Plant Physiology, April 2000, Vol. 122, pp. 1261–1268, www.plantphysiol.org © 2000 American Society of Plant Physiologists
Phytoene Desaturase Is Localized Exclusively in the
Chloroplast and Up-Regulated at the mRNA Level during
Accumulation of Secondary Carotenoids in Haematococcus
pluvialis (Volvocales, Chlorophyceae)1,2
Kay Grünewald*, Manfred Eckert, Joseph Hirschberg, and Christoph Hagen
Institute of General Botany, Friedrich-Schiller-University Jena, Am Planetarium 1, D–07743 Jena, Germany
(K.G., C.H.); Institute of General Zoology, Friedrich-Schiller-University Jena, Erbertstrasse 1,
D–07743 Jena, Germany (M.E.); and Department of Genetics, The Hebrew University of Jerusalem,
Jerusalem, 91904 Israel (J.H.)
The unicellular green alga Haematococcus pluvialis Flotow is
known for its massive accumulation of ketocarotenoids under various stress conditions. Therefore, this microalga is one of the favored organisms for biotechnological production of these antioxidative compounds. Astaxanthin makes up the main part of the
secondary carotenoids and is accumulated mostly in an esterified
form in extraplastidic lipid vesicles. We have studied phytoene
desaturase, an early enzyme of the carotenoid biosynthetic pathway. The increase in the phytoene desaturase protein levels that
occurs following induction is accompanied by a corresponding
increase of its mRNA during the accumulation period, indicating
that phytoene desaturase is regulated at the mRNA level. We also
investigated the localization of the enzyme by western-blot analysis
of cell fractions and by immunogold labeling of ultrathin sections
for electron microscopy. In spite of the fact that secondary carotenoids accumulate outside the chloroplast, no extra pathway specific
for secondary carotenoid biosynthesis in H. pluvialis was found, at
least at this early stage in the biosynthesis. A transport process of
carotenoids from the site of biosynthesis (chloroplast) to the site of
accumulation (cytoplasmatic located lipid vesicles) is implicated.
During the last decade, much of the research on carotenoid biosynthesis has been at the genetic level, i.e. the
elucidation of the respective genes and enzymes. Less is
known about the regulation of the processes involved. The
unicellular green alga Haematococcus pluvialis (Volvocales)
is known for its massive accumulation of the ketocarotenoid astaxanthin in the esterified form in response to, for
example, nutritional or light stress conditions. The increasing commercial interest in astaxanthin results from its antioxidative properties (Nishino, 1997), which are important
1
This study was supported in part by the Deutscher Akademischer Austauschdienst (short-term fellowship to K.G.), by the
Thüringer Ministerium für Forschung, Wissenschaft und Kultur
(grant no. B301– 69013), and by a graduate fellowship from the
Freistaat Thüringen for K.G.
2
This paper is dedicated to the occasion of the 65th birthday of
Prof. Dr. Wolfram Braune.
* Corresponding author; e-mail [email protected];
fax 49 –3641–949225.
for the pharmaceutical and cosmetic industries. It is also
used in huge amounts as a nutritional supplement in the
aquaculture of salmonoids (Wathne et al., 1998). Biological
functions of so-called secondary carotenoids (SC) as sunshade (Hagen et al., 1994) and as protection against photodynamic damage (Hagen et al., 1993) have been elucidated in H. pluvialis.
Carotenoids are synthesized from the universal isoprenoid precursor isopentenylpyrophosphate (IPP). The more
recently discovered DOX-P-pathway for IPP biosynthesis,
named after the first intermediate 1-deoxy-d-xylulose-5phosphate (synonyms: non-mevalonate pathway), is located in the chloroplast and seems to be the favored, if not
the only, pathway used for the synthesis of carotenoids in
higher plants and green algae (Lichtenthaler et al., 1999).
