was highest for 1-MAG (16:0) and lower for 2-MAG (16:0). The initial rate of reaction was high for 1-MAG (18:1) but the Vmax was small when compared with MAG containing saturated acyl chains. The rate of reaction declined sharply after 10 µM of 1-MAG (18:1). The 1-MAG (16:0) and 1-MAG (18:1) had higher Vmax values and the Km values were 16.39 and 5.65 µM, respectively. The other monoacylglycerols had lower Vmax and apparent Km values (Fig. 6A). These results suggest that the MAG acyltransferase preferentially use sn-1-monoacylglycerols.



View larger version (17K):
[in this window]
[in a new window]
 
Fig. 6.   Lineweaver-Burk plot of MAG acyltransferase activity toward monoacylglycerols and acyl-CoAs. A, the enzyme activity was measured as a function of MAG concentration, and the concentration of palmitoyl-CoA (20 µM) was kept constant. B, the enzyme activity was measured as the function of acyl-CoA concentrations, and 1-palmitoyl-sn-glycerol (20 µM) was kept constant. Each point is the average of two determinations.

MAG acyltransferase activity was the highest for palmitoyl-CoA when compared with stearoyl-, and oleoyl-CoAs, but the initial rate of reaction was higher for oleoyl-CoA. The apparent Km values for palmitoyl-CoA, oleoyl-CoA, and stearoyl-CoA were 17.54, 9.35, and 25.64 µM, respectively (Fig. 6B). Competition studies with myristoyl-CoA and lauroyl-CoA showed that the medium-chain acyl-CoAs and acetyl-CoA were not good substrates for the MAG acyltransferase (data not shown). Based on the Lineweaver-Burk plots, apparent Vmax and Km values for acyl acceptors and acyl donors at the standard assay conditions were obtained as summarized in Table II.


                              
View this table:
[in this window]
[in a new window]
 
Table II
Kinetic parameters of MAG acyltransferase

Effect of Fatty Acids, Phospholipids, and Sphingoid Bases on MAG Acyltransferase Activity-- Effect of fatty acids on the MAG acyltransferase activity was studied using C8 to C18 and C18:1 (Fig. 7A). Oleic acid stimulated the MAG acyltransferase activity at concentrations below 15 µM, but at the higher concentrations, the activity was reduced. Palmitic acid at concentrations between 20 and 50 µM had a stimulatory effect on the enzyme activity, and other fatty acids had no significant effect on the MAG acyltransferase activity. Phosphatidylcholine and phosphatidic acid had stimulatory effects on the purified MAG acyltransferase activity at lower concentrations from 2.5 to 15 µM, whereas at higher concentrations no stimulatory effect was observed (Fig. 7B). Phosphatidylethanolamine and phosphatidylinositol had no effect on the enzyme activity. At lower concentrations of 2.5-7.5 µM, sphingosine and sphingomylein activated the acyltransferase activity (Fig. 7C). The derivative of sphingosine, dehydrosphingosine showed lower stimulatory effects when compared with sphingosine. Dehydrosphingosine had no effect on the MAG acyltransferase activity.



View larger version (19K):
[in this window]
[in a new window]
 
Fig. 7.   Effect of fatty acids, phospholipids, and sphingoid bases on MAG acyltransferase activity. The MAG acyltransferase activity was measured under the standard assay conditions using [14C]palmitoyl-CoA and 1-palmitoyl-sn-glycerol in the presence of various concentrations of fatty acids (A), phospholipids (B), and sphingoid bases (C). Each point is the average of two determinations.



    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The biosynthesis of diacylglycerol is known to occur via the Kennedy pathway in the microsomal membranes (1-3, 30). The present study deals with the first identification, purification, and characterization of a soluble MAG acyltransferase from oilseeds. The soluble DAG biosynthesizing activity was observed in immature peanut cotyledons. The presence of a few soluble enzymes that provide precursors for lipid biosynthesis have been reported. A soluble G3P acyltransferase has been isolated from cocoa seed (31). PA phosphatase is found to be localized both in the soluble and the membrane fractions of Saccharomyces cerevisiae (32, 33) and higher plants (34). A soluble DAG biosynthetic activity has been demonstrated in developing rapeseed (25). LPA phosphatase (9, 10), DAG kinase (35, 36), inactive choline cytidyltransferase (37), and active ethanolaminephosphate cytidyltransferase (38) have also been found in the cytosol of animal systems.

