The accumulation of triacylglycerols within the endoplasmic reticulum of developing seeds ofHelianthus annuus

Authors


*For correspondence (fax +44 1275 394281; e-mail jon.napier@bbsrc.ac.uk). †These authors contributed equally to this paper.

Summary

Microsomes isolated from developing seeds ofHelianthus annuuswere prepared in a medium which ensured that endoplasmic reticulum (ER)-bound polysomes remained attached to the ER during homogenization. The microsomes were then incubated with the substrates necessary to sustain the synthesis of triacylglycerols (TAGs). Microsomes that contained high activities of the enzymes involved in the synthesis of TAGs (the enzymes of the Kennedy pathway) accumulated TAGs synthesizedin vitro, resulting in a decrease in their buoyant density. These light membrane fractions could therefore be separated on discontinuous sucrose density gradients from microsomes containing low activities of the enzymes of the Kennedy pathway. Analysis of the microsome fractions by 1H-NMR spectroscopy showed that the TAGs synthesized in the microsomesin vitrowere tumbling isotropically in an environment similar to that of the TAGs in oil bodies. Western blot analysis revealed that microsomes which synthesized large amounts of TAGsin vitrowere also substantially enriched in oleosins. In addition, labelling studies indicated that the oleosins newly synthesizedin vitroby ‘run-on' translation of ER-bound polysomes also localized to light membrane fractions. This indicates that oleosins are specifically enriched in regions of the ER involved in the biogenesis of the oil body.

Introduction

The developing seeds of most plant species synthesize triacylglycerols (TAGs) by the Kennedy pathway ( Kennedy 1961;Vogel & Browse 1996). The TAGs are deposited and stored in organelles, termed oil bodies, until germination, at which time they are catabolized as an energy source. Oil bodies comprise a core of TAGs surrounded by an outer coat consisting of a phospholipid (PL) monolayer and a single class of protein termed oleosin (reviewed in Napier et al. 1996 ). Oleosins have relative molecular masses of 15–26 kDa and are thought to stabilize the oil bodies to prevent coalescence, particularly during seed desiccation ( Murphy et al. 1993 ). The enzymes of TAG synthesis have been localized to the ER in the scutellum of developing seeds of maize (Zea mays) ( Cao & Huang 1986) and oil seed rape (Brassica napus) ( Lacey & Hills 1996), indicating that the ER is involved in oil body biogenesis. It is generally accepted that TAGs accumulate between the leaflets of the phospholipid bilayer of the ER ( Wanner & Theimer 1978), but this hypothesis has not been rigorously tested. Wanner & Theimer (1978) proposed that this accumulation results in swelling of the ER to form TAG-filled vesicles, which, upon reaching a critical size, bud from the ER to form an oil body surrounded by a PL monolayer. However, the timing of oleosin insertion into the oil body is unclear and there are different theories as to whether TAG deposition and oleosin synthesis are simultaneous or are spatially and/or temporally separated during seed development. For example, TAG synthesis has been shown to precede oleosin synthesis in B. napus and coriander (Coriandrum sativum) ( Murphy & Cummins 1989;Ross & Murphy 1992), while these events appear to occur concomitantly in Z. mays and soybean (Glycine max) ( Herman 1987;Tzen et al. 1993 ).

Recently, these different findings have been reconciled by a new model of oil body biogenesis, proposed by Sarmiento et al. (1997 ). These researchers suggest that the embryos of most temperate oilseeds will produce oil bodies with minimal protein coats in early stages of seed development. These may then fuse with more oleosin-rich oil bodies produced in mid- to late stages of seed development to form mature oil bodies. It has also been demonstrated that oleosin is co-translationally inserted into the ER ( Hills et al. 1993 ;Loer & Herman 1993) and that this is mediated by a signal recognition particle (SRP) ( Thoyts et al. 1995 ). Furthermore, the same portion of the oleosin seems to be inserted in the ER as in the oil body (our unpublished observations). Thus, the experimental evidence suggests that mature oil bodies form by budding from the ER, as hypothesized by Huang (1992).