By head-to-tail additions and the subsequent action of the
phytoene synthase, the first tetraterpene carotenoid, phytoene, is formed. The following desaturation steps leading
to ␨-carotene are catalyzed by phytoene desaturase (PDS),
the enzyme studied here. The subsequent biosynthetic
steps resulting in the different carotenoids also occurring in
H. pluvialis are summarized in Cunningham and Gantt
(1998). A short scheme of the sequences leading to astaxanthin esters and indicating also the ketocarotenoid specific enzyme ␤-carotene oxygenase (CRTO; synonym:
␤-carotene ketolase) is given in Figure 1. The gene for this
specific enzyme was cloned from two different strains of H.
pluvialis by Lotan and Hirschberg (1995) and Kajiwara et al.
(1995).
According to hydropathy analysis, substrate hydrophobicity, and chloroplast subfractionation, the active plant
PDS is assumed to be tightly bound or even intrinsic of
membranes (Camara et al., 1982; Al-Babili et al., 1996).
Linden et al. (1993) showed by immunogold labeling a
thylakoid localization of the enzyme in chloroplasts isolated from spinach. The regulation of the enzyme was
investigated for different organs and respective organelles
in a number of carotenoid-accumulating plants such as
tomato (Pecker et al., 1992; Giuliano et al., 1993), pepper
(Römer et al., 1993), and narcissus (Al-Babili et al., 1996). In
the green alga Dunaliella bardawil, both mRNA and protein
level of PDS remained constant during the massive
1261
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1262
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Plant Physiol. Vol. 122, 2000
MATERIALS AND METHODS
Organism and Growth Conditions
Haematococcus pluvialis Flotow (no. 192.80, culture collection of the University of Göttingen, Germany; synonym:
Haematococcus lacustris [Girod] Rostafinski) was grown autotrophically in 100-mL Erlenmeyer flasks at 20°C (⫾2°C)
in a medium described by Hedlich (1982) with the addition
of 0.3 ␮m thiamine. The flasks containing 50 mL of algae
suspension were neither shaken nor aerated. The accumulation of SC was achieved by nitrate deprivation of the
medium to 5% and stronger light intensities (150 ␮mol
photons m⫺2 s⫺1 of continuous white light) in the two-step
batch cultivation system described in Grünewald et al.
(1997).
Photon flux densities were measured using a photometer
(model LI-189, LI-COR, Lincoln, NE). Cell number was
determined using a cell counter (model Casy 1, Schärfe
Systems, Reutlingen, Germany).
Preparation of Cell Fractions
Figure 1. Pathway of secondary carotenoid synthesis in H. pluvialis.
Data are presented in this paper for the enzymes shown in black
boxes. Enzyme designation is according to the corresponding gene:
CRTL-B, lycopene ␤-cyclase; CRTO, ␤-carotene oxygenase; CRTR-B,
␤-ring hydroxylase; GGPS, geranylgeranyl diphosphate synthase; IPI,
isopentenyl diphosphate isomerase; PDS, phytoene desaturase; PSY,
phytoene synthase; ZDS, ␨-carotene desaturase.
Aliquots of cells were harvested by centrifugation at
1,400g for 2 min and re-suspended in break buffer consisting of 0.1 m Tris-HCl, pH 6.8, 5 mm MgCl2, 10 mm NaCl, 10
mm KCl, 5 mm Na2-EDTA, 0.3 m sorbitol, 1 mm aminobenzamidine, 1 mm aminohexanacid, and 0.1 mm phenylmethylsulfonyl fluoride (PMSF), and passed through a 10-␮m
isopore polycarbonate filter (Millipore, Eschborn, Germany). The filtrate was centrifuged for 10 min at 16,000g to
yield a chloroplast and cell debris pellet. All supernatant
was transferred to a fresh tube and centrifuged at 108,000g
for 1 h. The resulting three fractions, the microsome pellet,
the supernatant fraction, and the lipid vesicles floating on
top, were separated and stored at ⫺20°C.
Protein Analysis
␤-carotene synthesis on high-light conditions. In this unicellular alga the secondary ␤-carotene accumulation in intraplastidic lipid droplets is controlled by the formation of
the sequestering structures (Rabbani et al., 1998).
