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Ketocarotenoid Biosynthesis Outside of Plastids in the Unicellular Green Alga Haematococcus pluvialis

2001; Elsevier BV; Volume: 276; Issue: 8 Linguagem: Inglês

10.1074/jbc.m006400200

1083-351X

Kay Grünewald, Joseph Hirschberg, Christoph Hagen,

Antioxidant Activity and Oxidative Stress

The carotenoid biosynthetic pathway in algae and plants takes place within plastids. In these organelles, carotenoids occur either in a free form or bound to proteins. Under stress, the unicellular green alga Haematococcus pluvialisaccumulates secondary carotenoids, mainly astaxanthin esters, in cytoplasmic lipid vesicles up to 4% of its dry mass. It is therefore one of the favored organisms for the biotechnological production of these antioxidative compounds. We have studied the cellular localization and regulation of the enzyme β-carotene oxygenase in H. pluvialis that catalyzes the introduction of keto functions at position C-4 of the β-ionone ring of β-carotene and zeaxanthin. Using immunogold labeling of ultrathin sections and Western blot analysis of cell fractions, we discovered that under inductive conditions, β-carotene oxygenase was localized both in the chloroplast and in the cytoplasmic lipid vesicles, which are (according to their lipid composition) derived from cytoplasmic membranes. However, β-carotene oxygenase activity was confined to the lipid vesicle compartment. Because an early carotenogenic enzyme in the pathway, phytoene desaturase, was found only in the chloroplast (Grünewald, K., Eckert, M., Hirschberg, J., and Hagen, C. (2000)Plant Physiol. 122, 1261–1268), a transport of intermediates from the site of early biosynthetic steps in the chloroplast to the site of oxygenation and accumulation in cytoplasmic lipid vesicles is proposed. The carotenoid biosynthetic pathway in algae and plants takes place within plastids. In these organelles, carotenoids occur either in a free form or bound to proteins. Under stress, the unicellular green alga Haematococcus pluvialisaccumulates secondary carotenoids, mainly astaxanthin esters, in cytoplasmic lipid vesicles up to 4% of its dry mass. It is therefore one of the favored organisms for the biotechnological production of these antioxidative compounds. We have studied the cellular localization and regulation of the enzyme β-carotene oxygenase in H. pluvialis that catalyzes the introduction of keto functions at position C-4 of the β-ionone ring of β-carotene and zeaxanthin. Using immunogold labeling of ultrathin sections and Western blot analysis of cell fractions, we discovered that under inductive conditions, β-carotene oxygenase was localized both in the chloroplast and in the cytoplasmic lipid vesicles, which are (according to their lipid composition) derived from cytoplasmic membranes. However, β-carotene oxygenase activity was confined to the lipid vesicle compartment. Because an early carotenogenic enzyme in the pathway, phytoene desaturase, was found only in the chloroplast (Grünewald, K., Eckert, M., Hirschberg, J., and Hagen, C. (2000)Plant Physiol. 122, 1261–1268), a transport of intermediates from the site of early biosynthetic steps in the chloroplast to the site of oxygenation and accumulation in cytoplasmic lipid vesicles is proposed. secondary carotenoids β-carotene oxygenase digalactosyldiacylglycerol diacylglyceryltrimethylhomoserine 1-deoxy-d-xylulose-5-phosphate diphenylamine high performance thin layer chromatography high performance liquid chromatography isopentenylpyrophosphate light harvesting complex 2-methyl-d-erythritol-4-phosphate monogalactosyldiacylglycerol norflurazone phosphatidylcholine phytoene desaturase phosphatidylethanolamine phosphatidylglycerol phosphatidylserine triacylglycerol sulfoquinovosyldiacylglycerol Carotenoids play major roles in oxygenic photosynthesis where they function in light harvesting and protect the photosynthetic apparatus from excess light by energy dissipation (1Frank H.A. Cogdell R.J. Photochem. Photobiol. 