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ON THE EVOLUTION AND ECOLOGY OF THE GREEN
DINOFLAGELLATE GYMNODINIUM CHLOROPHORUM1
Daniel Grzebyk2, 3
Environmental Biophysics and Molecular Ecology Program, Institute of Marine and Coastal Sciences, Rutgers, The State University of New Jersey, 71 Dudley Road, New Brunswick, New
Jersey 08901, USA.
Robert R. Bidigare
Department of Oceanography, University of Hawai`i at Manoa, 1680 East-West Road POST 105, Honolulu, Hawai`i 96822, USA
Yibu Chen,
Environmental Biophysics and Molecular Ecology Program, Institute of Marine and Coastal Sciences, Rutgers, The State University of New Jersey, 71 Dudley Road, New Brunswick, New
Jersey 08901, USA.
Costantino Vetriani
Institute of Marine and Coastal Sciences and Department of Biochemistry and Microbiology, Rutgers, The State University of New Jersey, 71 Dudley Road, New Brunswick, New Jersey
08901, USA
Paul G. Falkowski
Environmental Biophysics and Molecular Ecology Program, Institute of Marine and Coastal Sciences, Rutgers, The State University of New Jersey, 71 Dudley Road, New Brunswick, New Jersey 08901, USA and Department of Geological Sciences, Rutgers, The State University of
New Jersey, Piscataway, New Jersey 08854, USA.
1 Received . Accepted .2 Author for correspondencecorrespondance: e-mail [email protected]? Current address: Muséum National d’Histoire Naturelle, USM0505, Systématique et Ecotoxicologie des Microalgues, 12 rue Buffon, Case 39, 75231 Paris Cedex 053
1
Abstract
We examined the molecular evolution and ecophysiological niche of the green dinoflagellate,
Gymnodinium chlorophorum. There is a single 18S rDNA in G. chlorophorum, suggesting that
the organism contains true plastids but not a nucleomorph. The 18S rDNA sequence from this
organism G. chlorophorum is identical to that of the only other green dinoflagellate cultured to
date, Lepidodinium viride, suggesting that the two species are derived from an identical host cell
and diverged a very short time ago. Using a PCR approach, followed by RFLP and sequencing,
no other 18S rDNA that would reveal a putative nucleomorph genome could be detected. Based
on Using pulsed-field gel electrophoresis and Southern blot analyses, the size of the plastid
genome was determined to be ~100 kb, which is among the smallest plastomes characterized to
date. Immunoblotting and gene sequencing indicates that G. chlorophorum contains the green
form 1B of RuBisCO., as determined using immunoblotting detection and verified via gene
sequencing. Phylogenetic analyses of plastid encoded rbcL and psbA genes indicates that the
plastid was not derived from a prasinophyte, but, more likely, was obtained from modern
chlorophytes, either the Chlorophyceae or the Trebouxiophyceae. This hypothesis is supported by
analysis of the HPLC analysis of-determined pigment composition; G. chlorophorum does not
contain any of the pigments commonly reported for prasinophytes (e.g., MgDVP and
prasinoxanthin), but the pigment profile is similar to modern chlorophytesChlorophyceae or
Trebouxiophyceae. However, the strain we analyzed (DIN3) lacks the photoprotective pigment,
lutein. Maximum growth rate was estimated to be 0.9-1.0 div•d-1. Optimum growth conditions
were 17-19° C, with a very narrow irradiance optimum of ~150-200 µmol quantaE•m-2•s-1. Based
on fast repetition rate fluorometry of photosynthetic parameters and cell-specific photosynthetic
pigment concentration, it would appear that this dinoflagellate has a very limited ability to
2
photoacclimatee. This lack of physiological plasticity appears to markedly reduce the niche
width of this species. The biochemical impact of acquiring its green plastid and the physiological
characteristics of G. chlorophorum isare discussed in an ecological perspective, with respect to
the capacity of this species to bloom in coastal environments.
Key index words: physiological ecology, plastid endosymbiosis, rDNA
Running title: GYMNODINIUM CHLOROPHORUM EVOLUTION AND ECOLOGY
3
Introduction
In a currently accepted view of eukaryotic phytoplankton evolution, the chlorophyll c2 and
peridinin-containing plastids in the ecologically dominant type of dinoflagellates were acquired
from a red alga by secondary endosymbiosis (Delwiche 1999, Durnford et al. 1999, Takishita and
Uchida 1999, Harper and Keeling 2003). However, a small number of dinoflagellate species have
unique plastid morphologies and photosynthetic pigmentation (Schnepf and Elbrächter 1999).