In plants, carotenoids are reported to be synthesized
exclusively within plastids (Lichtenthaler, 1999, and refs.
cited therein). H. pluvialis is unique among all algae and
plants that have been studied to date in that it accumulates
huge amounts of carotenoids in lipid vesicles outside the
plastid (Mesquita and Santos, 1984). This has given rise to
speculations about the possible existence of a biosynthetic
pathway specific for secondary carotenogenesis that is localized in the cytoplasm. This possibility is supported by
the cytosolic accumulation of the final product and was
addressed recently by Sun et al. (1998), who studied the
function of two IPP isomerases found in H. pluvialis. However, data on the cellular localization of both isoenzymes
were not presented by the authors.
We report on the regulation and compartmentation of
PDS in flagellates of H. pluvialis grown in the cultivation
system described previously (Grünewald et al., 1997).
Proteins were separated on 12% (w/v) SDS-PAGE gels.
For western analysis, the gels were electrophoretically
transferred semidry to nitrocellulose membranes (Schleicher & Schuell, Dassel, Germany), and treated with Ponceau S for staining the protein ladder transiently. Primary
antibodies were used at a dilution of the raw serum given
in the text. Secondary antibody conjugates with alkaline
phosphatase were used for immunostaining with 5-bromo4-chloro-3-indolyl phosphate (BCIP) and nitroblue tetrazolium chloride (NBT). Quantification of labeling was done
using densitometry (Scanpack 3.0, Biometra, Göttingen,
Germany). Total protein content was determined by means
of the DC protein assay kit (Bio-Rad, Munich).
RNA Extraction and mRNA Measurement
Standard RNA/DNA techniques were carried out according to the standard methods described in Sambrook et
al. (1989). RNA was isolated from aliquots of 107 frozen
cells harvested at different stages of SC accumulation using
the TriReagent (Sigma, St. Louis) according to the manu-
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Regulation of Phytoene Desaturase in Haematococcus pluvialis
facturer’s instructions. The concentration of total RNA was
determined spectrophotometrically at 260 nm, and aliquots
of the extracts were subjected to agarose gel electrophoresis
to test for ethidium bromide staining intensity. The results
of both methods were found to be identical. Reverse transcription followed by PCR (RT-PCR) was performed from
50 ng of RNA in a reaction buffer containing 1⫻ first-strand
buffer (GIBCO-BRL, Karlsruhe, Germany), 10 mm dithiothreitol (DTT), 1 mm dNTPs, 0.5 ␮m oligo-dT primer, 40
units of rRNasin (Promega, Madison, WI), and 200 units of
Superscript II (GIBCO-BRL) in a total volume of 40 ␮L at
42°C for 45 min after 10 min at room temperature. The
reaction was stopped by incubating the samples at 95°C for
5 min.
The PCR amplification was performed from 10 ␮L of the
RT reaction mixture in a total volume of 50 ␮L with 0.03 ␮m
of each primer, 2.5 ␮Ci of 32P-dCTP, 1⫻ Taq buffer, and 5
units of Taq polymerase. Primers were 5⬘-TCGCATCGGCCTGCTGC-3⬘ and 5⬘-GGCCAGGTGCTTGACGCT-3⬘
for yielding a 370-bp fragment for pds. The primers 5⬘-GCTGGTGAAGAGCCTGAC-3⬘ and 5⬘-TGGACCAGCTGCACTGGC-3⬘ were used for crtO, generating a 530-bp fragment. The PCR program was: 1 min at 94°C, 1 min at 58°C,
and 1 min at 72°C for 23 cycles. The PCR products were
analyzed on 7% (w/v) polyacrylamide gels in 1⫻ Trisborate/EDTA (TBE) buffer. Dried gels were exposed to
film and quantified using a phosphor imager and Macbas
software (Fuji, Tokyo). To demonstrate linearity between
the template mRNA and the final PCR product, different
amounts of RNA were used.