The incubation of NaF-treated soluble fraction with [14C]PA did not generate DAG; however, about 62-75% of [14C]palmitoyl-CoA incorporation into diacylglycerol was observed in the immature peanuts. The synthesized DAG did not originate from the hydrolysis of either triacylglycerol or phosphatidylcholine. These results provide evidences for an alternate enzymatic step for the synthesis of DAG. In this study, we show that DAG is synthesized by the acylation of MAG by acyl-CoA-dependent acyltransferase and this enzyme has not been purified from any source. The following observations revealed that the MAG acyltransferase is present in the soluble fraction. (i) The activity is associated with 150,000 × g supernatant. (ii) The enzyme is permeable in the gel filtration column, and (iii) the MAG acyltransferase is purified to homogeneity by successive column chromatographic separations without detergent. The role of this enzyme in intracellular processes and its regulation has yet to be elucidated. It appears that different tissues express isozymes with respect to subcellular location. For example, leaf enzyme is found in the particulate fraction, but the soluble form is found in immature seeds.

We have purified to apparent homogeneity a MAG acyltransferase activity from developing peanut cotyledons by successive chromatographic procedures. The key to the successful purification was the initial step of octyl-Sepharose column chromatography, and this step gave a 221-fold purification with a 2.6-fold increase in the total activity. The increase in total enzyme activity could be attributed to the elimination of an inhibitor or the enzymes competing for palmitoyl-CoA. The remaining purification steps showed a successive increase in the specific activity and the fold purification.

The purified peanut MAG acyltransferase showed the highest activity with palmitoyl-CoA, but oleoyl-CoA had a lower Km when compared with palmitoyl- or stearoyl-CoAs. Characterization of the partially purified MAG acyltransferase from rat liver (14), intestinal mucosa (39), and adipocytes (40) has also showed higher activity with palmitoyl-CoA. Unlike rat MAG acyltransferase (14), peanut enzyme showed a preference to sn-1-monoacylglycerol over the sn-2 isomer. This observation could also be attributed to the possible acyl migration from sn-2 to sn-1 of MAG during conditions of storage, assay, or extraction (41). However, we are not certain about either of the two possibilities. Kinetic experiments showed that the overall catalytic efficiencies (Vmax/Km) for 1-acyl acceptor was higher than that of the 2-acyl acceptor suggesting the 1-acyl acceptor was a good substrate. The analysis of acyl donors showed that the catalytic efficiency for oleoyl-CoA and palmitoyl-CoA were comparable.

The activity of most of the lipid biosynthetic enzymes is dependent on or modulated by the various lipid cofactors. In developing oilseeds, lipid biosynthesis is highly active, and various metabolic intermediates are accumulated during seed development. All these intermediates either stimulate or inhibit the enzymes involved in lipid metabolism. The characterization of the purified peanut MAG acyltransferase indicated that the lower concentrations of phospholipids and oleic acid stimulated the activity. Apart from oleic acid, palmitic acid also showed an activation effect on peanut MAG acyltransferase. In contrast to our results, it was shown in partially purified hepatic MAG acyltransferase that the higher concentrations of fatty acid inhibited the enzyme activity (15). Sphingosine was shown to inhibit rat hepatic MAG acyltransferase (42), but the peanut enzyme was activated in the presence of lower concentrations of sphingosine, and no inhibition was observed at higher concentrations.

Identification of MAG acyltransferase in peanut indicates the presence of the MAG pathway for DAG biosynthesis. It has been proposed in animal systems that the MAG pathway may play an important role in the regulation of lipid metabolism by controlling the chain length of fatty acids (43) or controlling the intracellular concentrations of acyl-CoA esters (44) or facilitating selective retention of essential fatty acids during hepatic oxidation (13). In plants, the MAG pathway may be involved in the synthesis of triacylglycerol and may also provide a regulatory link between signal transduction and synthesis of complex lipids. Another possibility is that the MAG pathway contributes to a separate intracellular pool of DAG for different sets of metabolic reaction (45). The identification of the MAG acyltransferase has significant implications in understanding the regulation of di- and triacylglycerol biosynthesis in plants.