1996; 63: 257-264Crossref PubMed Scopus (754) Google Scholar). Carotenoids that fulfill these processes are commonly referred to as primary carotenoids, because they are essential for the basic metabolism of the organism. In contrast, secondary carotenoids (SC)1 are defined functionally as carotenoids that are not obligatory for photosynthesis and are not localized in the thylakoid membranes of the chloroplast (2Krishna K.B. Mohanty P. J. Sci. Ind. Res. (India). 1998; 57: 51-63Google Scholar). SC function in specific stages of development (e.g. flower, fruit), mainly for coloration or under extreme environmental conditions. In plants, SC are often accumulated in special structures, for instance in plastoglobuli of chromoplasts. In some green algae, however, SC accumulate outside the plastid in cytoplasmic lipid vesicles. One typical example is the unicellular microalga Haematococcus pluvialis, well known for its massive accumulation of ketocarotenoids, mainly astaxanthin and its acylesters, in response to various stress conditions, e.g.nutrient deprivation or high irradiation (3Boussiba S. Physiol. Plant. 2000; 108: 111-117Crossref Scopus (474) Google Scholar). Different functions of SC in H. pluvialis such as acting as a sunshade (4Hagen C. Braune W. Björn L.O. J. Phycol. 1994; 30: 241-248Crossref Scopus (78) Google Scholar), protecting from photodynamic damage (5Hagen C. Braune W. Greulich F. J. Photochem. Photobiol. B. Biol. 1993; 20: 153-160Crossref Scopus (60) Google Scholar), or minimizing the oxidation of storage lipids (6Sun Z. Cunningham F.X. Gantt E. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 11482-11488Crossref PubMed Scopus (133) Google Scholar) have been proposed. There is growing commercial interest in the biotechnological production of astaxanthin because of its antioxidative properties and the increasing amounts needed as supplement in the aquaculture of salmonoids and other seafood (7Lorenz R.T. Cysewski G.R. Trends Biotechnol. 2000; 18: 160-167Abstract Full Text Full Text PDF PubMed Scopus (902) Google Scholar).H. pluvialis is one of the preferred microorganisms for this purpose because it accumulates SC at up to 4% of its dry mass (3Boussiba S. Physiol. Plant. 2000; 108: 111-117Crossref Scopus (474) Google Scholar). The pathway of astaxanthin biosynthesis in H. pluvialis was elucidated by inhibitor studies (8Harker M. Young A.J. Eur. J. Phycol. 1995; 30: 179-187Crossref Scopus (32) Google Scholar), and most of the involved genes are cloned (6Sun Z. Cunningham F.X. Gantt E. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 11482-11488Crossref PubMed Scopus (133) Google Scholar, 9Hirschberg J. Britton G. Liaaen-Jensen S. Pfander H. Carotenoids. Birkhäuser Verlag, Basel, Switzerland1998: 149-194Google Scholar, 10Linden H. Biochim. Biophys. Acta. 1999; 1446: 203-212Crossref PubMed Scopus (93) Google Scholar). In higher plants and green algae, the carotenoid precursor, isopentenylpyrophosphate (IPP) is derived from the DOXP pathway (synonyms are nonmevalonate or MEP pathway, Ref. 11Lichtenthaler H.K. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1999; 50: 47-65Crossref PubMed Scopus (1100) Google Scholar). For SC synthesis in H. pluvialis this was confirmed with inhibitor studies (12Hagen C. Grünewald K. J. Appl. Bot. 2000; 74: 137-140Google Scholar). The first specific steps in carotenogenesis lead to the formation of the tetraterpene phytoene. Following desaturation and β-cyclization, β-carotene is formed. The subsequent steps in the pathway leading to astaxanthin in H. pluvialis are catalyzed by β-carotene hydroxylase (10Linden H. Biochim. Biophys. Acta. 1999; 1446: 203-212Crossref PubMed Scopus (93) Google Scholar) and β-carotene oxygenase (CRTO, synonym is β-carotene ketolase, BKT; for a recent review see Cunningham and Gantt, Ref. 13Cunningham F.X. Gantt E. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1998; 49: 557-583Crossref PubMed Scopus (786) Google Scholar and Fig.1). Little is known about the regulation of SC synthesis in vivoin response to stress. The gene for CRTO, the enzyme studied in this paper, was cloned from two different strains of H. pluvialisby Lotan and Hirschberg (14Lotan T. Hirschberg J. FEBS Lett. 1995; 364: 125-128Crossref PubMed Scopus (155) Google Scholar) and Kajiwara et al. (15Kajiwara S. Kakizono T. Saito T. Kondo K. Ohtani T. Nishio N. Nagai S. Misawa N. Plant Mol. Biol. 1995; 29: 343-352Crossref PubMed Scopus (138) Google Scholar). A series of β-carotene oxygenases (among them one from H. pluvialis), and bacterial β-carotene hydroxylases were characterized in vitro with respect to substrate specificity and cofactor requirements (16Fraser P.D. Miura Y. Misawa N. J. Biol. Chem. 1997; 272: 6128-6135Abstract Full Text Full Text PDF PubMed Scopus (149) Google Scholar, 17Fraser P.D. Shimada H. Misawa N. Eur. J. Biochem. 1998; 252: 229-236Crossref PubMed Scopus (76) Google Scholar). Moreover, conversion of β-carotene by cell extracts of H. pluvialis was reported (18Chumpolkulwong N. Kakizono T. Ishii H. Nishio N. Biotechnol. Lett. 1997; 19: 443-446Crossref Scopus (24) Google Scholar). Recently, we have studied regulation and compartmentation of phytoene desaturase (PDS), an early enzyme of the carotenoid biosynthetic pathway (19Grünewald K. Eckert M. Hirschberg J. Hagen C. Plant Physiol. 2000; 122: 1261-1268Crossref PubMed Scopus (97) Google Scholar). The enzyme is up-regulated at the mRNA level during SC synthesis and localized exclusively in the chloroplast. This is consistent with the common hypothesis that in plants including algae carotenoids are synthesized exclusively within plastids (11Lichtenthaler H.K. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1999; 50: 47-65Crossref PubMed Scopus (1100) Google Scholar). H. pluvialis is distinguished in that it accumulates large amounts of carotenoids in lipid vesicles outside the plastid (3Boussiba S. Physiol. Plant. 2000; 108: 111-117Crossref Scopus (474) Google Scholar, 20Santos F. Mesquita J.F. Cytologia. 1984; 49: 215-228Crossref Scopus (81) Google Scholar). This has given rise to speculation about the possible existence of a biosynthetic pathway specific for secondary carotenogenesis that is localized in the cytoplasm, as was supported by the existence of two different IPP isomerases in H. pluvialis (6Sun Z. Cunningham F.X. Gantt E. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 11482-11488Crossref PubMed Scopus (133) Google Scholar). However, no extra pathway specific for SC biosynthesis in the cytosol of H. pluvialis was found at the level of PDS (19Grünewald K. Eckert M. Hirschberg J. Hagen C. Plant Physiol. 2000; 122: 1261-1268Crossref PubMed Scopus (97) Google Scholar). It was therefore hypothesized that carotenoids are transported from the site of biosynthesis (chloroplast) to the site of accumulation (cytoplasmic lipid vesicles). Here, we present a study of the origin of these lipid vesicles as well as regulation and compartmentation of the SC biosynthetic-specific ketolase CRTO in flagellates of H. pluvialis using immunolocalization and cell fractionation techniques. Our results indicate that the last oxygenation steps in the astaxanthin biosynthesis pathway take place outside the plastid in the cytoplasmic lipid vesicles and is discussed relative to the role of this sequestering structure in SC accumulation. H. pluvialis Flotow (No.192.80, culture collection of the University of Göttingen, Germany; synonym: Haematococcus lacustris (Girod) Rostafinski) was grown autotrophically in a two-step batch cultivation system as described (21Grünewald K. Hagen C. Braune W. Eur. J. Phycol. 