It also appears likely that oil body biogenesis occurs in a specific domain of the ER, as proposed by Herman (1995) and Napier et al. (1996 ). Imunocytochemical electron microscopy studies have suggested that oleosins are absent from the bulk cisternae ER in developing cotyledons of G. max ( Herman 1987) and oil bodies have been seen to form at the ends of tubular ER ( Herman 1995). In agreement with these findings, Sarmiento et al. (1997 ) noted that oleosins were absent from the bulk cisternae ER but that they were present in ER which is in close proximity to oil bodies. These observations might therefore suggest that oil bodies are formed at the ER with a dense coat of oleosins. However, the mechanism by which oil bodies were seen to form varied between species in a single electron microscopy study ( Wanner et al. 1981 ), and other researchers have observed oil bodies forming from naked oil droplets in the cytosol ( Adams et al. 1983 ;Bergfeld et al. 1978 ), indicating the need for care in interpreting data obtained from micrographs. Nonetheless, the proposal that oil bodies are synthesized in a subdomain of the ER ( Napier et al. 1996 ;Sarmiento et al. 1997 ) is supported by studies showing that the biosynthesis of TAGs and the enzymes involved in fatty acid modifications appear to be co-localized in specialized subdomains of the ER ( Lacey & Hills 1996;Richards et al. 1993 ;Whitfield et al. 1993 ). It has also been hypothesized that diacylglycerol acyltransferase (DAGAT), which is responsible for the synthesis of TAG from diacylglycerol (DAG), and CDP-choline:choline phosphotransferase (CDP-choline CPT), which is responsible for the synthesis of PC from DAG (and DAG from PC in the reverse direction), must be spatially separated in the ER ( Vogel & Browse 1996). This is because in oilseeds which accumulate unusual fatty acids in their TAGs, there is a concomitant exclusion of these fatty acids from the membrane PC. However, Vogel & Browse (1996) indicated that this exclusion could not be explained on the basis of substrate-selection by DAGAT or CDP-choline CPT alone; instead it may be due to subcellular separation of two the reactions within the ER. It is therefore possible that oil bodies are synthesized in a subdomain of the ER and that oleosins are directly targeted to these subdomains, similar to the segregation of transcripts encoding the prolamin and glutelin storage proteins to the specific regions of the ER observed in developing grains of rice (Oryza sativa) ( Okita et al. 1994 ).

The study by Lacey & Hills (1996) identified a low-density membrane (LDM) fraction which could be separated from the bulk ER due to its high lipid:protein ratio. This LDM fraction contained high specific activities of the enzymes of the Kennedy pathway and may therefore have been derived from regions of the ER that were actively synthesizing oil bodies. In order to extend those studies, it is necessary to separate the regions of the ER that are involved in TAG biosynthesis from those that are not committed to this biosynthetic pathway. Stobart et al. (1986 ) described an assay in which microsomes isolated from developing seeds of safflower (Carthamus tinctorius) synthesized TAGs at very high rates, secreting oil as small 20–40 nm droplets, as judged by light microscopy. We have therefore developed a similar assay using microsomes isolated from developing seeds of sunflower (H. annuus) in order to elucidate the mechanisms of oil body biogenesis. The microsomes that are synthesizing high amounts of TAGs in vitro can be separated using discontinuous sucrose density gradients from other microsomes that do not synthesize TAGs due to differences in their buoyant densities. Analysis of the different regions of the ER separated by this method has allowed us to re-evaluate current models of oil body biogenesis.

Results

Synthesis of TAGs by microsomal membrane fractions

Microsomes isolated from developing cotyledons of sunflower were initially assessed for their ability to synthesize total lipids in vitro, using methods based on those of Stobart et al. (1986 ). Radiolabelled glycerol-3-phosphate was incorporated into lipids of the Kennedy pathway (lyso-phosphatidic acid; phosphatidic acid; diacylglycerol; triacylglycerol), as determined by TLC and scintillation counting. This incorporation was stimulated by the addition of ATP but not GTP ( Fig. 1). The reaction in the presence of ATP was linear for up to 5 min, and the rate of incorporation (5.2 nmol min–1 mg–1 protein) compared well with that reported by Stobart et al. (1986 ) for Carthamus seed microsomes (4 nmol min–1 mg–1 protein). These results indicate that the sunflower microsome system can be used to study the synthesis of TAG and its relationship to oil body biogenesis.