Like diatoms and haptophytes, some Some species of dinoflagellates contain fucoxanthin and/or
its derivatives, which are also found in, like haptophytes and diatoms, from which these plastids
in the dinoflagellates were have been acquired by tertiary endosymbiosis (Chesnick et al. 1996,
Tengs et al. 2000). Other species have plastids containing phycoerythrin and a thylakoid
structure, and appear to similar to that found inhave been acquired from cryptophytes, likely also
by tertiary endosymbiosis (Takishita et al. 2002, Hackett et al. 2003). Regardless of the
subsequent modifications, aAll these plastid types belong to the red algal plastid lineage derived
fromoriginating from rhodophytes (Grzebyk et al. 2003). Gymnodiniales is an Order that contains
the highest plastid diversity, either as permanent organelles or as temporarily acquired plastids
(kleptochloroplasts) from prey microalgae (Schnepf and Elbrächter 1999). Interestingly, Aa few
Gymnodiniales species contain have green plastids withthat contain chlorophyll b as the major
accessory pigment. With respect to the position of gymnodinioid species with green or
fucoxanthin-derivative containing plastids in dinoflagellate/gymnodinioid molecular phylogenies
(Saunders et al. 1997, Grzebyk et al. 1998, Hansen et al. 2000), i It is assumed that these green
dinoflagellates are species were derived from taxa that previously had peridinin-containing
plastids, which werewere then lost and subsequently replaced with the current set of plastids
(Saldarriaga et al. 2001). If so, they should contain a nucleomorph.
4
Two species of green marine dinoflagellates, Gymnodinium chlorophorum (Elbrächter
and Schnepf 1996) and Lepidodinium viride (Watanabe et al. 1987, Watanabe et al. 1990), have
been described extensively in terms of ultrastructure. In both species, the chloroplasts are thought
to have originated from a prasinophyte alga, and therefore, would be the result of a serial
secondary endosymbioticiosis process. however, no No currently available molecular dataa,
however, are currently available to support this hypothosishypothesisassumption.
Green dinoflagellates have been known for a long time (Biecheler 1939, 1952). However,
reports of these organisms in marine phytoplankton communities were rare until the early 1980’s.
The first observation of a “green tide” by a gymnodiniale species, later identified as
Gymnodinium chlorophorum, was observed in summer of the Summer of 1982 in a Brittany
estuary, France (Sournia et al. 1982). Since then, blooms have been increasingly reported during
the sSummer on the Atlantic coast between the Gironde and Seine estuaries by the French
phytoplankton monitoring network, REPHY of IFREMER
(http://www.ifremer.fr/envlit/actualite/20011012.htm). G. chlorophorum blooms have also been
reported in the North Sea and Kattegat (Elbrächter and Schnepf 1996, Mouritsen and Richardson
2003) and possibly in Chile (Iriarte et al. 2005). Although not toxic, these blooms are locally
responsible for death of marine fauna as a result of generating anoxic conditions. There are only a
few cultured strains available and none of them has have been the subject of an ecophysiological
study.
In this paper, we present an extensive investigation of the species G. chlorophorum.
Ribosomal DNA sequences were used to determine the phylogenetic relationship of this species
in comparison with the only other cultured green dinoflagellate, L. viride. Biochemical and
molecular data obtained from the chloroplast were used to infer the origin of this organelle within
green algae. In order to place the phylogenetic data in an ecological framework, an
5
ecophysiological study was conducted to investigate the photo-physiology of this alga, in terms
of growth rate, photosynthetic parameters and pigment content, to environmental variations in
temperature and irradiance.
Material and methods
Algal cultures. The G. chlorophorum strain DIN3, deposited in the Algobank collection
(Laboratoire de Biologie et Biotechnologies Marines, Université de Caen, Caen, France), was
isolated from Luc-sur-Mer (Normandy, France) in 1995 by Jacqueline Fresnel. Cells were grown
in f/2 medium (Guillard and Ryther 1962) at a salinity of 31 (pratical practical salinity scale) and
at an irradiance of ~150 µmol quantaE•m-2•s-1 provided by fluorescent tubes on a 12/12 h
light/dark cycle. For pigment analysis and molecular biological studies, cells were harvested
either by continuous centrifugation or filtration onto Whatman GF/F glass fiber filters, and stored
at -70° C prior to analysis. For comparative nucleotide sequencing (nuclear rRNA and plastid
psbA genes), the G. chlorophorum type strain BAH ME 100 (Elbrächter and Schnepf 1996) was
obtained from Dr. Malte Elbrächter as a pelleted cell sample preserved in 95% ethanol with 100
µM EDTA at pH 8.0.