1997; 32: 387-392Crossref Scopus (3) Google Scholar). Following precultivation for 5 days at 25 μmol of photons m−2 s−1 of white fluorescent light (Osram L36/W25, Berlin, Germany), flagellates in the logarithmic growth phase were exposed to SC-inducing conditions (nitrate-deprived medium and 150 μmol of photons m−2s−1 of continuous white light) leading to accumulation of SC in the flagellated developmental state of H. pluvialis(21Grünewald K. Hagen C. Braune W. Eur. J. Phycol. 1997; 32: 387-392Crossref Scopus (3) Google Scholar). These flagellates surrounded by a thin extracellular matrix are more accessible to biochemical and ultrastructural analysis than the thick-walled and resistant aplanospore state. Photon flux densities were measured using a LI-189 photometer (LI-COR, Lincoln, NE), and cell number was determined using a Cell Counter Casy 1 (Schärfe Systems, Reutlingen, Germany). At the time points specified, sample aliquots corresponding to a defined cell number were collected by centrifugation at 1,400 × g for 2 min. Cell fractions were prepared by gentle filtration rupture that produced less contamination of the lipid vesicle fraction by light harvesting complexes (LHC) and chlorophylls than sonication. Aliquots of cells were harvested by centrifugation at 1,400 × g for 2 min and resuspended in break buffer consisting of 0.1 m Tris-HCl, pH 6.8, 5 mm MgCl2, 10 mm NaCl, 10 mm KCl, 5 mm Na2EDTA, 0.3m sorbitol, 1 mm aminobenzamidine, 1 mm aminohexanacid, and 0.1 mmphenylmethylsulfonyl fluoride. The hyperosmotically shocked cells were broken by passage through a 10-μm isopore polycarbonate filter (Millipore, Eschborn, Germany). The filtrate was centrifuged at 10,000 × g for 10 min at 4 °C to yield a chloroplast and cell debris pellet. The supernatant was transferred to a fresh tube and centrifuged again at 10,000 × g for 10 min at 4 °C. The suspension below the lipid vesicle fraction floating on top was transferred to a fresh tube and centrifuged at 76,000 × g for 2 h at 4 °C. The resulting microsome pellet was separated from the supernatant fraction. All fractions were stored at −20 °C. Cell aliquots or lipid vesicle preparations were extracted essentially as described (22Folch J. Lees M. Stanley G.H.S. J. Biol. Chem. 1957; 226: 497-509Abstract Full Text PDF PubMed Google Scholar). Lipids were then separated on HPTLC plates (Merk, Darmstadt, Germany), developed for two-thirds of the plate in chloroform/methanol/acetic acid/water, 73:25:2:4 (v/v/v/v) to separate the polar lipids and subsequently, in a second development, with hexane/diethylether/acetic acid, 85:20:1.5 (v/v/v) for the whole plate to separate the neutral lipids from the pigments. Lipids were identified by cochromatography of standard substances and by color reaction with different spray reagents (ninhydrin for free amines of phosphatidylethanolamine (PE) and phosphatidylserine (PS); α-naphthol for glyco- and sulfolipids; molybdenium blue for phospholipids; Dragendorff's reagent for quarternary amines, phosphatidylcholine (PC) and diacylglyceryltrimethylhomoserine (DGTS)). Quantification of individual lipids was performed densitometrically after visualization by Godin's spray reagent (23Godin P. Nature. 