Figure 1.

The effect of ATP and GTP on the synthesis of TAGs during short time-course assays.

Microsomes were incubated with the substrates required for TAG synthesis and ATP (▪), GTP (♦), both NTPs (st) or no NTPs (×).

To confirm that the sunflower microsomes were able to synthesize TAG in vitro, the synthesis of lipids over 4.5 h was measured by adding 1.52 kBq [1-14C]-oleate/50 nmol oleoyl-CoA to the reaction mixture, followed by flotation–centrifugation of the microsomes up through a discontinuous sucrose gradient. This generated four distinct fractions which are hereafter referred to as O (oil body fraction; 8% sucrose), L (light microsomal membrane fraction; 8–19% sucrose), E (standard ER fraction; 19–29% sucrose) and H (heavy microsomal fraction; 29–48% sucrose). These fractions were so named on the basis of where organelles or vesicles might be expected to migrate. Thus the oil body fraction was of such a sucrose density as to be expected to contain oil bodies, but with minimal contamination by ER microsomal vesicles. These ER-derived microsomal vesicles would be expected to be found in the denser sucrose fractions. The lipids were then extracted from these fractions and analysed by TLC and autoradiography. Most of the incorporated radiolabel had entered TAGs, which were present mainly in fractions O and L, though some labelled TAGs were also present in the more dense fractions E and H. A greater proportion of labelled polar lipids relative to TAGs was also present in the more dense fractions (E and H) compared to fractions O and L (Figure 2a). These results suggest that the synthesis of TAGs may be occurring in discrete subdomains of the ER, with microsomes derived from regions of the ER that synthesized TAGs becoming more buoyant during the reaction and thus recovered in fractions O and L. In contrast, the microsomes that did not synthesize high amounts of TAGs remained in lower regions of the gradient. This is illustrated in Fig. 2(b), which shows that the subdomains of the ER that had carried out TAG biosynthesis were clearly separated from the other regions of the ER. Note that although equal amounts of membranes were incubated with or without substrates to support TAG synthesis, accumulation of lipids by the (+) sample has resulted in increased turbidity and therefore apparent differences in amounts of membranes present in the two treatments (see also Stobart et al. 1986 ). No such change in membrane densities was observed in the case of heat-inactivated microsomes incubated in the presence of substrates (data not shown). The rate of TAG synthesis over 4.5 h was 0.4 nmol min–1 mg–1 protein. This is considerably lower than that measured over the first 5 min (5.2 nmol min–1 mg–1; see above) indicating that the rate of reaction had slowed during the later stages of the reaction.

Figure 2.

Figure 2.

Identification of light membrane fractions.

(a) The synthesis of lipids during the in vitro TAG assay. The synthesis of products followed by the incorporation of [1-14C]-oleate into TAGs, polar lipids and other lipids.

(b) Microsomal membrane distribution within a sucrose-step gradient. Membranes incubated in the presence (+) or absence (–) of substrates for TAG synthesis were separated as described in Experimental procedures; a shift in density (arrowed) is observed when the membranes synthesize TAG, resulting a light membrane fraction (L).

Figure 2.

Figure 2.

Identification of light membrane fractions.

(a) The synthesis of lipids during the in vitro TAG assay. The synthesis of products followed by the incorporation of [1-14C]-oleate into TAGs, polar lipids and other lipids.

(b) Microsomal membrane distribution within a sucrose-step gradient. Membranes incubated in the presence (+) or absence (–) of substrates for TAG synthesis were separated as described in Experimental procedures; a shift in density (arrowed) is observed when the membranes synthesize TAG, resulting a light membrane fraction (L).