Expression of H. pluvialis PDS in Escherichia coli and
Enzyme Purification
PDS of H. pluvialis (GenBank accession no. X86783) was
expressed in E. coli using the QIAexpress pQE 32 vector
(Qiagen, Hilden, Germany) possessing a His-tag sequence.
Plasmid ppdsHp was digested with BamHI-KpnI and the
excised 2,137-bp fragment was ligated into the BamHI-KpnI
1263
site of the vector to yield plasmid pQE32-pdsHp. Correct
ligation was confirmed by sequencing across the ligation
sites using the QIAexpress primers (Qiagen). The plasmid
was transfected into E. coli M15 cells carrying the lac repressor plasmid pREP4. Cells from a starter culture were
grown in 2⫻ YT medium at 37°C to an OD600 of 0.6. For
induction, isopropyl-␤-d-thiogalactopyranoside (IPTG)
was added to a final concentration of 2 mm.
After 5 h of cultivation at 30°C, the cells were harvested
by centrifugation at 4,000g for 20 min. Following cell lysis
by sonication in binding buffer (0.1 m NaH2PO4 and 0.02 m
Tris-HCl, pH 8.0), the homogenate was adjusted to 6 m
guanidine-HCl in binding buffer, kept on ice for 1 h, and
centrifuged at 15,000g for 30 min. The supernatant was
loaded onto a Ni-NTA agarose column (Qiagen). After
several washes, the protein was eluted with 50 mm imidazole. Aliquots of the fractions were analyzed for PDS by
SDS-PAGE. Positive fractions were pooled and loaded onto
a preparative SDS-PAGE. Gels were stained with Coomassie Brilliant Blue R250. The corresponding band of about 66
kD, verified by western blotting using a RGSHis antibody
(Qiagen), was excised from the gel and eluted using Elucon
(Biometra). The eluate was precipitated twice with ethanol,
re-dissolved in water, dialyzed against 1 mm PBS, and
stored lyophilized.
Antibody Preparation
Polyclonal antibodies against PDS overexpressed in E.
coli and purified as described above were raised in rabbits.
Immunization was as described in Eckert et al. (1996). The
raw serum was used without further purification.
Electron Microscopy and Immunolocalization
For immunogold labeling, algal cells were harvested at
550g for 3 min, and then fixed with 4% (w/v) paraformaldehyde and 0.5% (w/v) glutaraldehyde in growth medium
for 25 min. After several washes in distilled water, the
Figure 2. Induction of PDS during the synthesis of secondary carotenoids in H. pluvialis. A, Typical nitrocellulose blot
probed with PDS antiserum (1:6,000). From left: 0.05 ␮g of the antigen (A); marker (M); total protein extracts from 105
flagellates, before (0), 2, 4, or 7 d after the onset of inductive conditions for secondary carotenoid biosynthesis. B,
Densitometric results for the induction of PDS on a cell base (f) and the drop of total protein content per cell (䡺). Bars
indicate SE of 17, 15, 2, 17, and 17 experiments for 0, 2, 3, 4, and 7 d after onset of induction, respectively, and of four
experiments for total protein content.
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1264
Grünewald et al.
Plant Physiol. Vol. 122, 2000
microscope (model EM 900, Carl Zeiss, Oberkochen, Germany) at 80 kV.
RESULTS
Overexpression of PDS in E. coli and Antibody Generation
Antibodies directed against selected enzymes are indispensable tools for expression pattern and compartmentation studies in cell fractions and by cytoimmunological
techniques. We generated antibodies against His-tagged
PDS of H. pluvialis that was overexpressed in E. coli and
purified subsequently by different steps including Niaffinity chromatography. The plasmid ppdsHp, consisting
of the full-length cDNA of H. pluvialis PDS (accession no.