1954; 174: 134Crossref Scopus (174) Google Scholar) and calibration with standard substances. For quantification of DGTS and PS, calibration data of PE and PC were used, respectively. At the onset of SC-inducing conditions, diphenylamine (DPA, Sigma) was added to a final concentration of 30 μm. After 3 days, cells were washed three times, resuspended in fresh nitrate-deprived medium, and incubated for 2 more days under SC inductive conditions with 5 μm norflurazone (NF; SAN 9879; Sandoz Basel, Switzerland), 30 μm DPA or 30 μm DPA plus 5 μm NF, respectively. The 17-amino acid peptide LPHCRRLSGRGLVPALA, corresponding to the C terminus (residues 304–320) in the predicted sequence of BKT (15Kajiwara S. Kakizono T. Saito T. Kondo K. Ohtani T. Nishio N. Nagai S. Misawa N. Plant Mol. Biol. 1995; 29: 343-352Crossref PubMed Scopus (138) Google Scholar) and residues 315–329 (with the last three amino acids missing) in the predicted sequence of CRTO (14Lotan T. Hirschberg J. FEBS Lett. 1995; 364: 125-128Crossref PubMed Scopus (155) Google Scholar), was chemically synthesized and purified (Alpha Diagnostics International, San Antonio, TX). The peptide was coupled to thyroglobulin by means of glutaraldehyde and used for immunization of rabbits to raise polyclonal antibodies as described (24Eckert M. Gabriel J. Birkenbeil H. Greiner G. Rapus J. Gäde G. Cell Tissue Res. 1996; 284: 401-413Crossref PubMed Scopus (13) Google Scholar). The raw serum was deployed without further purification. Cell pellets were thawed on ice, suspended in break buffer, and broken by sonication for 1 min on ice (Vibra-Cell 72405 sonication processor, Sonics & Materials, Danbury, CT; pulse mode, 0.75 s on, 1 s off, 60 watts output). Break buffer with SDS was added to yield a final concentration of 2% SDS (w/v). Solubilization, especially of hydrophobic proteins like CRTO, was carried out upon shaking at 2,000 rpm for 2 h at 20 °C. Samples were centrifuged to remove unsolubilized material, and sample loading buffer to a final concentration of 50 mm Tris-HCl, pH 6.8, 2% SDS (w/v), 10% glycerol (v/v), and 0.01% bromphenol blue was added. Cell fractions were thawed on ice and resuspended in break buffer with 2% SDS (w/v), and solubilization was performed as described for total cell extracts. Before loading cell aliquots, samples were boiled for 5 min. Proteins were separated on 12% SDS-polyacrylamide gels essentially as described (25Laemmli U.K. Nature. 1970; 227: 680-685Crossref PubMed Scopus (207263) Google Scholar). For Western blot analysis, the gels were electrophoretically transferred semidry onto nitrocellulose membranes (Schleicher & Schuell, Dassel, Germany) and treated with Ponceau S for staining the protein ladder transiently. Membranes were blocked in blocking buffer containing 5% (w/v) nonfat dry milk, 1% Tween 20 (v/v), 150 mm NaCl, and 25 mm Tris-HCl, pH 7.6 at 4 °C overnight. Then the blots were challenged with anti-CRTO antibodies in blocking buffer at 1:250 dilution for 1 h at 4 °C and thereafter with secondary antibody alkaline phosphatase conjugates (Bio-Rad, Munich, Germany) used at 1:500 dilution. After the chromogenic reaction with 5-bromo-4-chloro-3-indolyl phosphate (BCIP) and nitro blue tetrazolium chloride (NBT), the labeling was quantified using densitometry (Scanpack 3.0, Biometra, Göttingen, Germany). Total protein content was determined by means of the detergent compatible protein assay kit (Bio-Rad, Munich, Germany). For ultrastructural examination, algal cells were harvested at 550 ×g for 3 min and then fixed with 0.7% glutaraldehyde, 0.8% paraformaldehyde, and 1% OsO4 simultaneously in growth medium for 25 min at 4 °C. After several washes in distilled water the specimens were dehydrated in graded ethanol series. The 70% ethanol step was performed in the presence of 3% uranylacetate for 10 min. Cells were embedded in LR Gold (London Resin, London) according to the manufacturer's instructions. Before immunogold labeling, ultrathin sections were cut as described (19Grünewald K. Eckert M. Hirschberg J. Hagen C. Plant Physiol. 