Electron microscopy. For transmission electron microscopy, G. chlorophorum cells were
centrifuged and processed through two fixation protocols, FIX. A and FIX. B, as used previously
for the description of the species (Elbrächter and Schnepf 1996).
RuBisCO detection by western blot analysis. For western blotting analyses, protein extracts
were prepared by the addition of one volume of 4% SDS, 0.1 M Na2CO3 to one cell pellet
volume, sonicated on ice, diluted with storage buffer (4% SDS, 15% glycerol, 0.05%
bromothymol blue, 0.05 volume 100 mM PMSF, 0.10 volume 1 M DTT), and boiled for 2 min
before flash freezing in liquid N2 and stored at –70° C. Proteins were separated on 12%
polyacrylamide gels and transferred onto a polyvinylidene fluoride (PVDF) membrane. Proteins
6
were challenged with two polyclonal antisera generated, for the first one, against the green type
RuBisCO (Form 1B) holo-enzyme from tobacco (kindly provided by Steve Mayfield, The
Scripps Research Institute, La Jolla, CA, USA) and, for the second one, against the red type
RuBisCO (Form ID) from Isochrysis galbana (Falkowski et al. 1989). A secondary antibody,
Affi-pure goat anti-rabbit HRP (Bio-Rad Laboratories, Hercules, CA, USA), and the SuperSignal
chemiluminescence reagent (Pierce, Rockford, IL, USA) were used for detection.
Pigment analyses. Pigment analyses were performed using the HPLC method of Bidigare et
al. (Bidigare et al. 2004). The chlorophyte Dunaliella tertiolecta (CCMP1320) and the
prasinophyte Pycnococcus provasolii (CCMP1203), from the Provasoli-Guillard National Center
for Culture of Marine Phytoplankton (Bigelow Laboratory, Boothbay Harbor, ME, USA), were
used as reference materials for the analysis of green algal photosynthetic pigments. For
extraction, the filters were placed in 3 mL acetone and ground using a glass/glass homogenizer.
The samples were then allowed to extract for 5 hr (4ºC, in the dark). Prior to analysis, the
pigment extracts were vortexed and centrifuged to remove cellular and filter debris. Samples
(200 L) of a mixture of 0.3 mL H2O plus 1.0 mL extract were injected onto a Varian 9012
HPLC system (Varian, Palo Alto, CA, USA) equipped with a Varian 9300 autosampler, a
Timberline column heater (26ºC), and Spherisorb 5 m ODS2 analytical (4.6250 mm) column
and corresponding guard cartridge. Pigments were detected with a ThermoSeparation UV2000
detector ( = 436 nm). A ternary solvent system was employed for HPLC pigment analysis:
eluent A (MeOH:0.5 M ammonium acetate, 80:20), eluent B (acetonitrile:water, 87.5:12.5) and
eluent C (ethyl acetate). The linear gradient used for pigment separation was a modified version
of that originally described by Wright et al. (Wright et al. 1991): 0.0’ (90%A, 10%B), 1.0’
(100%B), 11.0’ (78%B, 22%C), 27.5’ (10%B, 90%C), 29.0’ (100%B), and 30.0’ (100%B).
Eluents A and B contain 0.01% of 2,6-di-tert-butyl-p-cresol (BHT) (Sigma, St. Louis, MO,
7
USA.) to prevent the conversion of chlorophyll a into chlorophyll a allomers. HPLC grade
solvents (Fisher Scientific, Pittsburgh, PA, USA) were used to prepare eluents A, B and C. The
eluent flow rate was held constant at 1 mL min-1. Eluting peaks were identified by retention
time comparisons with standards and the reference algal extracts. Pigment identifications were
confirmed by online diode array spectroscopy. Pigment quantification was performed using
external standards with the exception of neoxanthin and violaxanthin that were estimated with the
violaxanthin response factor.