X86783) cloned in EcoRI-XhoI of pBluescript SK(⫹) (Stratagene, La Jolla, CA), was kindly provided by Tamar Lotan
(Department of Genetics, Hebrew University of Jerusalem,
Israel). The coding sequence of pds was expressed in E. coli,
as described in “Materials and Methods.” Protein extracts
from IPTG-induced E. coli cells were analyzed by SDSPAGE. A prominent polypeptide of about 66 kD, which
also showed immunoreactivity with an anti-His-tag antibody, appeared. After Ni-affinity chromatography, this
protein was further purified on a 12% (w/v) SDS-PAGE.
The protein from the excised gel band was electroeluted,
dialyzed, mixed with Freund’s complete adjuvants, and
used for immunization of rabbits.
Abundance of PDS during SC Accumulation
Figure 3. Changes in the mRNA content of H. pluvialis PDS and
CRTO during secondary carotenoid biosynthesis measured by RTPCR. A, Typical autoradiograph of the RT-PCR products. The relative
amount of RNA indicated as rRNA on an ethidium bromide-stained
agarose gel is shown below. Lanes from left: M, PCR from pds in
pBluescript (used as a marker); lanes 2 to 4, RT-PCR from 16.7, 50,
and 150 ng total RNA of the extract 2 d after onset of inductive
conditions; lanes 5 to 13, RT-PCR products from 50 ng total RNA for
the different days after induction start; M, PCR from crtO in pBluescript (used as marker); lane 15, PCR from 50 ng total RNA as used
in lane 2 (RT reaction omitted). B, Results from the phosphor imager
quantification analyses of the RTR-PCR products for crtO (f) and pds
(䡺). Bars indicate the SE of four parallels.
specimens were dehydrated in a graded ethanol series with
3% (w/v) uranylacetate at 70% (v/v) ethanol for 10 min.
Cells were embedded in LR White or LR Gold (London
Resin, London) according to the manufacturer’s instructions. Ultrathin sections were cut with a microtome (Ultrotome III, LKB, Stockholm) using a diamond knife and
placed on Ni grids. Immunogold labeling of LR Goldembedded sections was performed by floating the grids
section side down on the following aqueous solutions:
blocking on 5% (w/v) BSA in PBS for 1 h; primary antibody
at 1:500 (if not otherwise indicated in the figures) in 1%
(w/v) BSA in PBS for 10 h; five washes on 1% (w/v) BSA
in PBS; 20 nm of gold-conjugated anti-rabbit-IgG 1:100
in 1% (w/v) BSA in PBS for 1 h; and five washes on
distilled water. Sections were post-stained with 3% (w/v)
aqueous uranylacetate for 5 min, 1% (w/v) aqueous lead
citrate for 30 s, and were then examined in an electron
Accumulation of SC under stress conditions leads to a
dramatic color shift of the flagellates from green to red. To
investigate whether this process is accompanied by a parallel increase in enzyme we quantified the abundance of
PDS by western analysis. Carotenogenic enzymes show
high turnover rates, and their cellular protein levels are
very low (Sandmann, 1997). The antibodies raised against
the recombinant His-tagged PDS recognized traces of the
antigen at very high dilution (Fig. 2A, lane A). We separated total protein extracts of start samples and of samples
drawn 2, 4, and 7 d after the onset of conditions inductive
for SC biosynthesis on SDS-PAGE and detected the amount
of PDS protein immunologically (Fig. 2A, lanes 0, 2, 4, and
7). The antiserum recognized a single band at about 53 kD,
representing the mature enzyme cleaved of its N-terminal
transit sequence (Sandmann, 1994). This band appeared in
all stages and showed typical induction kinetics in the time
course of SC biosynthesis, being constitutively abundant in
the green flagellates, reaching a maximum after about 3 to
4 d after the start of induction, and declining notably
during the later SC synthesis period (Fig. 2B). The preimmune serum did not give any signal in the immunological
analysis whether in the total extracts or with the antigen
(data not shown).