2000; 122: 1261-1268Crossref PubMed Scopus (97) Google Scholar) and etched to unmask antigenic determinants (26Marotta L. Shero M. Carter J.M. Klohs W. Apicella M.A. J. Immunol. Methods. 1984; 71: 69-82Crossref PubMed Scopus (2) Google Scholar). Etching was done by floating grids section side down on 2% H2O2 for 2 min at room temperature followed by three washes on distilled water. The grids were exposed to anti-CRTO antibody at 1:100 dilution, and immunogold labeling was performed as described (19Grünewald K. Eckert M. Hirschberg J. Hagen C. Plant Physiol. 2000; 122: 1261-1268Crossref PubMed Scopus (97) Google Scholar). Subsequent to poststaining with 3% aqueous uranylacetate (w/v) for 5 min and 1% aqueous lead citrate (w/v) for 20 s, immunogold-labeled sections were examined in a Zeiss EM 900 electron microscope (Carl Zeiss, Oberkochen, Germany) at 80 kV. Incubations were carried out in a total volume of 600 μl under conditions essentially as reported (16Fraser P.D. Miura Y. Misawa N. J. Biol. Chem. 1997; 272: 6128-6135Abstract Full Text Full Text PDF PubMed Scopus (149) Google Scholar,17Fraser P.D. Shimada H. Misawa N. Eur. J. Biochem. 1998; 252: 229-236Crossref PubMed Scopus (76) Google Scholar). Cell fraction aliquots of 107 cells were suspended in break buffer (0.1 m Tris-HCl, pH 6.8, 0.3 msorbitol, 1 mm aminobenzamidine, 1 mmaminohexanacid, 1 mm dithiothreitol, 0.1 mmphenylmethylsulfonyl fluoride) in a total volume of 300 μl. Following the addition of 295 μl of cofactor buffer (5 mm ascorbic acid, 1 mm dithiothreitol, 0.5 mmFeSO4, 0.1% deoxycholate (w/v), 0.5 mm2-oxoglutarate) and brief mixing, the reaction was initiated by addition of 5 μl of a 1% β-carotene stock solution (w/v) in chloroform. In parallel samples, 100 μm DPA were added to inhibit β-carotene oxygenase. Incubation was performed under continuos stirring for 2 h in the dark at 30 °C. Reactions were terminated by freezing the samples in liquid nitrogen. Cell pellets were extracted quantitatively in 100% acetone at 4 °C, and the pigment content was determined spectrophotometrically according to Lichtenthaler (27Lichtenthaler H.K. Methods Enzymol. 1987; 148: 350-382Crossref Scopus (9280) Google Scholar). Fractions were freeze-dried, and carotenoids were extracted with 200 μl of acetone (the chloroplast fraction was extracted with 500 μl of acetone) at 4 °C. In vitroincubations were freeze-dried, and pigments were extracted with acetone, at 4 °C. Prior to HPLC analysis, samples were filtered and 20% water (v/v) was added. HPLC analysis was performed as described (21Grünewald K. Hagen C. Braune W. Eur. J. Phycol. 1997; 32: 387-392Crossref Scopus (3) Google Scholar). Lipid profiles of total extracts from cells drawn after 4 days of exposure to conditions inductive for SC synthesis revealed massive accumulation of TAG during SC synthesis (Fig. 2). Concomitantly, the amount of most membrane lipids, especially of MGDG, decreased whereas that of DGDG and DGTS increased slightly (TableI). Analysis of the lipid vesicles formed under inductive conditions revealed triglycerides as their predominant lipid class. Membrane lipids accounted for less than 5% (w/w) of total lipids in this fraction. No MGDG was detectable, and DGDG and DGTS made up half of the membrane lipids in this fraction besides significant amounts of PC and of PE.Table IChanges in the content of polar and neutral lipids during accumulation of SC in H. pluvialis flagellates and their distribution in SC-containing lipid vesiclesTime after onset of inductionExtractLipids1-aAbbreviations of lipids are as in Abbreviations footnote.TAGMGDGDGDGSQDGDGTSPEPCPGPSdaypg per cell1-cSE (n = 4 to 8) did not exceed 10%.0Total cellsn.d.1-bn.d., not detectable.49.974.826.687.407.793.652.291.214Total cells76.0912.205.816.208.406.792.111.511.32% of total lipid mass4Lipid vesicles95.46n.d.1.070.541.190.660.680.140.261-a Abbreviations of lipids are as in Abbreviations footnote.1-b n.d., not detectable.1-c SE (n = 4 to 8) did not exceed 10%. Open table in a new tab Application of low concentrations of DPA under conditions inductive for synthesis of SC led to accumulation of β-carotene instead of ketocarotenoids inH. pluvialis (8Harker M. Young A.J. Eur. J. Phycol. 