    ACKNOWLEDGEMENTS

We thank Dr. P. N. Rangarajan for helpful discussions and for allowing us to use the FPLC facility. We are grateful to Dr. Savithramma, University of Agricultural Sciences, Bangalore for giving us peanut and castor seeds.


    FOOTNOTES

* This research was supported by the Dept. of Science and Technology, and the Dept. of Biotechnology, New Delhi, India.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Dagger To whom correspondence should be addressed: Tel.: 91-80-3092881; Fax: 91-80-3602627; E-mail: lipid@biochem.iisc.ernet.in.

Published, JBC Papers in Press, January 18, 2001, DOI 10.1074/jbc.M100005200


    ABBREVIATIONS

The abbreviations used are: G3P, glycerol-3-phosphate; DAG, diacylglycerol; FFA, free fatty acid; LPA, lysophosphatidic acid; MAG, monoacylglycerol; PA, phosphatidic acid; TAG, triacylglycerol; FPLC, fast protein liquid chromatography; CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES


1. Bell, R. M., and Coleman, R. A. (1980) Annu. Rev. Biochem. 49, 459-487[CrossRef][Medline] [Order article via Infotrieve]
2. Browse, J., and Somerville, C. (1991) Annu. Rev. Plant Physiol. Plant Mol. Biol. 42, 467-506[CrossRef]
3. Lehner, R., and Kuksis, A. (1996) Prog. Lipid Res. 35, 169-201[CrossRef][Medline] [Order article via Infotrieve]
4. Webber, K. O., and Hajra, A. K. (1992) Methods Enzymol. 209, 92-98[Medline] [Order article via Infotrieve]
5. Athenstaedt, K., Paltauf, F., Weys, S., and Daum, G. (1999) J. Bacteriol. 181, 1458-1463[Abstract/Free Full Text]
6. van Crovan, E. J., Groenink, A., Jalink, K., Eichholtz, T., and Moolenaar, W. H. (1989) Cell 59, 45-54[Medline] [Order article via Infotrieve]
7. Jalink, K., Eichholtz, T., Postma, S. R., van Crovan, E. J., and Moolenaar, W. H. (1993) Cell Growth Differ. 4, 247-255[Abstract]
8. Jalink, K., Moolenaar, W. H., and Van Duun, B. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 1857-1861[Abstract]
9. Xie, M., and Low, M. G. (1994) Arch. Biochem. Biophys. 312, 254-259[CrossRef][Medline] [Order article via Infotrieve]
10. Hiroyama, M., and Takenawa, T. (1998) Biochem. J. 336, 483-489[Medline] [Order article via Infotrieve]
11. Hiroyama, M., and Takenawa, T. (1999) J. Biol. Chem. 274, 29172-29180[Abstract/Free Full Text]
12. Nishizuka, Y. (1992) Science 258, 607-614[Medline] [Order article via Infotrieve]
13. Pelech, S., and Vance, D. (1989) Trends Biochem. Sci. 14, 28-30[CrossRef]
14. Xia, T., Mostafa, N., Bhat, B. G., Florant, G. L., and Coleman, R. A. (1993) Am. J. Physiol. 265, R414-R4198[Abstract/Free Full Text]
15. Coleman, R. A., and Haynes, E. B. (1984) J. Biol. Chem. 229, 8934-8938
16. Bhat, B. G., Wang, P., and Coleman, R. A. (1994) J. Biol. Chem. 269, 13172-13178[Abstract/Free Full Text]
17. Tsujita, T., Miyazaki, T., Tabei, R., and Okuda, H. (1996) J. Biol. Chem. 271, 2156-2161[Abstract/Free Full Text]
18. Mattson, F. H., and Volpenhein, R. A. (1961) J. Lipid Res. 2, 58-62[Abstract/Free Full Text]