Transcript Levels of PDS and CRTO during
SC Accumulation
In addition to data on protein abundance, for further
examination of the regulation of carotenoid genes during
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Regulation of Phytoene Desaturase in Haematococcus pluvialis
1265
Figure 4. Immunogold labeling of PDS in a flagellate of H. pluvialis exposed for 2 d to SC-inducing conditions. A, Typical
part of a flagellate showing all major cell compartments. Bar represents 1 ␮m. B, Magnification of the chloroplast. Bar
represents 0.2 ␮m. Ch, Chloroplast; Ex, extracellular matrix; Li, lipid vesicles; Nc, nucleolus; Nu, nucleus; St, starch; Va,
vacuole.
idly. The maximal amount after 4 d matched the maximum
observed for the PDS, and the subsequent decrease was
comparable to that observed in PDS.
induction of SC synthesis in H. pluvialis, the changes of
mRNA levels were investigated. Due to very low abundance of transcripts of carotenogenic genes (Cunningham
and Gantt, 1998), the sensitive method of RT-PCR was used
for quantification. We measured the steady-state levels of
mRNA for PDS and CRTO by RT-PCR of total RNA. During the course of exposure, changes in total RNA per cell
content did not exceed 10%. Co-amplification RT-PCR of
the two transcripts was performed using four specific
primers. As demonstrated for the RNA isolated from samples drawn 2 d after the onset of SC induction, the amount
of amplification product was linearly correlated with the
initial amount of total RNA used for the RT reaction (Fig.
3A, lanes 2–4). No amplification product appeared if the RT
reaction prior to PCR was omitted (Fig. 3A, last lane). The
steady-state PDS mRNA level was found to be linearly
increased from the preexisting basal level immediately
upon onset of inductive conditions, reaching its maximum
after 4 d, and then decreasing slowly. In the case of CRTO,
no mRNA was detected in the green flagellates of the start
samples. After a delay of about 1.5 d of incubation the
steady-state level of the CRTO transcripts increased rap-
Immunogold Localization of PDS
We found that the best structural preservation and maintenance of antigenic structures was achieved when cells
were fixed at increasing concentrations of paraformaldehyde and glutaraldehyde and embedded after dehydration
in LR Gold resin. Accessibility of antigenic determinants
was proven for membrane-bound antigens by probing with
a anti-light-harvesting protein (LHC) antibody raised
against spinach LHC, and for soluble proteins with the
anti-Rubisco small subunit (data not shown). Both antibodies were kindly provided by U. Johanningmeier (Institute for Plant and Cell Physiology, University of Halle,
Germany).
Probing the sections with the polyclonal antibodies
raised against PDS resulted in a typical pattern of gold
decoration (Fig. 4A; Table I). Two main parts of the cells
were specifically labeled, the chloroplast and the outer
Table I. Distribution of PDS immunogold labeling in the major compartments of the flagellates at different stages of SC biosynthesis
Time after Onset of
Induction
Primary Antibody
Relative Immunogold Labeling
Chloroplast
Extracellular matrix
39.9 ⫾ 2.2
42.7 ⫾ 4.0
50.1 ⫾ 4.0
24.2 ⫾ 1.4
56.3 ⫾ 2.9
51.9 ⫾ 1.7
47.7 ⫾ 4.0
41.1 ⫾ 3.7
68.9 ⫾ 1.8
17.2 ⫾ 4.1
d
0
2
4
7
2
a
Cytoplasma
Nucleus
Cells, Labeling
Counted for
Mean of Gold
Particles per Cell
6.0 ⫾ 0.6
8.2 ⫾ 1.7
6.7 ⫾ 0.7
6.5 ⫾ 0.7
20.3 ⫾ 1.7
2.2 ⫾ 0.7
1.3 ⫾ 0.5
2.1 ⫾ 0.5
0.5 ⫾ 0.2
6.2 ⫾ 2.1
10
11
12
11
11
69
51
66
113
22
%
Anti-PDS 1⬊1,000
Anti-PDS 1⬊1,000
Anti-PDS 1⬊1,000
Anti-PDS 1⬊1,000
Preimmune 1⬊250
Includes lipid vacuoles.
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1266
Grünewald et al.