1995; 30: 179-187Crossref Scopus (32) Google Scholar, 21Grünewald K. Hagen C. Braune W. Eur. J. Phycol. 1997; 32: 387-392Crossref Scopus (3) Google Scholar, 28Fan L. Vonshak A. Gabbay R. Hirschberg J. Cohen Z. Boussiba S. Plant Cell Physiol. 1995; 36: 1519-1524Google Scholar). Lipid vesicles in the cytoplasm of treated cells appeared yellow instead of red in control samples, suggesting that β-carotene accumulated in the cytoplasm (8Harker M. Young A.J. Eur. J. Phycol. 1995; 30: 179-187Crossref Scopus (32) Google Scholar). To substantiate this observation, we determined the pigment composition in different cellular compartments. Results revealed a predominant accumulation of β-carotene inside the lipid vesicles of DPA-treated cells (Fig. 3). To test if this extraplastidic β-carotene can be converted to astaxanthin, DPA was removed concomitantly with the addition of NF to inhibit carotenoidde novo synthesis at the level of phytoene desaturation. Beside the known bleaching effect of NF leading to a reduced amount in total carotenoids, a significant decrease in the ratio of β-carotene to ketocarotenoids occurred inside the lipid vesicles (Fig. 3). The pattern of SC in this fraction did not differ significantly from untreated samples consisting mostly of mono- and diesters of astaxanthin. Compartmentation studies and regulation analysis of the late steps in SC biosynthesis inH. pluvialis require specific antibodies against the enzymes involved. Attempts to obtain antibodies against the His-tagged C terminus of CRTO, encompassing two-thirds of the polypeptide overexpressed in Escherichia coli, were unsuccessful. Despite poor expression and isolation difficulties because of pronounced hydrophobic behavior of the protein, necessary amounts of the antigen were recovered by Ni2+-affinity chromatography and subsequent purification steps. The generated polyclonal antibodies recognized a series of proteins on Western blots and did not meet the needs for localization experiments, even after parallel immunization experiments and various purification approaches by affinity chromatography. Interestingly, we noticed an increasing oligomerization tendency of the overexpressed antigen up to the octamer, even under denaturating SDS-polyacrylamide gel electrophoresis conditions, dependent on storage time. Finally, we immunized rabbits with a 17-mer synthetic oligopeptide corresponding to the C-terminal part of the predicted structure of CRTO. Database searches revealed no counterparts of this peptide among plant amino acid sequences. The ability of the antibodies to recognize less than 30 ng of CRTO in Western blots was verified with the E. coli-overexpressed C-terminal part of the enzyme (data not shown). Abundance of CRTO was examined in total cell extracts of start samples and of samples taken 1, 2, 3, 4, and 7 days after inducing SC synthesis in the flagellates of H. pluvialis by intense illumination and nitrate deprivation (Fig.4 A). No CRTO was observed before the second day after induction. After this time, the amount of a 34-kDa protein increased rapidly in parallel to SC accumulation (Fig.4 B). The apparent molecular mass of the recognized protein was ∼3 kDa smaller than predicted from the cDNA sequence of CRTO (14Lotan T. Hirschberg J. FEBS Lett. 1995; 364: 125-128Crossref PubMed Scopus (155) Google Scholar). The preimmune serum did not detect this polypeptide (not shown). The antibodies against the C-terminal 17-mer of CRTO were tested on LR Gold sections of H. pluvialis flagellates that have previously been shown to present the best combination for structural preservation and maintenance of antigenic structures (19Grünewald K. Eckert M. Hirschberg J. Hagen C. Plant Physiol. 