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Originally published In Press as doi:10.1074/jbc.M311305200 on February 5, 2004

J. Biol. Chem., Vol. 279, Issue 16, 16101-16110, April 16, 2004
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On the Export of Fatty Acids from the Chloroplast*

Abraham J. K. Koo, John B. Ohlrogge, and Mike Pollard{ddagger}

From the Department of Plant Biology, Michigan State University, East Lansing, Michigan 48824-1312

Received for publication, October 14, 2003 , and in revised form, January 22, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The model for export of fatty acids from plastids proposes that the acyl-ACP (acyl carrier protein) product of de novo fatty acid synthesis is hydrolyzed in the stroma by acyl-ACP thioesterases and the free fatty acid (FFA) released is then transferred to the outer envelope of the plastid where it is reactivated to acyl-CoA for utilization in cytosolic glycerolipid synthesis. Experiments were performed to assess whether the delivery of nascent FFA from the stroma for long chain acyl-CoA synthesis (LACS) occurs via simple diffusion or a more complex mechanism. The flux through the in vivo FFA pool was estimated using kinetic labeling experiments with spinach and pea leaves. The maximum half-life for FFA in the export pool was <=1 s. Isolated pea chloroplasts incubated in the light with [14C]acetate gave a linear accumulation of FFA. When CoASH and ATP were present there was also a linear accumulation of acyl-CoA thioesters (plus derived polar lipids), with no measurable lag phase (<30 s), indicating that the FFA pool supplying LACS rapidly reached steady state. The LACS reaction was also measured independently in the dark after in situ generated FFA had accumulated yielding estimates of LACS substrate-velocity relationships. Based on these experiments the LACS reaction with in situ generated FFA as substrate is only about 3% of the LACS activity required in vivo at the very low concentrations of the FFA export pool calculated from the in vivo experiment. Furthermore, bovine serum albumin rapidly removed in situ generated FFA from chloroplasts, but could not compete effectively for "nascent" FFA substrates of LACS. Together the data suggest a locally channeled pool of exported FFA that is closely linked to LACS.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
De novo fatty acid biosynthesis in plants occurs mainly in the plastid (1). The immediate product, acyl-ACP1 (acyl carrier protein) thioester, may be used directly by plastid-localized acyltransferases for the synthesis of "prokaryotic" lipids that are assembled within the chloroplast. Alternatively the acyl moiety of the acyl-ACP can be exported to the cytoplasm where it is primarily incorporated into glycerolipids at the endoplasmic reticulum (eukaryotic pathway) (2, 3). In leaves of plants such as Arabidopsis and spinach (16:3 plants) both pathways contribute about equally to the synthesis of leaf glycerolipids (3), whereas in non-photosynthetic tissue of all plants and in leaves of 18:3 plants such as pea the eukaryotic pathway is the major pathway for glycerolipid synthesis (4).

The current model for export of fatty acids produced by de novo synthesis within the plastid proposes that acyl-ACP hydrolysis occurs in the plastid stroma by the action of acyl-ACP thioesterases, possibly at the inner leaflet of the inner envelope, and that the free fatty acid (FFA) released is transferred to the outer envelope of the plastid where it is reactivated to acyl-CoA for utilization in cytosolic glycerolipid synthesis (Fig. 1). The chemical principle of this model was first proposed by Shine et al. (5). As subsequent experiments showed that ACP-dependent fatty acid synthesis was almost entirely a chloroplast function (1) this implied that the soluble acyl-ACP thioesterase was a stromal enzyme activity. Long-chain acyl-CoA synthesis (LACS) activity is associated with the chloroplast envelope (6, 7) and more specifically with the outer envelope (8, 9). In addition, the major product of fatty acid synthesis in isolated chloroplasts is FFA, unless ATP and CoASH are both present, when acyl-CoA becomes a major end product (10, 11). In vivo confirmation that FFA is indeed an intermediate in fatty acid export from the plastid was accomplished by the use of 18O labeling experiments (12).



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  		FIG. 1.
Schematic for fatty acid export from the site of de novo fatty acid synthesis in the chloroplast stroma to the cytosol. Two simplified models are shown. The lower route is for simple diffusion of FFA and upper route is via a protein mediated export mechanism. RCO, long chain fatty acyl group.

 
There are several unknowns when considering the details of a fatty acid export process. First, it is unclear if the acyl-ACP thioesterases associate with the inner leaflet of the inner membrane of the envelope, thus releasing the acyl group to the inner leaflet (as shown in Fig. 1). Second, the vectorial nature of the long chain acyl-CoA synthesis (LACS) reaction is unknown. The outer envelope contains pores which can accommodate molecules of up to 8-10 kDa (13). Thus metabolites such as ATP, CoASH, and acyl-CoAs may diffuse into the intermembrane space between the inner and outer envelope. It is uncertain whether soluble acyl-CoA-binding proteins, which have a mass of 10-11 kDa (14), will be able to move as freely. We do not know whether the plastid long chain acyl-CoA synthetase can bind FFA at the inner leaflet of the outer envelope, then transfer FFA to the outer leaflet for activation to acyl-CoA, or whether the transfer of FFA across the outer envelope is accomplished by other means, or if the active site of the long chain acyl-CoA synthetase is at the inner leaflet of the outer envelope. We note by way of analogy that pairs of proteins, one of which is always an acyl-CoA synthetase, have been implicated in the vectorial import of fatty acids in Escherichia coli (15), in yeast (16) and in animal cells (17, 18). Third, there appears to be no direct connections between the inner and the outer envelope of the plastid, although there may be some "contact points" (19). Thus at the minimum we would expect that FFA will have to flip-flop across the inner envelope membrane, and then possibly disassociate into the periplasmic space to reach the long chain acyl-CoA synthetase at the outer envelope (Fig. 1, lower route). Given these unknowns a major question is whether the movement of FFA out of chloroplasts is a transporter-facilitated process or involves free diffusion of the FFA intermediates. This is a debate that has parallels in other systems where FFA transport is required (20-23). We report a variety of experiments designed to examine the question of FFA export from the plastid, using assays with intact leaves and chloroplasts.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials-Pea seeds (Pisum sativum L. cv Little marvel) were germinated on moist vermiculite. For chloroplast preparations the peas were grown for 8-10 days in the growth chamber at 25 °C and an 8 h photoperiod with occasional watering. For the in vivo experiments the peas were grown for 12 days in the growth chamber watering with nutrient solution. Young pea leaves were removed that had yet to open. These were cut in half laterally, immediately placed in assay medium and preincubated as described below. [1-14C]Acetic acid (57.2 Ci/mol) was purchased from American Radiolabeled Chemicals Inc. (St. Louis, MO).