Plant Physiol. Vol. 122, 2000
Figure 5. Distribution of PDS in cellular fractions of H. pluvialis flagellates exposed for 4 d to
N deprivation and high light. A, Coomassiestained SDS-PAGE gel of chloroplastic (1), supernatant (2), microsomal (3), and lipid vesicle
fraction proteins (4). Each lane was loaded with
15 ␮g of protein. B, Western-blot analysis with
antibodies raised against recombinant PDS (1:
4,000). Lanes 1 to 4 are as indicated in A, but
were loaded with cell fraction aliquots of 105
cells.
paracrystalline layer of the extracellular matrix (Siegmund,
1999). No specific signal was obtained in or around the
lipid vesicles or in the cytoplasm. Statistics of the distribution of the gold particles in the course of SC biosynthesis
are presented in Table I. The signal in the extracellular
matrix was evident at all stages and is interpreted as crossreactivity due to a antigenic determinant occurring in PDS
and a structural protein of the extracellular matrix. Interestingly, western-blot analysis did not reveal any crossreactivity of such proteins. No specific labeling was observed
when sections were probed with preimmune serum (Table
I). At higher magnifications, the chloroplast labeling
proved to be specific to the thylakoids (Fig. 4B).
Compartmentation of PDS
To prove the results from the immunogold localization
experiments, we probed cell fractions with the anti-PDS
antibodies. Following a gentle cell disruption by passing
flagellates through polycarbonate isopore filters, fractions
were isolated by differential centrifugation and phase separation. Four fractions were obtained (Fig. 5A, lanes 1–4).
Light-microscopic observations identified them as: (a) a
pellet of starch granules, cell debris, and larger cell organelles, mainly of the chloroplast; (b) a supernatant fraction; (c) a pellet of microsomes and cytoplasmic membranes; and (d) the lipid vesicle fraction. When probed with
PDS antibodies, an immunoreactive band occurred only in
the chloroplast membrane fraction (Fig. 5B). The results
shown here are for fractions of flagellates incubated for 4 d
under inductive conditions for SC synthesis, representing a
state at which the amount of PDS in total extracts was
elevated compared with the values before the onset of
induction.
DISCUSSION
In the present study we addressed the question of
whether the biosynthesis of secondary carotenoids in H.
pluvialis proceeds via an independent second pathway that
operates outside the chloroplast, as the accumulation of the
astaxanthin esters in cytosolic lipid vesicles might indicate.
According to our data for PDS, we conclude that, at least
for this relatively early biosynthetic step of carotenogenesis, no second cytosolic pathway exists and that higher
biosynthetic activity is coupled to higher amounts of enzyme. Up-regulation on both the mRNA and protein levels
was observed upon induction of SC synthesis. Because of
slight variability in the extent of SC accumulation between
parallels (Grünewald, 1997), we do not interpret the small
difference between the maxima in PDS mRNA and protein
levels as a sign for post-translational regulation events—at
least the main part of up-regulation takes place at the
mRNA level.
Bouvier et al. (1998) showed that pepper PDS mRNA
increased under different stress conditions. During accumulation of capsanthin and capsorubin, Römer et al. (1993)
observed after an increase between the mature green pepper fruit and the first defined ripening stage a constant
level of PDS mRNA despite the massive increase of carotenoids occurring during the further ripening stages, and
this resembles our findings here. Although SC continued to
accumulate, PDS mRNA did not increase further and,
moreover, even slightly decreased. Despite the delay in the
increase and the absence in green, unstressed flagellates, an
analogous pattern was observed for CRTO mRNA. In respect to the absence in green flagellates, our results are
different from those reported by Sun et al. (1998), who
detected mRNA of CRTO even in noninduced green cells.
This might have been caused by different cultivation conditions used. Especially, the presence of acetate in the
medium before induction could be responsible for the basic
level of CRTO mRNA reported by these authors. Unfortunately, at the present time, antibodies directed specifically
against the ␤-carotene oxygenase or other enzymes involved in the late steps of SC synthesis are not available.