Nucleotide sequencing and phylogenetic analyses. For DNA extraction, cells were
resuspended in a lysis buffer (1.2% SDS, 30 mM EDTA, 50 mM Tris-HCl, 220 mM NaCl, 50
mM -mercaptoethanol) for 15 min at room temperature and then extracted using a buffered
phenol/chloroform procedure. Selected gene fragments were amplified through 30-35 PCR
cycles, using the cloning primers and conditions given in Table 1. PCR products were resolved
on a 1% agarose electrophoresis gel, and the DNA purified from the gel using the QIAquick Gel
Extraction Kit (Qiagen, Valencia, CA, USA), then cloned into either the pCR 2.1 or the pCR 4-
TOPO vector using the Topo TA Cloning kit (Invitrogen, Carlsbad, CA, USA) before E. coli
transformation. Plasmids were purified from select clones using the QIAprep Spin Miniprep Kit
(Qiagen). Terminator cycle sequencing reactions (BigDye version 3.0 or 3.1, Applied
Biosystems, Foster City, CA, USA) were carried out using the M13 reverse and forward primers,
and the appropriate internal sequencing primers (Table 1). Sequencing was performed using an
ABI PRISM 3100-Avant Genetic Analyzer (Applied Biosystems). Sequence data were assembled
using ContigExpress from the Vector NTI Suite 7 software package (Invitrogen, Carlsbad, CA,
USA). The assembled sequence (3,458 nucleotides) of the rDNA from G. chlorophorum DIN3
was deposited in GenBank under the accession number AY331681; it includes the 18S, 5.8S and
partial 28S (D1-D3 region) rRNA genes and the internal transcribed spacers ITS1 and ITS2.
8
Nucleotide sequence alignments, with data retrieved from Genbank, were performed using
ClustalX (Thompson et al. 1997) and refined by eye. Phylogenetic analyses were carried out
using a variety of programs and methods for comparison. Maximum likelihood (ML) analyses
were carried out using the fastDNAml (version 1.2.2) (Olsen et al. 1994) program available
online (http://bioweb.pasteur.fr/seqanal/phylogeny/intro-uk.html), using the default setting but
with the weighting option set to limit the analysis to the first and second codon positions.
Bootstrapped ML analyses were performed using the program PHYML v2.4.3, according to the
general time reversible (GTR) substitution model and applying a “invariant + ” model for the
among-site substitution rate distribution approximated with 8 categories of substitution rates,
allowing the program to estimate all analytical parameters (proportion of invariant sites,
transition/transversion ratio, -shape parameter ) (Guindon and Gascuel 2003). One hundred
(rbcL) to one thousand (psbA) non-parametric bootstrap analyses were performed and the
consensus tree was assembled using the program CONSENSE from the PHYLIP package
(Felsenstein 1993). A Bayesian likelihood inference of phylogeny was performed using the
program MrBayes, version 3.0B4 (Ronquist and Huelsenbeck 2003), using the substitution and
an among-site substitution rate distribution model, as before. One million MCMC cycles were
computed with trees sampled every 500 generations. The consensus tree was calculated after
burning the first 100 sampled trees.
Screening for endosymbiotic 18S rDNA genes. Our approach combined the use of PCR and
DNA restriction methods in an attempt to favor the amplification and the detection of a putative
endosymbiont nucleomorph (sensu Elbrächter and Schnepf, 1996) 18S rRNA gene. Nine
restriction enzymes, each producing a single cut of the G. chlorophorum 18S rDNA sequence,
were selected using the NEBcutter2 tool (http://tools.neb.com/NEBcutter2/index.php, New
England Biolabs, Beverley, MA, USA): HindIII, BamHI, BstXI, EcoRI, SwaI, ClaI, RsrII, BcgI
9
and PpuMI. Restriction The enormous dinoflagellate nucleus contains a great number of 18S
rDNA copies that could outnumber those from a putative endosymbiont nucleomorph. Hence,
PCR-RFLP analysis might only reveal fragmentation patterns corresponding to those predicted
from the dinoflagellate sequence. We made the assumption that if a nucleomorph 18S rRNA gene
is present, in order to decrease the number of dinoflagellate rRNA gene copies, digestion of G.