In Vivo Incubations-Leaf assays with halved pea leaves were run in 5.0 ml of medium containing 25 mM NaMES buffer, pH 5.7, with 0.3-0.4 g of fresh weight of tissue, at 28 °C and under strong illumination (180 µmol of photons/m2/s). Tween 20 was added as a wetting agent to a concentration of 0.01% w/v. The reaction was initiated with sodium [1-14C]acetate solution. Time course assays were run with 0.03-0.06 mCi of labeled acetate (0.105-0.21 mM). These assays were terminated by partial removal of the medium and quenching with hot isopropyl alcohol while the tissue was under illumination.

Chloroplast Preparation and Assays-Intact chloroplasts were isolated on a continuous Percoll gradient (24). All procedures were conducted at 0-4 °C. About 10-20 g of 8-day-old pea seedlings were homogenized in 100 ml of semi-frozen homogenization buffer (50 mM HEPES, pH 8.0, 330 mM sorbitol, 0.1% BSA, 1 mM MgCl2, 1 mM MnCl2, 2 mM EDTA) using a polytron (PT 10/35) and filtered through 2 layers of miracloth (Calbiochem). Crude chloroplasts were collected by centrifugation at 1200 x g max for 5 min. These were purified by centrifugation through pre-made gradients of Percoll generated by centrifugation of 30 ml of 50% Percoll in homogenization buffer for 30 min at 40,000 x g. Crude chloroplasts (2-3 ml) were centrifuged through the pre-made gradients at 13,000 x g max for 5 min in an HB-4 rotor without brake. The intact chloroplast band formed near the bottom of the gradient was recovered and washed with resuspension buffer containing 50 mM HEPES, pH 8.0 and 330 mM sorbitol. The resulting isolated chloroplasts can incorporate acetate into fatty acids at rates corresponding to in vivo rates of fatty acid synthesis, import and process in vitro translated protein precursors, sustain high rates of photosynthesis (25), and give about 90% intactness as determined by phase contrast light microscopy and the reduction of ferricyanide before and after an osmotic shock (26, 27). Quantification of chlorophyll was performed as described by Arnon (28).

Chloroplast incubations to synthesize fatty acids were performed largely according to the methods described by Roughan (29). Chloroplasts (40-50 µg of chlorophyll) were incubated under illumination (180 µmol of photons/m2/s) with shaking at 25 °C in a medium (200 µl) containing 0.33 M sorbitol, 25 mM HEPES-NaOH, pH 8.0, 10 mM KHCO3, 2 mM Na3EDTA, 1 mM MgCl2, 1 mM MnCl2, 0.5 mM K2HPO4, and 2 mM acetate containing 2 µCi of [14C]acetate. Where noted in the text assays also contained 1 mM ATP, 0.5 mM CoA, and/or 0.5 mg/ml BSA added at specific times. An aliquot was removed before the illumination and used to measure the actual radioactivity added. Reactions were terminated by placing the incubation mixture into liquid nitrogen and were kept in -80 °C freezer until lipid analysis.

In the BSA/FFA binding assays, chloroplasts were separated from the BSA in the medium by centrifugation at 13,000 rpm (Sorvall Biofuge pico) for 15 s. The supernatant was aspirated from the chloroplast pellet and frozen in liquid nitrogen. Fatty acids were extracted from the supernatant and the chloroplast pellet separately.

Lipid Analysis-After quenching the tissue by heating in isopropyl alcohol at 80-90 °C for 5 min lipid extraction from the [14C]acetate fed leaf strips was carried out using hexane-isopropyl alcohol (30). An aliquot of the lipid extract was suspended in 4:1 acetone/water and the OD at 652 nm measured to determine chlorophyll (28); another aliquot was assayed by liquid scintillation counting to determine radioactivity incorporated. Suspension of the lipid residue in organic solvent and re-evaporation in the presence of a few drops of glacial acetic acid was crucial to drive off residual acetate substrate contamination to give accurate radioactivity determinations. Lipid classes and fatty acid methyl esters were analyzed by thin layer chromatography (TLC) and isolated by preparative TLC. TLC plates were scanned for radioactivity using a Packard Instant Imager (Canberra Instruments), both to quantify radioactivity and to locate the appropriate bands for recovery from preparative TLC plates. For pea leaf extracts a non-saponifable band (4-6% of total label) co-migrated very close to FFA on TLC, so the samples were first treated with ethereal diazomethane and then fatty acid methyl esters determined to measure the free fatty acid pool. Neutral lipids were analyzed on silica plates developed half way with hexane/diethyl ether/acetic acid 70:30:1 (v/v/v) and then fully with hexane/diethyl ether/acetic acid 180:20:1 (v/v/v). For analysis of individual labeled fatty acids the methyl ester fraction was isolated by preparative TLC and separated using either C18-reversed phase TLC, developing half way and then fully with acetonitrile/methanol/water 130:70:1 (v/v/v), or by argentation-TLC using plates impregnated with a 5% silver nitrate in acetonitrile solution, dried and activated, and developed with hexane-diethyl ether mixtures.

The method for analysis of radiolabeled products from chloroplast assays was adapted from Roughan (29). Samples were saponified by heating in 80-90 °C for 60 min in 0.8 ml of 10% KOH (w/v) in methanol per 0.2 ml reaction mixture. After cooling, acidification and extraction with hexane and then evaporation of the hexane under nitrogen the total radioactivity recovered was determined by liquid scintillation counting. Since there are negligible non-saponifiable products this radioactivity is a measure of the incorporation of [14C]acetate into total fatty acids. Lipid class analysis was performed by mixing 0.2 ml of reaction mixture with 2 ml of chloroform/methanol 1:1 (v/v) and 0.72 ml of 0.2 M H3PO4 in 2 M KCl. The lower organic phase was collected, and the upper aqueous methanol phase was back-extracted with hexane (4 ml). The combined organic extracts were evaporated to dryness and an aliquot assayed for radioactivity by liquid scintillation counting. Neutral and polar lipids were separated by TLC using K6 silica plates (Whatman, Clifton, PA) developed in hexane/diethyl ether/glacial acetic acid 80/20/1 (v/v/v). To determine the polar lipid composition, the polar lipid band was recovered and analyzed on K6 silica TLC plates impregnated with 0.15 M ammonium sulfate and activated at 120 °C for 3 h, and developed in acetone/toluene/water 91/30/8 (v/v/v) (31). The aqueous methanol phase contains acyl-CoA as the major saponifiable lipid. Over 80% of the label was shown to co-migrate with long chain acyl-CoA standard when extracted by butan-1-ol and analyzed by TLC (K6 plate developed in butan-1-ol/acetic acid/water 50:20:25 (v/v/v)). Routinely the aqueous phase was saponified, acidified, extracted with hexane and the extractable lipids (i.e. free fatty acids) assayed for radioactivity (11, 32). Control extractions with chloroplasts spiked with known amounts of radiolabeled oleic acid or oleoyl-CoA yielded essentially complete recovery of oleic acid and >85% recovery of oleoyl-CoA either in the presence or absence of BSA.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
The Total in Vivo Pool of FFA in Leaves Is Very Small and Implies a Very Short Half-life for the FFA Pool Involved in Export from the Chloroplast-Although it is generally known that labeled FFA are minor products when leaf tissue is incubated with acetate, the amount of FFA has not previously been quantified. We have shown that FFA are intermediates in fatty acyl group export from the chloroplast (12) and to better understand this export process it was important to quantify this FFA pool. Fig. 2 presents a time course for acetate incorporation into total fatty acids and into FFA by pea leaves over a 20 min period.