Lipid composition of microsomal fractions

Microsomes incubated with or without the substrates for TAG synthesis were fractionated by centrifugation and flotation. Fractions were then incubated in the presence of [1-14C]-glycerol-3-phosphate and the radioactive products separated by TLC and quantified by liquid scintillation counting. The same fractions were also analysed by TLC to determine their neutral lipid content while the activity of CDP-choline CPT, which serves as a marker for the ER, was also determined.

Analysis of the lipids in incubated fractions, via TLC, showed that, in the absence of substrate, the major components of the L, E and H fractions were polar lipids which did not migrate (labelled –L, –E and –H; Figure 3a) with low levels of DAG, but little or no TAG. This is consistent with their comprising mainly ER-derived microsomes. Little lipid, either neutral or polar, was recovered from fraction O (labelled –O; Figure 3a), reflecting the low proportion of total lipids present in this fraction.

When total membranes were incubated with substrates for TAG synthesis and then fractionated, clear differences in their composition were observed. Higher proportions of the total membrane lipids were present in fractions O and L which also contained high levels of TAG (see +O and +l), in agreement with the analysis shown in Fig. 2. However, the ratio of TAG:polar lipid in fraction O+ was still lower than in oil bodies from developing sunflower embryos (OB, Figure 3a).

The activities of enzymes of the Kennedy pathway (glycerol-3-phosphate acyltransferase; lyso-phosphatidic acid acyltransferase; phosphatidic acid phosphatase; diacylglycerol acyltransferase) were compared by determining the patterns of labelling in the products (lyso-phosphatidic acid; phosphatidic acid; diacylglycerol; triacylglycerol) present in membrane fractions (O, L, E and H) incubated in the presence of [1-14C]-glycerol-3-phosphate (Figure 3b). The activity of CDP-choline CPT in the membranes was also measured (Figure 3c). Clear differences were observed when the incubation was carried out with or without substrates required for TAG synthesis. In the absence of substrates, lipid synthesis and CDP-choline CPT activity were present mainly in fractions E and H (–E and –H; Figure 3b,c), whereas these activities were concentrated in the lighter fractions (+O and +L; Figure 3b,c) when substrate was supplied. In both cases, the relative activities of CDP-choline CPT corresponded to those of the enzymes of the Kennedy pathway, as indicated by the patterns of incorporation of radiolabelled substrate into TAG, DAG, PA and LPA. This may imply that the enzymes of phosphatidyl choline and TAG synthesis are not spatially separated in the ER of sunflower cotyledons. Sunflower may also differ in this respect from oilseeds (such as Ricinus or Cuphea) which store ‘unusual' fatty acids, in which spatial separation may be required to prevent their incorporation into membranes ( Vogel & Browse 1996).

NMR studies of TAG accumulated in membrane fractions

In order to determine whether the TAG that had been synthesized in vitro by the sunflower microsomes during the 4.5 h reaction had accumulated as discrete droplets within the ER membrane, or whether it was distributed throughout the phospholipid membrane, the buoyant microsomal membrane fractions were investigated by 1H-NMR, using a methodology which had previously been used to examine low density lipoproteins (LDLs) and very low density lipoproteins (VLDLs) from animals. The high-resolution 1H-NMR spectra obtained for oil bodies (Figure 4a) and the membrane fraction L prepared after incubation with substrates for TAG synthesis (Figure 4b) were similar, with a set of signals in the aliphatic region assignable to acyl chains of unsaturated mobile fatty acids. The spectra are very similar to those obtained from animal systems and assigned to mobile TAG using two-dimensional NMR methods ( Williams et al. 1985 ). The spectra obtained for the microsomes in fraction L (Figure 4b) had narrow line-widths and were free of broad underlying signals from proteins, similar to spectra previously obtained from chylomicra ( Williams et al. 1985 ), indicating a highly mobile fraction of TAG free of significant amounts of associated protein. The TAG signals in the spectra from the oil bodies (Figure 4a) had larger line-widths, similar to signals in spectra previously obtained from VLDL and LDL ( Williams et al. 1985 ), indicating lower mobility in this TAG pool with possible spectral contributions from associated oil body proteins. The heavy microsomal fraction (H) did not give significant signals in this region of the 1H-NMR spectrum, consistent with the absence of TAGs (Figure 4c; cf Figure 3a).