chlorophorum genomic DNA with the selected restriction enzymes prior to PCR amplification
should increase the probability of amplification of the nucleomorph 18S rRNA gene. If
Following the PCR, in anthe RFLP analysisnucleomorph rDNA was present, if at least one
selected restriction enzyme can also cut the nucleomorph gene, restriction of 18S rDNA PCR
products should reveal two distinctive patterns of DNA fragmentation compared to undigested
genomic DNA, one corresponding to the nucleomorph gene, the other corresponding to the
pattern predicted for the nuclear gene. Restrictions of genomic DNA were performed overnight
(15 h), in 20µL reactions, using 200 ng DNA and 2 enzyme units in the conditions recommended
by the manufacturers (New England Biolabs, and Promega, Madison, WI, USA). Digested
genomic DNA (20 ng) was used as a template for PCR amplification of the 18S rDNA (18S-1F
and 18S-1R primers, 0.5 pmol•µL-1 reaction), which was performed as indicated in Table 1, for
35 cycles, using JumpStart RedAccuTaq (Sigma, St. Louis, MO, USA). An aliquot of 18S rDNA
PCR products was used for the restriction reactions by the nine selected enzymes. Each PCR
product obtained from digested dinoflagellate DNA was digested with the same enzyme,
respectively, using 4 enzyme units. After visualization of restriction patterns, 1 µL of each
reaction was used as a template for reamplification of 18S rDNA (25 PCR cycles). The PCR-
amplified fragments were purified via agarose gel electrophoresis and gel extraction as described
above, then used as a template for sequencing analysis, which was performed as outlined above.
In parallel, a PCR reaction was performed with non-digested genomic DNA as a template: the
10
18S rDNA PCR products were used as size markers, and nine aliquots were digested by the
selected restriction enzymes, in order to compare the restriction patterns with those obtained
previously.
Chloroplast genome sizing. Frozen cell pellets (100-200 mg) were re-suspended in 2 mL lysis
buffer: 100 mM EDTA (pH 8.0), 20 mM Tris-HCl (pH 8.0), 1% SDS, 1% sodium lauryl
sarcosine, 1% Tween 20, 0.5% Triton X-100, DNase inhibitors (1% Polyvinylpyrrolidone PVP-
40, 10 mM EGTA, 5 mM 1,10-Phenanthroline monohydrate), 0.01 volume of polyamine solution
(from a 100x stock solution: 30 mM spermine and 75 mM spermidine in water, 0.2 µm-filtered
and stored at –20° C), and 1 mg•mL-1 proteinase K. Cell suspensions were incubated at 50° C
with rotation in a hybridization oven for 2-15 h. The suspensions were electro-dialyzed overnight
in 0.5xTBE buffer at 4.5 V•cm-1 using an ElectroPrep System (Harvard Apparatus, Holliston,
MA, USA). After a brief spin to pellet undigested cells and cell debris, solutions were gently
mixed (1:1 by volume) with melted 2% InCert agarose (Cambrex, Walkersville, MD, USA) at
55° C and distributed in plug molds. Plugs were washed in the lysis buffer without proteinase K,
using 5-10 mL per mL agarose, at 50° C with gentle rotation for 1-3 times, until discoloration was
observed, then several times in the washing buffer (50 mM EDTA pH 8.0, 20 mM Tris-HCl pH
8.0) at 50° C. Pulsed field gel electrophoresis (PFGE) was run using the CHEF III DR system
(Bio-Rad Laboratories, Hercules, CA, USA) under the following conditions: 1.25% agarose gel
(Bio-Rad Laboratories), 0.5 x TBE buffer at (10.5° C), 6 V•cm-1, 15 h run time, a switch time
starting at 0.2 sec and rising up to 15 sec. DNA was visualized after staining with ethidium
bromide using a Typhoon 9410 scanner (Amersham Biosciences, Sunnyvale, CA, USA), then
transferred by Southern blot onto a positively charged Nylon membrane. Chloroplast genomes
were detected with a blend of 32P labelled psbA probes obtained from G. chlorophorum,
Trichodesmium sp. and I. galbana. Hybridization was performed overnight at 48° C. Detection
11
was performed by exposing storage phosphor screen autoradiography plates for 3-4 h and
visualization using the Typhoon 9410 scanner.