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  		FIG. 2.
Time course for acetate incorporation into acyl lipids by pea leaves. Pea leaf tissue was incubated with non-saturating concentrations of [14C]acetate (0.105 mM). The data points are the average of triplicate determinations, and error bars represent the range of values. A, shows acetate incorporation into total lipids by halved pea leaves over time. At each time point 80-85% of the label in total lipids was present as labeled fatty acid. B, shows acetate incorporation into the labeled FFA in the same experiment, and is measured as pmol of acetate incorporated/mg of chlorophyll.

 
With expanding pea leaves a linear rate of lipid synthesis was established within 2 min (Fig. 2A). At all time points the labeled total lipid extract from pea leaves contains 80-85% labeled fatty acids, mainly as phosphatidylcholine. Presumably the short lag phase indicates the time to reach the steady state balance between transport processes and pool filling in the biosynthetic utilization of acetate. To measure the FFA pool the leaf assays were quenched while still in the presence of substrate in the light (Fig. 2B). Control extractions spiking unlabeled tissue with [14C]oleic acid gave >90% recovery of label. In the experiment in Fig. 2 the labeled FFA pool at 2 min was 4.3% of total labeled lipids and contained essentially only palmitic and oleic acids. In pea most of the fatty acids synthesized in the chloroplast are exported (~90%). The turnover time of FFA during plastid export for pea can be estimated as follows. Let the steady state rate of lipid synthesis be 100 units per min, of which ~85 units are fatty acids. Therefore 76.5 units (90% of 85%) must pass through the FFA transfer pool for export to the eukaryotic pathway. At the 2 min time point the steady state level of labeled FFA in the transfer pool has been reached, and because the steady state rate extrapolates back to the x-axis at 1 min (Fig. 2A) the total amount of lipids synthesized is equivalent to about 100 units. Thus 100 units x 4.3% of labeled FFA are present (4.3 units). At 2 min we have a flux of 76.5 units per min through a transfer pool of 4.3 units. This gives a turnover time of 60 x 4.3/76.5 = 3.4 s and a t1/2 value of 2.35 s (t1/2= turnover time x 0.693). A repeat of the time course with pea leaves gave the onset of steady state labeling within 1 min, while the labeled FFA pool at 2 min was 2.05 ± 0.26% (4 determinations), giving a t1/2 value of 1.7 s. A similar time course and FFA pool analysis experiment was performed with spinach leaf strips (data not shown). A turnover time 0.96 s and a t1/2 value of 0.66 s were determined.

These calculations for pea and spinach leaves represent estimated values for the FFA transfer pool turnover times that are an upper limit. The observed FFA pool may include the transient pool of FFA being exported from the plastid, FFA released from labeled acyl thioester pools during the heat quench step, plus FFA derived from other sources. The kinetics of labeled FFA appearance in the time course for pea leaves (Fig. 2B) give us an indication that much of the labeled FFA pool is not a transfer pool. If the observed FFA pool were all the transfer pool then this pool would saturate by 2 min. Instead the pool continues to grow substantially. We speculate that some of the FFA pool actually derives from fatty acid synthesis at the damaged margins of the leaf strips in the assay. Given the approximations in the calculations it seems reasonable to state that the t1/2 for FFA in any chloroplast export pool is about 1 s, or less, and possibly much less. Having established this limiting in vivo pool size we then examined the detailed relationship between products of fatty acid synthesis in isolated, intact pea chloroplasts. In pea, compared with spinach, much more of the fatty acid produced is exported from the chloroplast.

Time Course for Acetate Incorporation into Fatty Acyl Products by Isolated Pea Chloroplasts-The synthesis of fatty acids from acetate by intact, illuminated chloroplasts can produce a variety of acyl products (34, 35), depending on the plant species from which the chloroplasts are isolated and on the cofactors in the chloroplast incubation medium. In a minimal chloroplast incubation medium (no CoASH, glycerol-3-phosphate, nor UDP-galactose) the major product is free fatty acid (FFA). Smaller amounts of 1,2-diacylglycerol (DAG) (~5-20%) and polar lipids (PL) (~10-20%) have also been observed. When CoASH and ATP are added, up to 60% of the label may be found as acyl-CoA (11).

The incorporation of acetate into acyl lipids by pea chloroplasts is shown in Fig. 3. In minimal chloroplast incubation medium the [14C]acetate incorporation into total fatty acids was 15.5 pmol of fatty acid/s/mg chlorophyll (Fig. 3A), but when CoASH and ATP were present this rate increased 1.4-fold to 21.5 pmol of fatty acid/s/mg chlorophyll (Fig. 3B). In the minimal medium the major product was FFA (about 90% of total label). When CoASH and ATP were present up to 80% of the label was found as acyl-CoA plus polar lipids (PL). Fig. 3B shows that over a 35 min period in the light with ATP and CoASH total fatty acid synthesis was linear, and that the appearance of the individual products, namely FFA (~25%), acyl-CoA (~40%), and total PL (~40%) were also linear, with negligible lag phase detected. The labeled PL fraction was composed of about 60% phosphatidylcholine and 10% phosphatidylglycerol. To confirm the lack of a lag phase earlier time points were taken (Fig. 3C). Any lag phase required for the establishment of the steady state condition of product accumulation in the presence of CoASH plus ATP was less than 30 s. In Fig. 3, B or C if the bulk FFA pool were a precursor of acyl-CoA, which is subsequently used for PL synthesis, a measurable lag would occur in acyl-CoA plus PL accumulation. Thus there was no observable precursor-product kinetic relationship between labeled FFA and acyl-CoA. This implies that the FFA concentration to run the LACS reaction at steady state rate has been reached rapidly (<30 s) and that the bulk-labeled FFA pool accumulating in Fig. 3, B or C is not a precursor for the LACS reaction. The linear rates in Fig. 3, B or C further imply that neither the bulk FFA nor acyl-CoA are acting as feedback inhibitors for fatty acid synthesis by isolated chloroplasts. Our lag phase estimates are currently limited to >10 s by the gentle mixing required for delicate organelles like chloroplasts, for pool filling from acetate through to end products of fatty acid synthesis, and quench times.