Synthesis of oleosin proteins

SDS–PAGE of the microsomal fractions failed to show the presence of appreciable amounts of oleosin proteins (data not shown), which is consistent with an absence of contaminating oil bodies. However, it has been proposed that the synthesis of oleosins is associated with oil body biogenesis ( Napier et al. 1996 ). It is therefore important to determine whether oleosin biosynthesis also occurred in the TAG-synthesizing microsomal fractions.

The four microsomal fractions were initially prepared after incubation with or without substrates for TAG synthesis, and their protein components separated by SDS–PAGE, transferred to nitrocellulose membrane and probed with a polyclonal antiserum specific for sunflower oil body proteins ( Thoyts et al. 1995 ); this antisera recognizes the several different oleosin proteins present on sunflower oil bodies. This showed ( Fig. 5, top panel) that, after incubation with substrate, the highest levels of oleosin were present in the fractions that were most active in TAG synthesis, i.e. fractions O and L (Figure 3a). In contrast, oleosins were predominantly detected in fraction E (i.e. normal ER) when incubated without substrate. The detection of oleosins in the microsomal membrane fractions confirms the role of the ER in oleosin synthesis, as proposed by Thoyts et al. (1995 ) and Napier et al. (1996 ), and also confirms the observations of Sarmiento et al. (1997 ).

Figure 5.

Western blots of fractions O, L, E and H fractionated after incubation with (+) or without (–) the substrates required for TAG biosynthesis probed with either anti-oleosin serum or anti-BiP monoclonal antibodies.

The oleosin antisera recognizes all isoforms of the protein, hence several different cross-reacting bands. Molecular weights of the antigens are indicated.

We wished to determine whether oleosins were selectively accumulating within specific ‘TAG-synthesizing' domains of the ER, or alternatively, that oleosins were present in nascent oil bodies. Therefore, the distribution of oleosins in the microsomal membrane fractions after incubation with the substrates was compared to the distribution of the ER marker protein BiP (binding protein). Since BiP is considered to be evenly distributed throughout the ER, but not found in any other organelles ( Denecke et al. 1995 ;Staehelin 1997), it can be used as a marker for the distribution of the ER in the membrane fractions. Figure 5 (lower panel) clearly shows that in the absence of substrates for TAG synthesis, BiP is found predominantly in the heavy ER fractions (–H). However, in the presence of substrates for TAG synthesis, BiP is redistributed across the fractions in a similar manner as for the oleosin. Thus, oleosins specifically accumulated in buoyant regions of ER that were synthesizing TAGs and are not present as nascent, budded oil bodies. Their pattern of accumulation also mirrored that observed for both the TAGs and the enzymes responsible for their synthesis (cf Figs 3 and 5).

Figure 3.

Figure 3.

In vitro lipid synthesis and distribution.

(a) The lipid composition of fractions O, L, E and H fractionated by TLC after incubation with (+) or without (–) the substrates required for TAG biosynthesis, compared with oil body (OB) control.

(b) The activities of the enzymes of the Kennedy pathway in the above fractions measured by determining the incorporation of [1-14C]-G-3-P into TAG, DAG, PA and LPA.

(c) The activity of the CDP-choline CPT measured by determining the incorporation of [1-14C]-CDP-choline into PC.

Figure 3.

Figure 3.

In vitro lipid synthesis and distribution.

(a) The lipid composition of fractions O, L, E and H fractionated by TLC after incubation with (+) or without (–) the substrates required for TAG biosynthesis, compared with oil body (OB) control.

(b) The activities of the enzymes of the Kennedy pathway in the above fractions measured by determining the incorporation of [1-14C]-G-3-P into TAG, DAG, PA and LPA.

(c) The activity of the CDP-choline CPT measured by determining the incorporation of [1-14C]-CDP-choline into PC.

Figure 3.

Figure 3.

In vitro lipid synthesis and distribution.