Influence of temperature and irradiance on growth rate. The influences of temperature and
irradiance on the growth rate of G. chlorophorum was determined using a custom built
aluminium block where 90 one-hundred mL culture tubes can be exposed to simultaneous
crossed gradients of temperature and light. Ten temperature conditions were set up ranging from
10° C to 30° C (10, 12, 15, 17, 19, 21.6, 23.9, 26.1, 28.1 and 30° C). The irradiance gradient was
set up using fluorescent tubes (Sylvania Dulux-L, 55 W, 4800 Lumens) and neutral density
filters; irradiance values were measured using a QSL-100 quantum meter (Biospherical
Instruments Inc., San Diego, CA, USA). Six ranges of irradiance values were selected for each
temperature: 7.5-12.7, 17-30, 40-63, 65-104, 120-226 and 379-522 µmol quantaE•m-2•s-1,
providing 60 different temperature-light conditions on a 14/10 h light/dark cycle. The experiment
was started with a growing culture, which was diluted (55 times) in filter-sterilized f/10 medium
to the initial concentration of ~200 cell•mL-1, then 50 mL aliquots were aseptically distributed
into experimental tubes. Growth was monitored daily by measuring in vivo fluorescence using a
10-AU Turner Designs fluorometer (Brand et al. 1981). Cultures were sampled for cell counts
that were performed after sedimentation in Lab-Tek Chamber Slides (Nalge Nunc International,
Rochester, NY, USA). The growth rate was calculated according to (Guillard 1973) as kmax
(div•d-1) using data taken during the exponential growth phase and over 2-day periods. Growth
response surface vs. irradiance and temperature was inferred using SigmaPlot (SPSS Inc.,
Chicago, IL, USA), followed by manual adjustment. During exponentiael growth, photosynthetic
parameters were measured using a fast repetition rate fluorometer after placing cells in darkness
for a 30 min. period (Kolber et al. 1998).
12
Influence of irradiance on pigment content. The low and medium irradiance conditions (range
20-40 and 130-170 µmol quantaE•m-2•s-1, respectively) were obtained by using regular cool white
fluorescent tubes. The high irradiance condition (range 930-1060 µmol quantaE•m-2•s-1) was
obtained by using high-output cool white fluorescent tubes (GE F48T12-CW-HO). An
exponentially growing culture (f/2 medium) was divided into three 900 mL subcultures in
polycarbonate Erlenmeyer flasks, which were then exposed to the three light treatments and on a
12/12 h light/dark cycle. The experimental temperature was set at 19° C. Cells were collected
onto Whatman GF/F glass fiber filters for pigment analysis before reaching the stationary phase.
High-light grown cells were collected after seven days in the treatment while the low- and
medium-light grown cells were collected after 10 days.
Results
TEM ultrastructure. Ultrastructural features observed in our strain are consistent with the
original description of the species G. chlorophorum (Elbrächter and Schnepf 1996, Fig. 1). No
body scales were observed on the surface of the cells or in formation inside of the cells. The cell
contains plastid structures on the periphery, with paired thylakoid lamellae that are typical of
green plastids. We could not conclusively determine the number of plastid envelope membranes
from the thin sections.
Ribosomal DNA sequences. The rDNA sequence from G. chlorophorum DIN3 is identical to the
homologous sequence obtained by direct sequencing of PCR products from the species type
strain (BAH ME 100). Moreover, the 18S rRNA gene sequence is identical to that reported from
Lepidodinium viride (accession number AF022199), except for 3 mismatching nucleotides
corresponding to the reverse primer binding site. The sequence of the 28S rDNA D1/D3 region
previously obtained from the G. chlorophorum type strain (accession number AF200669) has 3
mismatching nucleotides derived from the forward primer binding site.
13
Screening for an endosymbiotic nucleomorph genome. Digestion After the digestion of
dinoflagellate genomic DNA, of amplified the 18S rDNA PCR-RFLP analysis resulted in visible
restriction patterns identical to those obtained from non-digested genomic DNA and matching
those predicted from the gene sequence (Fig. 2). and identical to those obtained after restriction
of the 18S rDNA PCR fragments from non-digested genomic DNA. In the subsequent
sequencing Analysis analysis of sequenced fragments, following obtained from PCR
reamplification of 18S rDNA from left in the digestion reactions and in the undigested control,
the sequencing chromatograms showed the absence of genetic polymorphism and revealed only
the dinoflagellate 18S rDNA.
Pigment analysis. The profile of elution profiles of photosynthetic pigments in G.
chlorophorum, D. tertiolecta and P. provasolii are shown in Figure 3. In G. chlorophorum, the
small peak eluting before zeaxanthin was not confirmed not to be lutein in a subsequent HPLC
analysis of a concentrated pigment extract. Prasinoxanthin and MgDVP (Mg-[3,8-divinyl]-
phytoporphyrin-132-methylcarboxylate) were not detected in the extract. Relative abundances of
accessory photosynthetic pigments vs. Chl a are shown in Table 2. In G. chlorophorum, the total
carotenoid ratio vs. Chl a was 3-4 fold smaller than observed for the two other green algal
species. In the absence of lutein and prasinoxanthin, which are the major carotenoids in the two
other algae, the dinoflagellate carotenoid content was shifted towards high proportions (> 20%)
of neoxanthin and xanthophyll cycle pigments (antheraxanthin and violaxanthin). In addition,
while the neoxanthin-, antheraxanthin- and violaxanthin-to-Chl a ratios were in the same range as
measured in the other two algae, the ratio for zeaxanthin was approximately 10-20 fold lower.