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  		FIG. 3.
Time course for acetate incorporation into fatty acyl products by pea chloroplasts. In A, pea chloroplasts (40 µg of chlorophyll) were incubated under illumination in 0.2 ml of basal medium with 2 mM [14C]acetate. For B and C, 1 mM ATP and 0.5 mM CoASH were also added to the assay medium. The fractions assayed were total lipids ({blacksquare}), FFA ({square}), acyl-CoA ({circ}), and total polar lipids (•).

 
The Long Chain Acyl-CoA Synthesis (LACS) Reaction in the Dark Utilizing in Situ Generated FFA-When FFA is exported from the plastid it is converted to acyl-CoA for utilization in cytosolic reactions of lipid biosynthesis. The LACS reaction has previously been studied after preparation of chloroplast envelope vesicles and addition of FFA substrate (7). In this study we investigated the LACS reaction in intact pea chloroplasts with in situ generated FFA. Fig. 4 presents the effect of switching off the light after 15 min of incubation of chloroplasts in media containing no CoASH during the light period but added at the onset of the dark period, or in media containing CoASH during both light and dark periods. As observed by many researchers (33, 36) total fatty acid synthesis was completely halted in the dark. What is noteworthy is the very different behavior of the individual acyl pools during the dark period. In the absence of CoASH the major fatty acid product in the light was FFA (Fig. 4A). When CoASH (0.5 mM) was added at the transition to darkness there was a very rapid conversion of FFA to acyl-CoA and its metabolites (PL). The initial rate of FFA depletion (from 15 to 20 min) was at least twice the rate of total fatty acid synthesis in the light. By contrast, in Fig. 4B the initial rate of FFA depletion on transition to darkness was less than 30% that of the rate of fatty acid synthesis in the light. The combined acyl-CoA and PL accumulation had essentially stopped in the dark period.



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  		FIG. 4.
Depletion of FFA pool after transition from light to dark. Pea chloroplasts (40 µg of chlorophyll) were incubated with 2 mM [14C]acetate under illumination in 0.2 ml of basal medium containing 1 mM ATP for 15 min. After 15 min the lights were removed, and the assay continued. A and C, CoASH (0.5 mM) was added at the moment of transition into dark. B, CoASH (0.5 mM) was added from the beginning of incubation in the light. The data points are average of three independent determinations. In A and B, the fractions assayed were total lipids ({blacksquare}), FFA ({square}), acyl-CoA ({circ}), and total polar lipids (•). C, shows only the reaction after the CoASH addition on transfer to the dark, and the fast and slow rates of FFA depletion are indicated. In C, the fractions assayed were FFA ({square}), acyl-CoA ({circ}), and acyl-CoA plus polar lipids ({blacktriangleup}).

 
Because the LACS reaction with in situ generated FFA was very fast in Fig. 4A this experiment was repeated with additional measurements immediately after the light to dark transition (Fig. 4C). There were variations between five independent experiments but upon the dark transition with coincident addition of CoASH, about 25-50%, and sometimes >50% of the FFA label was removed by the LACS reaction in the first 30 s. Thus the LACS reaction in chloroplasts has the capacity to run at a rate over 10 times that of fatty acid synthesis. However, the LACS reaction did not rapidly go to completion. Inspection of Fig. 4C shows that disappearance of FFA does not follow simple first order kinetics. There are at least two components to the disappearance of FFA; a rapid rate of depletion accounting for at least 60-70% of the labeled FFA but sometimes up to 80-85% depending on experiment, and a slow decay for the remainder.

Evidence for Distinct Kinetic Pools of FFA in Chloroplast Assays-Based on Fig. 4A we know the LACS reaction is highly active in the dark and therefore the slow rate of LACS in Fig. 4B appears somewhat paradoxical, especially since in Fig. 4A the amount of labeled FFA at the dark transition is 4.7 nmol/mg chlorophyll while in Fig. 4B it is still 3.5 nmol/mg chlorophyll. This paradox is also seen in Fig. 3B with the linear accumulation of FFA up to 12 nmol/mg chlorophyll in the presence of ATP and CoASH, and is best explained by multiple FFA pools. The results shown in Figs. 3 and 4 reveal three kinetically distinct pools of FFA products. In all of these pools FFA is assumed to be associated with the chloroplasts and partitioned largely or completely into membranes. These pools are: A, the very small FFA pool supplying the LACS reaction in the light in the presence of CoASH and ATP (Fig. 3, B and C); B, a bulk FFA pool that builds up if the LACS reaction cannot run in the light because of the absence of CoASH but which is accessible to LACS when CoASH and ATP are present (Fig. 3A); and C, a bulk FFA pool (about 20-30% of total label) that is not readily accessible to LACS reaction either in the light or the dark in the presence of ATP and CoASH. If the appearance of acyl-CoA in assays run in the presence of ATP and CoASH in the light is indeed catalyzed by the same enzyme that catalyzes the LACS reaction in the dark, then bulk pool (B) will provide substrate for pool (A). Thus in the light in the presence of ATP and CoASH, newly synthesized FFA rapidly moves through pool (A), or accumulates in a linear fashion in pool (C) (Fig. 3, B and C). Stopping fatty acid synthesis by placing the assay in the dark does not cause much additional synthesis of acyl-CoA and PL because pool (B) is absent (Fig. 4B). In the absence of CoASH during fatty acid synthesis in the light, both pool (B) and pool (C) accumulate in a linear fashion, and when the assay is placed in the dark with CoASH, pool (B) is available for a rapid LACS reaction (Fig. 4, A and C). The physical nature of these distinct FFA pools becomes an important question. Presumably in vivo only pool (A) is present. Pool (B) is artifactual in that in vivo the LACS reaction is not cofactor limited, but as we will discuss later serves as a useful model if FFA is allowed to diffuse freely within the organelle. The LACS reaction of pool (B) FFA would be a mass action driven activation of FFA. Pool (C) is an artifact, perhaps as a consequence of a population of damaged chloroplasts.