(a) The lipid composition of fractions O, L, E and H fractionated by TLC after incubation with (+) or without (–) the substrates required for TAG biosynthesis, compared with oil body (OB) control.

(b) The activities of the enzymes of the Kennedy pathway in the above fractions measured by determining the incorporation of [1-14C]-G-3-P into TAG, DAG, PA and LPA.

(c) The activity of the CDP-choline CPT measured by determining the incorporation of [1-14C]-CDP-choline into PC.

Although these results demonstrated that oleosins accumulated in regions of the ER that synthesized and accumulated TAGs, it is possible that they were synthesized throughout the ER and then migrated to the sites of oil accumulation. We therefore used an in vitro cell-free translation system which allows for the ‘run-on' translation of endogenous transcripts associated with the bound polysomes of microsomal fractions. Rough microsomes were prepared from sunflowers and an aliquot was translated in a cell-free wheatgerm extract containing [35S]-methionine and [35S]-cysteine. This fraction, which contains ‘nascent' oleosin protein, was then recombined with the remaining microsomes, and the TAG synthesis reaction was performed. After the TAG synthesis reaction was completed, the membranes were separated by flotation–centrifugation and the samples were immunoprecipitated using anti-oleosin serum. This allowed the distribution of ‘run-on' synthesized oleosin (i.e. the product of bound ribosomes and their associated endogenous transcripts) to be examined in the case of TAG-synthesizing membranes. Figure 6 clearly shows that the newly synthesized oleosins have distributions dependent on whether or not the microsomal membranes are undergoing TAG synthesis. In the absence of substrates, the bulk of newly synthesized (‘nascent') oleosin is found in the ER fraction (E), whereas in the presence of substrates, the oleosins are found in fractions O and L.

Figure 6.

ER-derived microsomes were incubated in the presence of [35S]-methionine and [35S]-cysteine to allow translation of the associated transcripts; the membranes were then incubated with (+) or without (–) the substrates required for TAG synthesis.

Membranes were then fractionated into fractions O, L, E and H and the oleosins present in these fractions were immunoprecipitated under denaturing conditions, separated by SDS–PAGE and blotted onto nitrocellulose. The radiolabelled oleosins translated in vitro were detected by autoradiography. A clear shift in distribution of newly synthesized oleosin was observed in the presence of substrates for TAG synthesis.

Discussion

An in vitro system to dissect the mechanisms of oil biosynthesis and oil body biogenesis has been developed, based on the safflower microsomal system of Stobart et al. (1986 ). Microsomes isolated from developing seeds of H. annuus synthesized large amounts of TAG over an extended time course, leading to the formation of light fractions which may correspond to intermediates in the formation of oil bodies in vivo. The structures did not develop into true oil bodies in vitro but this is perhaps not surprising considering that the approximate diameter of a microsome is 0.2 μm ( Lord et al. 1973 ) whereas the diameter of an oil body is approximately 0.6–2 μm ( Huang 1992). Thus, a high and sustained level of oil biosynthesis would be required to form true oil bodies and this is unlikely to be achieved in vitro. SDS–PAGE of the ‘intermediate' light microsomal fractions (L) showed no major differences in their polypeptide compositions compared to normal ER fractions which were not actively synthesizing oil (data not shown). However, analyses of the products of incorporation of [1-14C]-glycerol-3-phosphate indicated that they were clearly enriched in enzymes of the Kennedy pathway.

NMR analyses of the light ER fractions showed that the TAGs accumulated in domains where the molecules can freely tumble, similar to the situation in oil bodies. This indicates that the TAGs are not intercalated as monomers within the phospholipid bilayer, since the acyl chains would then be expected to have low mobility, similar to that of acyl chains of membrane phospholipids, with very large line-widths by NMR. Instead, the NMR data are consistent with deposition of the oil as droplets within the phospholipid bilayer of the ER as proposed by Wanner et al. (1981 ) on the basis of ultrastructural studies. However, other ultrastructural studies have interpreted similar results as evidence that oil bodies are formed in the cytosol ( Bergfeld et al. 1978 ), reflecting the problems in using that approach. Here we provide biophysical evidence for the accumulation of TAGs between the bilayers of the ER.