When G. chlorophorum cells were exposed to three different irradiances, large changes in
pigment composition and cell-specific pigment concentrations were observed (Table 3). Cell-
specific chlorophyll and total carotenoid concentrations were the lowest under the high-light
14
treatment, and increased gradually with decreasing growth irradiance. Under the high-light
treatment, cells werecolor turned to pale yellow,; , chlorophyll a and b contents were about 5-6
fold lower, and the total carotenoid content about 3 times lower, than observed for cells
acclimated tothe the low -light treatment. Carotenoid pigment composition and ratios were
significantly influenced by the light regime. Under the high-light conditions, the total carotenoid
vs. Chl a ratio was higher, with zeaxanthin and violaxanthin being the more abundant pigments.
Under the medium-light treatments, compared to values obtained in cells exposed to high light,
the violaxanthin and neoxanthin levels increased ~3-fold, whereas the zeaxanthin content
remained relatively unchanged compared with cells grown at high -light.at the same level. Under
low-light exposure, relative to the medium-light treatment, violaxanthin and neoxanthin levels
approximately doubled again, together totalling ~70% of the total carotenoid content. The -
carotene content also significantly increased, by 7.5-fold, becoming the third most abundant
pigment, whereas it was virtually absent in the cells exposed to the high-light treatment.
RuBisCO protein. Polyclonal antibodies raised against the Type 1B form of RuBisCO from
tobacco recognized a single protein with an mw of ca. 55 kDa in G. chlorophorum whole cell
protein extracts (Fig. 4). The size of the protein is similar to that of Dunaliella tertiolecta and
Pyramimonas parkeae. Antibodies raised against the Type 1D enzyme form from Isochrysis
galbana did not cross-react with protein extracts from G. chlorophorum orand from the green
algae, but did cross react with proteins from I. galbana (i.e., a positive control; results not
shown). Neither antibody cross-reacted with protein extracts from the peridinin-containing
dinoflagellate Amphidinium carterae or the diatom Skeletonema costatum.
Chloroplast genome size. The chloroplast plastid chromosome of G. chlorophorum and three
other phytoplankton organisms was isolated by pulsed-field gel electrophoresis and identified by
Southern blot analysis using a psbA gene probe (Fig. 5). Most of the plastid chromosomes, likely
15
present as minicircular molecules, did not migrate in the pulsed-field gel and remained in the
plugs, which were subsequently intensively probed in the Southern analysis (Fig 5B). A fraction,
however, was in a linear form that migrated in the pulsed-field gel. The chloroplast genome of G.
chlorophorum migrateds with an apparent size of ca. 100 kb (Fig 5B). The chloroplast genome
was virtually undetectable after staining with ethidium bromide, but was clearly visible as a weak
band after 32P labelling (Fig 5A-B). For comparison, the plastid genome size from Isochrysis
galbana, Nannochloropsis oculata and Heterosigma akashiwo were estimated as being
approximately 112, 125 and 165 kb, respectively. No plastid genome could be resolved and
detected in the two other species (D. tertiolecta and Amphidinium carterae).
Plastid gene phylogeny. To determine the origin of the chloroplast, we sequenced the rbcL
and psbA genes from G. chlorophorum DIN3 (accession numbers AY331683 and AY331682,
respectively). The chloroplast genome clearly encodes for the green type 1B RuBisCO. In order
to include the most representative unicellular green taxa, phylogenetic analyses were performed
from a partial sequence alignment spanning 1070 nucleotide positions (Fig. 6). Secondary plastid
data from euglenophytes and chlorarachniophytes, characterized by high genetic divergences
compared to all primary plastid rbcL genes, were not included in the analysis. Given the high
genetic divergence observed for G. chlorophorum, all the secondary green plastid taxa grouped
together with Mesostigma, likely as a result of long branch attraction (trees not shown). In all of
the analyses, the Prasinophyceae constituted a solid cluster, that included Tetraselmis when using
the first and second positions of amino acid codons (Fig. 6A), but not when using all three codon
positions (Fig 6B). In all analyses, G. chlorophorum was rejected from the prasinophyte cluster,
and branched within the Trebouxiophyceae with strong (Fig. 6A) to moderate (Fig 6B) support.