The Dependence of the LACS Reaction in the Dark on the Concentration of in Situ Generated FFA-The rapid depletion of FFA by the LACS reaction seen in Fig. 4C will require a series of FFA transport steps (membrane flip-flops and association and disassociation steps) to deliver the FFA to the site of the LACS reaction. We do not know the distribution of FFA within the chloroplast, or the orientation of the LACS active site in the outer envelope, so we cannot speculate on the complexity of these steps. However, if we approximate the FFA fast depletion reaction to a single (averaged) time constant for exponential decay of pool (B) then that time constant is ~30 s, or sometimes less. The concentration dependence of the FFA depletion is shown in Fig. 5, using a 30 s dark LACS reaction assay. At the levels of FFA we allowed to accumulate in the light preincubation period, the rate of the dark LACS reaction was essentially proportional to FFA concentration and thus not saturated with respect to FFA substrate.



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  		FIG. 5.
FFA concentration dependence of the LACS reaction. Isolated pea chloroplasts were incubated with [14C]acetate under illumination in 0.2 ml of basal medium containing 1 mM ATP for varying times in the absence of CoASH and then lights were removed. One set of assays was quenched at this point to determine the FFA pool accumulated. To a second set of assays CoASH (0.5 mM) was added at the moment of transition into dark. After another 30 s in the dark the reaction was stopped by freezing immediately with liquid nitrogen and the FFA assayed to assess its depletion by the LACS reaction.

 
Accessibility of Chloroplast FFA Products to Sequestration by BSA in the Dark-BSA is well known for its ability to bind FFA (37, 38). We have used this FFA-binding capacity in competition experiments with chloroplasts. Chloroplasts that were preloaded with in situ generated [14C]FFA were incubated with different concentrations of defatted BSA for 5 min in the dark and centrifuged to separate the chloroplast pellet from the media. Radiolabeled lipids in the media and in the chloroplast pellet were assayed. At the beginning of the assay 80-85% of the total labeled product was FFA and >85% of the total labeled lipid was associated with the chloroplast. However when the chloroplasts containing labeled FFA were mixed with increasing concentrations of BSA, increasing amounts of FFA were found in the supernatant. This redistribution saturated at about 80% of the label released into the medium at a BSA concentration of 0.3 mg/ml (4.5 µM). The concentration of labeled FFA produced in situ in the assay was 2.5 µM. Next the rate of FFA transfer from the chloroplasts to the BSA was examined by varying the co-incubation time at 0.5 mg/ml BSA (Fig. 6). Over 90% of the labeled FFA partitioned into the media bound to the BSA within 45 s. This was the quickest practical time to conduct transfers and centrifugation. Thus the t1/2 for FFA sequestration by BSA is about 10 s or less. Presumably the FFA:BSA binding reaction is mediated via BSA binding to the outer leaflet of the outer envelope and requires FFA to be present in the outer leaflet and to move through to the outer leaflet rapidly from other sites in the chloroplast.



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  		FIG. 6.
Sequestration of FFA by binding to BSA. Isolated pea chloroplasts were first loaded with in situ generated [14C]FFA by incubation in minimal media with [14C]acetate for 15 min in light. Upon transition to the dark BSA (0.5 mg/ml) was added to the assay medium, which was then centrifuged (16,000 x g) for 15 s at various time points. The supernatant was aspirated and collected, and the transfer reaction was terminated by quenching both the supernatant and pellet immediately with liquid nitrogen. The fractions assayed were FFA ({blacksquare}), acyl-CoA ({circ}), and polar lipids ({square}).

 
BSA Competes with LACS for the Bulk in Situ FFA Pool but Is Not as Effective for the Nascent FFA Pool-Because BSA can rapidly and almost completely remove FFA generated in situ by chloroplasts (Fig. 6), experiments were conducted to measure the rate of the LACS reaction with FFA substrate bound to BSA. FFA was first generated in situ in the presence or absence of BSA for 15 min, then CoASH was added in the dark (Fig. 7). Comparing the initial rates for the LACS reaction the control reaction utilizing FFA bound to the chloroplast was at about 20 times faster than the rate for FFA substrate sequestered in the medium bound to BSA. This difference may be greater still if the results are normalized to the same assay concentration of FFA generated in situ. The data from Fig. 7 indicate that the release of FFA to the chloroplast by BSA is a relatively slow step.



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  		FIG. 7.
Competition between LACS and BSA for in situ generated FFA. Isolated pea chloroplasts were first loaded with in situ generated [14C] labeled fatty acids by incubation in minimal medium containing [14C]acetate as substrate without CoASH but in the presence ({blacksquare}) or absence ({square}) of 0.5 mg/ml BSA for 15 min in light. Upon transition to dark, CoASH (0.5 mM) was added. Reactions were terminated after further incubations with CoASH for various times. Label in FFA was measured. The initial rate in the presence of BSA was 4050 dpm/min and in the absence of BSA (dotted line) 77000 dpm/min.

 
The results of a continuous labeling assay under light in the presence and absence of BSA are shown in Fig. 8. When ATP, CoASH, and BSA are present from the beginning of the assay, LACS and BSA are expected to compete for de novo synthesized "nascent" FFA from the export pool (A). If the FFA diffuses out to the outer envelope, BSA may have a chance to bind a fraction of the FFA and thereby increase the proportion of FFA in the total product. As observed in a previous study (32) BSA stimulated total fatty acid synthesis activity 1.6-fold (average of four different experiments, data not shown). With the addition of BSA the acyl-CoA product increased from 26 to 37% and that of PL decreased from 52 to 41% while FFA level remained unchanged at 22%. The fact that BSA was not able to increase the proportion of FFA in the products again argues against the idea that there was a bulk freely diffusing pool of FFA during the process of FFA export from the chloroplast and suggests that LACS has "prior" access to exported FFA before it diffuses out to the outer leaflet of the outer envelope membrane.