Our results also indicate that oleosins are specifically targeted to and/or accumulate in the region of the ER where oil is being synthesized and deposited, agreeing with models in which the oil bodies are formed from specific domains of the ER with the concurrent formation of an oleosin coat. The molecular basis for the segregation of oleosin to these regions is clearly of interest. Abell et al. (1997 ) recently demonstrated that mutations in the conserved ‘proline knot' sequence ( Huang 1992) present in the hydrophobic central domain of oleosins resulted in the inability of the mutant form of the protein to accumulate in oil bodies, although it was still able to insert into the ER membrane. It will be of great interest to see whether this mutation results in mis-localization of the oleosin within the ER, and/or a reduced ability to interact with the nascent TAG droplets forming between the ER bilayer leaflets.

Our results confirm a number of observations made by Sarmiento et al. (1997 ), most importantly that the oleosin protein can be observed to accumulate in the endomembrane system, presumably prior to oil body assembly. It is interesting to note that in this present study we detected the majority of the oleosin proteins in the ‘normal' ER fractions, with BiP being present in the heavier ER fractions. In the study by Sarmiento et al. (1997 ) this distribution was reversed, with BiP occurring in lighter fractions than oleosin, although the significance of this difference is unclear. However, it is clear from our present studies that oleosin distribution within the ER is altered as a result of TAG synthesis, with oleosin accumulating in lighter, TAG-synthesizing fractions. We also observed that the redistribution of newly synthesized oleosin (as determined by run-on translation of rough ER) occurred in microsomes synthesizing TAG. This provides further evidence for the hypotheses linking TAG accumulation and oleosin synthesis with oil body biogenesis ( Napier et al. 1996 ;Sarmiento et al. 1997 ). One interesting question raised by the model proposed by Sarmiento et al. (1997 ) is why nascent ‘oil bodies' (small phospholipid/TAG droplets with low concentrations of oleosin) do not re-fuse with the ER membrane, which must have a far greater surface area than the small ‘oil bodies' with which fusion is proposed to occur to give rise to mature oil bodies. This may imply the presence of another, as yet unidentified, level of regulation of oil body biogenesis.

However, it is clear that our results, in line with other recent studies ( Abell et al. 1997 ;Sarmiento et al. 1997 ) provide support for models in which the biogenesis of oil bodies is intimately linked with the ER. Perhaps more tellingly, we provide biochemical evidence for the synthesis of TAGs and oleosin by discrete subdomains of the ER.

Experimental procedures

Isolation of microsomes from developing embryos of H. annuus

Plants were grown under the same conditions as the study by Thoyts et al. (1995 ). Embryos were isolated in mid-maturation, approximately 14 days after flowering. The embryos were then homogenized at a ratio of 0.5 g embryo fresh weight to 0.5 ml of homogenization buffer (HB) containing 50 m m HEPES (pH 7.5), 0.25 m sucrose (RNAase-free), 10 m m KCl, 5 m m EGTA, 62.5 m m KC2H3O2, 5 m m MgCl2, 1% (w/v) BSA, 5 m m DTT and an RNAase inhibitor added at a dilution of 1:5000. Cell debris was then removed by centrifugation at 2000 g for 10 min and the supernatant was then centrifuged at 100 000 g for 1 h. The membrane pellet was resuspended in HB and centrifuged at 100 000 g for 1 h to ensure that all oil bodies were removed from the microsome fraction. The microsomes were then resuspended in the same volume of HB as used for the initial homogenization of the embryos.