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Regulation of Phytoene Desaturase in Haematococcus pluvialis
The increase in the transcripts of carotenogenic genes, as
shown here for ␤-carotene oxygenase and PDS, and the
parallel increase in the protein level, observed at least for
PDS, obviously enable the alga to maximize SC synthesis
under stress conditions. Support for this point of view
comes from Linden (1999), who cloned ␤-carotene hydroxylase from H. pluvialis and observed an increase of the
corresponding mRNA in parallel to astaxanthin synthesis.
Contrary findings have been reported in Dunaliella bardawil
(Rabbani et al., 1998). On stress induction, this alga massively accumulates ␤-carotene in intraplastidic droplets.
No significant parallel increases in PDS protein or mRNA
levels were observed. One reason for this striking difference between the regulation patterns of carotenogenesis in
these two Volvocales might be the different localization of
the final product. The intraplastidic accumulation in D.
bardawil seems to be regulated by the presence of the
sequestering structures (Rabbani et al., 1998). In contrast,
the extraplastidic accumulation in H. pluvialis might allow
the common transcriptional control of biosynthesis of SC
and cytoplasmic lipid vesicle material. This reflects the
importance of the localization of the processes involved in
terms of regulation.
Only very few immunogold localization experiments
have been done for carotenogenic enzymes, and most have
dealt with isolated and enriched organelles. Typical examples are geranylgeranylpyrophosphate synthase in pepper
(Cheniclet et al., 1992) and PDS in isolated spinach chloroplasts (Linden et al., 1993). We present, for the first time to
our knowledge, immunogold labeling data on the localization of PDS in algal cells. Most of the labeling was found
within the chloroplast. Moreover, the absence of specific
signals in the cytoplasm or inside or around the SCcontaining lipid vesicles ruled out the possible occurrence
of PDS there. Higher magnifications of chloroplast regions
showed the signal predominantly located in close contact
to the thylakoids. The non-thylakoidal-localized gold particles might indicate a soluble inactive population of PDS in
the stroma (Al-Babili et al., 1996).
Our results suggest that PDS isoenzymes, possibly located in the cytoplasm, are not involved in the synthesis of
SC (for IPP isomerase, compare with Sun et al., 1998). From
the highly conserved structure of plant type PDS (Sandmann, 1994), and the polyclonal nature of the antibodies,
we conclude that isoenzymes of PDS would have been
recognized in our cytoimmunochemical experiments.
Moreover, changes in the enzyme amount were in accordance with SC synthesis and the mRNA pattern.
Cell fractionation experiments supported results from
immunogold cytochemistry. Fractionation of cells with increased PDS amounts resulted in four fractions, of which
only the chloroplast membranes gave a signal in westernblot analysis. This confirms previous data on PDS localization in different plants and sequence information of a
chloroplast import signal that is found in all PDS (Bonk et
al., 1997).
We conclude that PDS in H. pluvialis is restricted to the
chloroplast. We also demonstrated that the regulation of
PDS occurs primarily at the mRNA level, most likely by
transcriptional control, and that the enzyme is induced in
1267
response to stress conditions leading to SC biosynthesis.
The localization results implicate for H. pluvialis a transport
process of SC over compartment borders from the chloroplast as the site of synthesis to the lipid vesicles located in
the cytoplasm as the site of accumulation. Further elucidation of this process and the intermediate step at which it
occurs will help to complete our knowledge on the regulation of SC biosynthesis.
ACKNOWLEDGMENTS
We thank Prof. Dr. W. Braune who has suggested and encouraged us to work on the H. pluvialis carotenoid problem. We are
indebted also to S. Schmidt for technical assistance and to M.
Goldstein, V. Mann, M. Utting, and J. Müller for helpful comments
during the course of this work, and especially to U. Johanningmeier (Institute for Plant and Cell Physiology, University of Halle,
Germany) for kindly providing the antispinach LHC and the antiRubisco small subunit antibodies.
Received July 27, 1999; accepted December 27, 1999.
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