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  		FIG. 8.
Competition between LACS and BSA for nascent FFA in fatty acid synthesis assays. Isolated pea chloroplasts were incubated with [14C]acetate under illumination in a basal incubation medium containing 1 mM ATP and 0.5 mM CoASH, in the presence or absence of 0.5 mg/ml BSA. Samples were collected at 5-, 10-, and 20-min time points, and the percentage of total labeled lipids present in FFA, acyl-CoA and polar lipid fractions was analyzed. Values represent the means of 15 measurements from four independent experiments, with standard deviations shown. The Student's t test comparing control and BSA-added samples showed that differences are significant for acyl-CoA: *, p < 0.001 and PL: **, p < 0.01 but are insignificant for FFA: ***, p > 0.5.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The Endogenous Free Fatty Acid Pool-The principal products of de novo fatty acid synthesis exported from most chloroplasts are palmitate and oleate (4). In our previous study of leaf fatty acid synthesis from [18O]acetate (12), the fatty acyl export from the plastid was demonstrated to involve a hydrolytic step to produce a free carboxylate anion prior to the fatty acid reactivation for incorporation into eukaryotic lipids. This was an expected result but had not been previously demonstrated in vivo. The confirmation makes consideration of FFA transport mechanisms in plastids a relevant physiological problem. An immediate question to address is the size of this FFA transport pool. In all carefully conducted labeling studies of plant tissues only a very small radiolabeled FFA fraction is observed in experiments with [14C]acetate. From our kinetic analysis in this study we estimate that the t1/2 for FFA in any plastid export pool is <= 1 s.

At this point this upper limit placed on the t1/2 value for FFA is not sufficient to rule out any mechanistic possibility. At one extreme it is instructive to consider the stoichiometry for any channeled, protein-mediated transport process. Heber and Heldt (39) have estimated that the inner envelope protein concentration is 200-300 µg/mg chlorophyll. The endogenous rate of fatty acid export is about 15-55 pmol/s/mg chlorophyll (see below). Assuming a transport protein of 60 kDa, with a 1:1 protein:FFA binding stoichiometry and a 50% occupancy of binding sites, then if the average transfer time is about 1 s, the transfer protein concentration is 2-6 µg/mg chlorophyll, or about 0.5-3% of the inner envelope protein. Thus the time constant is consistent with the possibility of a channeled transport system. Should there be a FFA transporter protein (passive or active) in the inner envelope, it is probable given the 2-10 nm periplasmic space between the two envelopes that it may be in direct contact with the acyl-ACP thioesterase at the stromal interface of the inner envelope and with the LACS protein on the outer envelope (Fig. 1, upper route).

At the other extreme, consider the process of free diffusion from the plastid. The possible steps are shown in Fig. 1, lower route. A debate between a facilitated process and free diffusion for FFA uptake by various types of animal cells has persisted for many years (20, 21, 23, 40), and some points are instructive to the issues of FFA export from the plastid. Very fast rates of FFA flip-flop (t1/2 <=25 ms) are observed for small (0.025-µm diameter) and large (0.1-µm diameter) unilamellar PC vesicles, (21, 41, 42). However, Kleinfeld et al. (22) pointed out that there is an inverse relationship between t1/2 for FFA flip-flop and the radius of curvature of the liposomes studied. In giant unilamellar vesicles (>0.2-µm diameter) t1/2 values of 0.7-7 s were measured for FFA flip-flop, values that were later found to be similar for erythrocyte ghosts (40). Chloroplasts typically have diameters of 1-4 µm. An important point is that there is no direct physicochemical measurements for FFA flip-flop or binding and desorption from liposomes composed primarily of galactolipids, such as to mirror the composition of the chloroplast inner envelope or the inner leaflet of the outer envelope. In the absence of such measurements it is difficult to speculate whether the physical rates of FFA flip-flop and membrane desorption might be adequate to account for the rate of FFA export from the plastid, or for any movement of FFA into and from thylakoids. Even with a series of physical measurements the chloroplast is a complex organelle to model when compared with liposomes and erythrocyte ghosts, in that the envelope is a double membrane. In addition within each chloroplast there is the massive endomembrane system of thylakoids that accounts for >90% of the organelle's membrane and which might sequester FFA. Furthermore, in the light the pH of the stroma is about 8.5 whereas the pH of the cytosol is about 7.5. Thus freely diffusing FFA will preferably partition to stromal facing leaflets in chloroplast membranes, that is, to those of the inner envelope and thylakoids.

FFA Pools in Incubations with Intact Chloroplasts-In assays with spinach chloroplasts Roughan et al. (43) noted that when chloroplasts were incubated in the light for 20 min in basal medium and then transferred to the dark for 20 min with CoASH and ATP, the FFA labeled product was reduced by half with the concomitant appearance of acyl-CoA as the major product (or PC when microsomes were also added). These assays were not run as kinetic assays, since their purpose was product identification. The implication in this and other articles and reviews is that the acyl-CoA formed upon addition of ATP plus CoASH is derived from the "bulk" FFA pool (32). Because our kinetic analysis of in vivo leaf assays indicated that the t1/2 for FFA in any plastid export pool is <= 1 s this very small pool is inconsistent with the large FFA pools seen in assays with isolated chloroplasts, and particularly with their being functional transport intermediates. We have investigated this apparent contradiction and showed there is not in fact an inconsistency if the total FFA produced in chloroplast assays is described as the sum of several distinct FFA pools.

The results from our kinetic assays with intact chloroplasts (Figs. 3, 4, 5, 6, 7, 8) are interpreted in terms of three fates (or pools) for FFA. A very small FFA pool (A) that provides "nascent" FFA for LACS is implicated by the lack of precursor-product kinetics between total FFA and acyl-CoA, as shown in Fig. 3B. This pool size (i.e. t1/2 value) is too small to measure accurately but we postulate it is analogous to the in vivo situation. The LACS reaction is not feedback inhibited by continued accumulation of acyl-CoA. The bulk accumulating FFA (Fig. 3A) can be best described as a combination of two pools, neither of which represents a physiological situation. One pool (B) is a LACS-accessible pool, which fills when the LACS reaction cannot function because of lack of cofactors. The other pool (C), is present in all assays, represents 20-40% of the total labeled fatty acids and is not rapidly accessible to LACS. Because Pool C is likely a function of damaged chloroplasts, it is not discussed further.