TAG synthesis reaction

All substrates and co-factors were purchased from Sigma, UK. The reaction was performed at 30°C for 4.5 h and was initiated by adding the following substrates and co-factors to each 0.5 ml portion (approximately 500 μg total protein as estimated by the method of Bradford 1976) of the microsomes: 4 m m G-3-P, 100 μm oleoyl-CoA, 1.2 m m ATP and 100 μm guanosine 5′-triphosphate (GTP). Oleoyl-CoA (50 nmol) was then added every 30 min, ATP (600 nmol) every 60 min and GTP (50 nmol) every 120 min from the initiation of the reaction. Negative controls were incubated with equivalent volumes of H2O added instead of substrates and co-factors. After 4.5 h, the reaction mixtures were brought to a volume of 4 ml with a buffer containing 5 m m EDTA, 48% (w/w) sucrose, and overlaid consecutively with three 2 ml layers of buffer containing 5 m m EDTA and 29% (w/w), 19% (w/w) or 8% (w/w) sucrose, respectively. The discontinuous sucrose density gradients were then centrifuged at 100 000 g for 1 h and the top of the 8% sucrose medium and the interphase of each sucrose step was removed in a volume of 1 ml; these fractions are termed O, L, E and H which correspond to the top of the 8% sucrose medium and the 8/19%, 19/29%, 29/48% sucrose density interphases, respectively.

SDS–PAGE and immunoblotting

Proteins and polypeptides were separated in 0.75 mm biphasic slab gels consisting of a 10% (w/v) polyacrylamide stacking gel and a 16% (w/v) polyacrylamide separating gel using the method of Schagger & von Jagow (1987). Polypeptides were either visualized by staining with Coomassie brilliant blue R-250 or blotted onto nitrocellulose (Anderman, UK) using a semi-dry blotter (Genetic Research Instrumentation Limited, UK) according to the manufacturer's instructions. The blots were then probed with a 1:5000 dilution of rabbit serum raised against H. annuus urea-washed oil bodies ( Thoyts et al. 1995 ) or mouse monoclonal 2E7 which recognizes the ER-retention signal of BiP ( Napier et al. 1992 ). Antibody binding was revealed using the appropriate alkaline phosphatase-conjugated secondary antibody (Sigma, UK).

Enzyme assays

The enzymes of the Kennedy pathway were assayed according to the methods of Lacey & Hills (1996). Time courses were performed in 600 μl reactions containing approximately 300 μg of total protein as estimated by the Bradford assay, and 100 μl portions were taken at the time points shown in Fig. 1. CDP-choline CPT was assayed according to Lord et al. (1973 ).

Lipid analyses

Membrane lipids were extracted and separated according to the methods of Lacey & Hills (1996). After separating lipids by TLC, they were either visualized by iodine staining ( Lacey & Hills 1996) or autoradiography.

NMR analysis

1H-NMR spectra were recorded at 37°C using the JEOL 500 MHz NMR spectrometer at the Bristol Centre for Molecular Recognition. Free induction decays (64 scans) were recorded into 16 K real data points and multiplied by a mild Gaussian window function before Fourier transformation.

Translation of membrane bound polysomes and their localization to ER subdomains

Microsomal membranes (enriched in rough ER and attendant transcripts) were isolated as described above and 20 μl of the resulting 0.5 ml of microsomes were added to a wheatgerm cell-free lysate (Promega, UK) which had previously been clarified by centrifugation at 100 000 g for 15 min to remove endogenous ribosomes. This in vitro translation reaction was incubated for 30 min, according to the supplier's instructions, then added back to the rest of the microsomal fraction and the in vitro TAG synthesis reaction was performed with or without the addition of substrates. After 4.5 h, the membranes were fractionated in discontinuous sucrose density gradients into fractions O, L, E and H. The oleosins in the recovered membranes were then immunoprecipitated under denaturating conditions (50 m m Tris–HCl pH 7.5, 150 m m NaCl, 0.1% w/v SDS, 1% w/v non-idet P-40, 0.5% w/v deoxycholic acid). The immunoprecipitates were separated by SDS–PAGE and blotted onto nitrocellulose and detected by autoradiography.

Acknowledgements

IACR Long Ashton Research Station receives grant-aided support from Biotechnology and Biological Sciences Research Council (BBSRC) of the UK. D.J.L. was supported by a BBSRC ROPA grant CELO4640 ‘Mechanisms of oil body biogenesis in sunflower seeds'. We thank Mike Lewis (MRC, Cambridge) for the gift of the 2E7 antibody, and Keith Stobart for helpful discussions.

Ancillary

Advertisement