2000; 122: 1261-1268Crossref PubMed Scopus (97) Google Scholar). To prevent the extraction of cellular lipids during dehydration steps and to ensure full preservation, high pressure cryofixation in combination with cryodehydration was applied. However, despite a number of modifications of the preparation protocol, the structure of lipid vesicles could not be improved. Thus, an etching technique was chosen as described (26Marotta L. Shero M. Carter J.M. Klohs W. Apicella M.A. J. Immunol. Methods. 1984; 71: 69-82Crossref PubMed Scopus (2) Google Scholar). During the ethanolic dehydration process before embedding, the lipid vesicles remained intact because of lipid cross-linking by OsO4. Probing the sections with polyclonal antibodies against different photosynthetic proteins (19Grünewald K. Eckert M. Hirschberg J. Hagen C. Plant Physiol. 2000; 122: 1261-1268Crossref PubMed Scopus (97) Google Scholar) did not reveal any signals because of masking of antigenic determinants by the fixative. To unmask antigenic determinants, sections were exposed to hydrogen peroxide for a defined time span. Accessibility of antigenic determinants after etching was confirmed with anti-LHC and anti-PDS antibodies, which detected the corresponding polypeptides as reported previously (19Grünewald K. Eckert M. Hirschberg J. Hagen C. Plant Physiol. 2000; 122: 1261-1268Crossref PubMed Scopus (97) Google Scholar). After challenging the sections with the polyclonal antibodies raised against the CRTO C-terminal 17-mer, two cell compartments became specifically immunogold-labeled in the course of SC synthesis, namely the chloroplast and, becoming dominant, the lipid vesicles (Fig.5, TableII). Labeling of the latter compartment was not restricted to the periphery, but was scattered throughout the vesicles. The only notable signal in the cytosol was obtained after 2 days of inductive conditions (15%) and was localized in close contact to the Golgi cisternae (not shown). No specific labeling was observed when sections were probed with preimmune serum (not shown).Table IIDistribution of β-carotene oxygenase in the major compartments of flagellates at different stages of SC synthesisTime of exposureMean of gold particles per cellDistribution of immunogold labeling in various compartments2-aAfter subtraction of the corresponding count found in preimmune labeled sections.2-bS.E. of day 4 and day 0; days 2 and 7 are given for n = 24 and n = 12, respectively.ChloroplastNucleusCytosolLipid vesiclesResidueday%Start3.2 ± 2.231.6 ± 60.15.3 ± 13.465.8 ± 17.40.0 ± 0.0−2.6 ± 9.2236.8 ± 2.664.6 ± 5.40.0 ± 0.014.7 ± 2.315.0 ± 2.15.7 ± 1.3479.8 ± 1.055.1 ± 11.50.7 ± 0.55.1 ± 0.737.1 ± 3.42.0 ± 0.3729.8 ± 2.930.7 ± 7.4−4.2 ± 3.0−3.0 ± 1.974.7 ± 4.61.8 ± 1.92-a After subtraction of the corresponding count found in preimmune labeled sections.2-b S.E. of day 4 and day 0; days 2 and 7 are given for n = 24 and n = 12, respectively. Open table in a new tab To ascertain the results from the immunogold localization experiments, four cellular fractions were obtained: (i) a pellet containing mainly the chloroplast, (ii) a supernatant fraction, (iii) microsomes and cytoplasmic membranes, and (iv) the lipid vesicles (19Grünewald K. Eckert M. Hirschberg J. Hagen C. Plant Physiol. 2000; 122: 1261-