At saturating concentrations BSA rapidly removes FFA from the chloroplast (t1/2 < 10 s) (Fig. 6). This allowed us to test it as a competitor against the putatively channeled LACS reaction, since the time constant for FFA removal from the chloroplast by BSA was equal to or less than that for the independent LACS reaction. In addition, once FFA is bound to BSA, the LACS reaction is reduced at least 20-fold (Fig. 7). When fatty acid synthesis occurs simultaneously in the presence of ATP, CoASH and BSA, BSA does not inhibit the appearance of acyl-CoA products by sequestration of the exported FFA (Fig. 8). A simple explanation can be advanced for this result. Since we believe the BSA binding of FFA is a physical transfer process it is likely that the rate of removal of FFA from chloroplasts by BSA is proportional to the bulk FFA concentration, which initially is about 2.5 µM (Fig. 6). In intact chloroplasts BSA cannot penetrate the outer envelope, and therefore must load FFA from the outer leaflet of the outer envelope or from the very low amounts of FFA partitioned into the medium. We can estimate pool (A), the pool of FFA required to sustain the channeled LACS reaction, as filling within 30 s or 1 s (using the in vitro or in vivo estimates), which gives the bulk concentration of FFA product in the chloroplast assay as <35 nM or <1 nM. With either calculation, on simple kinetic grounds BSA should not effectively "intercept" FFA in pool (A). That is, the lack of effect by BSA on products of continuous chloroplast assays containing ATP and CoASH (Fig. 8) is consistent with our pool (A) interpretation.

LACS Activity in Isolated Chloroplasts and a Comparison with Leaf Tissue Labeling Experiments Suggest a Channeled FFA Export Pool-The independent LACS reaction was measured in isolated intact chloroplasts by first loading chloroplasts in situ with FFA accumulated from de novo fatty acid synthesis in the light after which the LACS reaction was run in the dark by adding CoASH (Fig. 4, A and C). The data in Fig. 5 show that the initial LACS rate for a 15 min loading period is not saturated with respect to FFA (it is saturated with respect to ATP and CoASH) and therefore the depletion of pool (B) can be very approximately described by a first order rate equation k(lacs) x [FFA]B. Because in situ-generated FFA can be rapidly removed by BSA (Fig. 6) pool (B) must be freely diffusible. We also know it is mostly associated with the chloroplasts (Fig. 6) unless the reaction mixture contains FFA scavengers. However, we cannot be certain that FFA in pool (B) has partitioned into the thylakoid membranes. Thus the intra-organelle location of FFA pool (B) may be complex and FFA depletion in the dark by LACS may reflect complex kinetic parameters, that is, various FFA diffusion rates (lateral diffusion, flip-flop, partitioning between membrane and media), LACS enzymatic properties, possible LACS feedback inhibition and the possibility of multiple LACS enzymes with different Km values etc. As we assert below that pool (A) represents a channeled system, pool (B) may be considered analogous to what might happen in vivo if there was no channeling (e.g. in a mutant).

The endogenous rate of fatty acid synthesis in leaves is of the order of 1-2 µmol of C2 units/h/mg of chlorophyll (12, 33, 44). This translates to about 15-55 pmol C18 fatty acids/s/mg chlorophyll for plastid export, assuming that 50-90% of newly synthesized fatty acids are exported. In our pea chloroplast assays with exogenous acetate (Fig. 3B) acyl-CoA synthesis (acyl-CoA plus PL) reached a steady state rate of 17.5 pmol C18 fatty acids/s/mg chlorophyll in less than 30 s. This value is lower than the LACS Vmax measured by Joyard and Stumpf (7). These authors characterized the LACS reaction in envelope membrane vesicles isolated from spinach leaf chloroplasts. Assuming that the protein to chlorophyll ratio is 20:1 and that envelope protein accounts for about 1% of total plastid protein their Vmax for 18:1-CoA formation was 85-140 pmol/s/mg chlorophyll. In Fig. 4C the total rate of fatty acid synthesis is 16.7 pmol of fatty acid/s/mg of chlorophyll during the light period, and when the dark reaction is run the initial LACS rate is 220 pmol of fatty acid/s/mg chlorophyll. This is 13-fold greater than the rate of fatty acid synthesis. The maximum rate of the dark LACS reaction is estimated from Fig. 5 as 670 pmol of fatty acid/s/mg chlorophyll. We estimate that our assay concentration of FFA to drive the LACS reaction of intact pea leaf chloroplasts at half maximum velocity as 3 µM, which is well below the 200 µM Km reported by Joyard and Stumpf (7) for oleic acid activation by isolated chloroplast envelopes from spinach. However comparing bulk FFA concentrations between assay systems for envelope vesicles and chloroplasts causes inconsistencies, particularly with respect to the ratio of FFA to membrane lipids etc. The numerical comparison should not be taken literally, but the comparison, albeit for different plant species, does suggest that the LACS enzyme has a much greater affinity for FFA than measured by studies with isolated envelope vesicles.

A more significant comparison is to assess the LACS rates predicted by Fig. 5 when using estimates of the pool size of FFA available for the reaction. For the chloroplast continuous labeling assays in the light the steady state rate (Fig. 3, B and C) is established within 30 s. At this time point the assay concentration of FFA is 0.17 nmol/mg chlorophyll (Fig. 3, B and C), which, according to Fig. 5 should sustain a LACS rate of 4.6 pmol/s/mg chlorophyll. However, the steady state rate of acyl-CoA production of 17.5 pmol/s/mg chlorophyll has been established. Clearly, LACS operating on a freely diffusing FFA pool within the chloroplast cannot explain the onset of the steady state condition. Furthermore, if we take the t1/2 value of 1 s, derived from the in vivo experiments, as the time to fill the transfer FFA pool then the in vivo FFA transfer pools are about 15-55 pmol of C18 fatty acids/mg chlorophyll. These FFA concentrations would support 0.4-1.4 pmol of C18 fatty acids/s/mg chlorophyll according to the concentration dependence curve for chloroplast LACS activity shown in Fig. 5. The discrepancy between acyl-CoA synthesis rates in chloroplasts when fatty acid synthesis is run in the presence of CoASH and ATP (Fig. 3, B and C) and the independently measured LCAS reaction (Fig. 5) of 3.8-fold or greater increases to 37-fold for the comparison involving estimates of in vivo FFA transfer pools. This may in turn be interpreted as 37-fold (or greater, as we are discussing upper limits) concentrated localization of FFA in the export pool in vivo (Fig. 2) compared with freely diffusing condition (Fig. 5). We believe that this leaf tissue to chloroplast comparison is valid because (1) the chloroplasts are synthesizing fatty acids at rates corresponding to in vivo rates of fatty acid synthesis, (2) because the major products from chloroplast assays in the presence of ATP and CoASH are acyl-CoA or acyl-CoA-derived products, (3) because higher maximum velocity and lower FFA affinity for LACS activity are measured with intact organelles loaded with FFA in situ rather than with vesicles challenged with exogenous FFA, and (4) because the t1/2 value to establish the steady state in isolated chloroplasts could easily be an order of magnitude or more lower if rapid kinetic techniques were available for organelles.