19. Fletcher, M. J. (1968) Clin. Chim. Acta 22, 393-397[Medline] [Order article via Infotrieve]
20. De La Roche, I. A., Andrews, C. J., and Kates, M. (1973) Plant Physiol. 51, 468-472
21. de Duve, C., Pressmann, B. C., Giannetto, R., Wattiaux, R., and Applemans, F. (1955) Biochem. J. 60, 604-617
22. Smith, P. K., Krohn, R. I., Hermanson, G. T., Mallia, A. K., Gartner, F. H., Provenzano, M. D., Fujimoto, E. K., Goeke, N. M., Olsen, B. J., and Klenk, D. C. (1985) Anal. Biochem. 150, 76-85[Medline] [Order article via Infotrieve]
23. Bligh, E. G., and Dyer, W. J. (1959) Can. J. Biochem. Physiol. 37, 911-917
24. Ichihara, K., Norikura, S., and Fujii, S. (1989) Plant Physiol. 90, 423-419
25. Murphy, D. J. (1988) Lipids 23, 157-163[Medline] [Order article via Infotrieve]
26. Sparace, S. A., and Moore, T. S. (1979) Plant Physiol. 63, 963-972
27. Gallagher, S. R., and Leonard, R. T. (1982) Plant Physiol. 70, 1335-1340
28. Briskin, D. P., Leonard, R. T., and Hodges, T. K. (1987) Methods Enzymol. 148, 542-558
29. Bischoff, K. M., Shi, L., and Kennelly, P. J. (1998) Anal. Biochem. 260, 1-17[CrossRef][Medline] [Order article via Infotrieve]
30. Kennedy, E. P. (1961) Fed. Proc. Fed. Am. Soc. Exp. Biol. 20, 934-940
31. Fritz, P. J., Kauffman, J. M., Robertson, C. A., and Wilson, M. R. (1986) J. Biol. Chem. 261, 194-199[Abstract/Free Full Text]
32. Hosaka, K., and Yamashita, S. (1984) Biochim. Biophys. Acta 796, 110-117[Medline] [Order article via Infotrieve]
33. Carman, G. M. (1997) Biochim. Biophys. Acta 1348, 45-55[Medline] [Order article via Infotrieve]
34. Ichihara, K., Murata, N., and Fujii, S. (1990) Biochim. Biophys. Acta 1043, 227-234[Medline] [Order article via Infotrieve]
35. Lapetina, E. G., and Hawthorne, J. N. (1971) Biochem. J. 122, 171-179[Medline] [Order article via Infotrieve]
36. Kanoh, H., Kondoh, H., and Ono, T. (1983) J. Biol. Chem. 258, 1767-1774[Abstract/Free Full Text]
37. Kent, C., and Carman, G. M. (1999) Trends Biochem. Sci. 24, 146-150[CrossRef][Medline] [Order article via Infotrieve]
38. Sundler, R. (1975) J. Biol. Chem. 250, 8585-8590[Abstract]
39. Coleman, R. A., and Haynes, E. B. (1986) J. Biol. Chem. 261, 224-228[Abstract/Free Full Text]
40. Jamdar, S. C., and Cao, W. F. (1992) Arch. Biochem. Biophys. 296, 419-425[Medline] [Order article via Infotrieve]
41. Stimmel, B. F., and King, C. G. (1934) J. Am. Chem. Soc. 56, 1724-1725
42. Bhat, B. G., Wang, P., and Coleman, R. A. (1995) Biochemistry 34, 11237-11244[Medline] [Order article via Infotrieve]
43. Knudsen, J., Clark, S., and Dils, R. (1975) Biochem. Biophys. Res. Commun. 65, 921-926[Medline] [Order article via Infotrieve]
44. Gross, R. W. (1983) Biochemistry 22, 5641-5646[Medline] [Order article via Infotrieve]
45. Gurr, M. I. (1980) in The Biochemistry of Plants (Stumpf, P. K., ed), Vol. 4 , pp. 205-248, Academic Press, New York


Copyright © 2001 by The American Society for Biochemistry and Molecular Biology, Inc.