Possible Mechanisms for Fatty Acid Export-The analysis of our data presented above allow us to discount the simple free diffusion-based model for FFA export from the plastid, as shown in Fig. 1. There are three generic models (and combinations thereof) to replace the simple diffusion model, namely (1) intervening transfer proteins or (2) a facilitated diffusion process that would mediate the export of fatty acyl groups between the stromal acyl-ACP thioesterase and the outer envelope long chain acyl-CoA synthase, or even (3) a cryptic inner envelope LACS. These models (i.e. for pool (A) in the intact chloroplast assays) are briefly discussed.

For facilitated diffusion without any intervening protein FFA must be deposited in the envelope leaflet where it is most readily used by the LACS enzyme. The limiting in vivo FFA transfer pool size of 1 s calculates to about 0.1 to 0.01 mol% FFA if deposited in a single envelope leaflet lipid phase. In this leaflet the LACS enzyme has a very high affinity for FFA (low Km), so that very little FFA has time to move elsewhere. When the LACS reaction is made inoperative in isolated chloroplasts by lack of CoASH substrate then FFA diffusion to other chloroplast membranes creates pool (B). Little is known about the actual localization, topography and/or vectorial nature of acyl-ACP thioesterases and long-chain acyl-CoA synthetases. Plastid fatty acid synthesis is likely a channeled process (45) associated in vivo with envelope membranes as has been observed for acetyl-CoA carboxylase (46) and as suggested by association of acyl-ACP thioesterase with membrane fractions when imported into the pea chloroplast by in vitro reconstitution experiments.2 There are at least nine LACS genes in Arabidopsis (47). One of them, LACS9 (At1g77590), is demonstrated to be localized to the envelope of the plastid (it is not shown whether LACS9 is located on the outer or inner envelope membrane) and was shown to be responsible for the major in vitro LACS activity on the plastid envelope (48). This gene product may therefore belong to the class responsible for the outer envelope LACS activity described previously (8, 9). To have facilitated FFA diffusion without any additional polypeptides it seems necessary to propose that the fatty acyl-ACP thioesterase binds to the inner envelope, possibly causing a localized membrane perturbation, which can facilitate FFA flip-flop and loads the hydrolyzed product directly onto the inner envelope. Also, the outer envelope long-chain acyl-CoA synthetase would have a domain that would extend across periplasmic space and interact with the inner envelope in a way that would facilitate FFA transfer.

Conceptually it is easier to envisage a protein-mediated channel (Fig. 1). There is evidence from bacterial, yeast and mammalian systems that pairs of proteins are often involved in the vectorial import of fatty acids across biomembranes. In E. coli an outer envelope-localized protein (FadL) was shown to be necessary for the saturable binding and permeation of exogenous long chain fatty acids (49-51). The corresponding acyl-CoA synthetase resides in the inner envelope. In yeast both the FATP protein and the fatty acyl-CoA synthetases of the plasma membrane are interacting components of the fatty acid import machinery (16). In animal cells FATP and acyl-CoA synthetases, both plasma membrane proteins, are required for optimum fatty acid uptake (17). FATP is an AMP-binding domain protein with homology to the very long chain acyl-CoA synthases (18). By contrast, activated long chain fatty acid import across the peroxisomal membrane in yeast is mediated by an ATP binding cassette (ABC) transporter (52, 53). The data we have obtained are consistent with such an intermediate protein between the acyl-ACP thioesterase and LACS. It is apparent that in the absence of the cofactors for the LACS reaction, nascent FFA (pool (A)) is released (to become pool (B)). This release may appear rapid on the time scale of the assays, but could be quite slow relative to the time taken to channel FFA across the plastid envelope to the LACS protein in vivo. If there is only one LACS enzyme then the transport system would also have a finite rate of loading from non-channeled FFA to be able to independently function with freely diffusing FFA. Possible gene candidates for fatty acid transporters might include At4g14070, At3g23790, and the ABC transporters associated with the plastid envelope (54).

The final generic model proposes a latent inner envelope LACS enzyme. Although LACS activity was localized to the outer envelope membrane (8, 9) we can not definitively rule out the possibility of an inner envelope LACS that is intimately linked to FFA export from the inner leaflet of the inner envelope via an acyl-ACP thioesterase complex, but whose activity is cryptic in a simple LACS assay. Among the possible Arabidopsis gene candidates for "cryptic" LACS might be LACS8 (At2g04350) (48) or At4g14070 and At3g23790 (AMP-binding proteins with predicted plastid leader sequences). Disruption of a transporter or cryptic LACS might result in an increased FFA pool in vivo, and if it has a time constant for removal of the order of seconds, analogous to the removal of pool (B) in chloroplast assays by the LACS dark reaction, then the rapid in vivo labeling technique might be useful for the discovery of Arabidopsis lines mutated in these gene products.

In conclusion, we believe that this study defines a set of parameters needed to more fully understand the molecular mechanism of free fatty acid transfer from the plastid. There are several possibilities, as described above, and, given the absence of physical-chemical data on FFA transfer steps for galactolipid-rich membranes and the complexity introduced by the double membrane envelope of plastids, these will not be easy experimental systems. However, the possibility of using reverse genetics in Arabidopsis should open up tractable new avenues to combine with the traditional ways of studying such problems.


   FOOTNOTES
 
* This work was supported by Grant MCB-9817882 from the National Science Foundation, Grant DE-FG02-87ER13729 from the Department of Energy and the Michigan Agricultural Experimental Station. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{ddagger} To whom correspondence should be addressed: Dept. of Plant Biology, Michigan State University, East Lansing, MI 48824-1312. Tel.: 517-355-5237; Fax: 517-353-1926; E-mail: pollard9{at}msu.edu.

1 The abbreviations used are: ACP, acyl carrier protein; FFA, free fatty acid; BSA, bovine serum albumin; acyl-CoA, acyl coenzyme A; LACS, long chain acyl-CoA synthesis; MGDG, monogalactosyldiacylglycerol; PC, phosphatidylcholine; PL, polar lipids; MES, 4-morpholineethanesulfonic acid. Back

2 Unpublished observation communicated by Dr. John Froehlich, MSU-DOE Plant Research Laboratory. Back


   ACKNOWLEDGMENTS
 
We thank Dr. Frederic Beisson and Dr. Ajay W. Tumaney (MSU Department of Plant Biology) for helpful comments and discussions on the article.



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 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
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