The psbA nucleotide sequence obtained from G. chlorophorum strain DIN3 is identical to that of
the type strain BAH ME 100. This gene is clearly affiliated with the green plastids, though
16
showing high divergence, as observed for rbcL. In the phylogenetic trees of primary chloroplast
genes (Fig 7), the G. chlorophorum sequence branched into a cluster including the
Chlorophyceae-Trebouxiophyceae, which also included Tetraselmis marina CCMP898
(Prasinophyceae, Chlorodendrales), but was distinctly separate from the sequences obtained for
the Prasinophyceae (Nephroselmis), Streptophyta (Mesostigma) and Glaucophyta (Cyanophora).
Effect of temperature and light on growth rate. During the three-week incubation period in
the temperaturtute-light gradient, significant growth was observed in only a few of the 60
temperature-light pairs. Growth was observed between 15 and 24° C, with irradiances ranging
from 10 to 520 µmol quantaE•m-2•s-1 (Fig. 8). Growth rate peaked sharply at 17-19° C and at 150-
200 µmol quanatE•m-2•s-1 irradiance regimes, respectively. The fastest growth rate, ~0.9 div•d-1,
was measured at 17° C and 180 µmol quantaE• m-2•s-1. However, extrapolation from the observed
k values suggested that this strain might be able to grow at 1 div•d-1. Outside the range of
optimum conditions, the growth rate decreased dramatically to <0.3 div•d-1.
PhotopPhysiological parameters related to photosystem II (PSII) were estimated from the
analysis of variable fluorescence from cultures in the exponential phase of growth (Table 4). The
functional cross-section of PSII (PSII) did not show variations that were related to growth
irradiance. Under high- and low–light acclimationirradiance exposure (relative to the optimum
range of 150-200 µmol quantaE•m-2•s-1), the photosynthetic efficiency (Fv/Fm) was significantly
lower, as was the time constant determined for electron transport on the acceptor side of PSII (i.e.
Qa, the time for plastoquinone Qa re-oxidation).
Discussion
Ultrastructural and molecular characterization of the studied strain. TEM analysis
ofUltrastructure of thin sections of strain DIN3, as seen using transmission electron microscopy,
reveals that the ultrastructure is similar to thatthe originally describedption for of the species G.
17
chlorophorum by (Elbrächter and Schnepf (1996), with no evidence of either external or internal
body scales outside the cells, or in formation inside the cells. In contrast, the green dinoflagellate
Lepidodinium viride appears to possess basket-shaped scales (Watanabe et al. 1987, Watanabe et
al. 1990). The identification of strain DIN3 as G. chlorophorum was confirmed by the identity of
a 3.4 kb rDNA sequence which was obtained from the type strain for this species. However, the
identity of the 18S rDNA sequence is also identical towith that deposited from L. viride
(Saunders et al. 1997), with the exception of a few mismatching nucleotides at the 3’-end of the
latter sequence:; these mismatches are consistent with the reverse PCR primer used byin
Saunders et al.’s work. (1997). Therefore, thei morphological differences butand yets genetic
identity raises the question about the identity of these two clonesstrains; :are the two strainsy in
fact, the same species?
The L. viride strain used for rDNA sequence analysis was identified by light microscope
only (G. SaundersD.R.A Hill, personal communication); however, the morphology of this
dinoflagellate is very similar to that of G. chlorophorum. To detect the major morphological
feature that differentiates the two species, namely the presence of body scales, requires TEM
and/or SEM analysis (Botes et al. 2002). Based on morphology, L. viride is considered to belong
to the Gymnodinium clade (Daugbjerg et al. 2000). Furthermore, the carotenoid composition of
our G. chlorophorum DIN3strain is similar to that described for L. viride (Watanabe et al. 1991).
In this latter species, an unidentified small peak eluting at the same time as lutein was also
reported, and the absence of -carotene can be explained if relatively high irradiances were used
for cultivation (cf. Table 3). Therefore, it is possible that G. chlorophorum and L. viride are truly
sibling species that diverged too recently for the introduction of mutations in their 18S rDNA
sequence. In order to substantiate this conclusion, the homologous rDNA sequence obtained from
a formally identified L. viride strain is required.
18
Does the G. chlorophorum contain a vestigial endosymbiont or a true plastidcontain a
nucleomorph genome? The double-membrane surrounded endosymbiont, described in G.
chlorophorum and L. viride, appears to enclose multiple and/or highly reticulated chloroplasts. In
addition, a vesicle with a double membrane contained within the plastidic periplasm has been
suspected to be a vestigial nucleus-like structure, but the presence of genomic DNA within it has
not been verified (Watanabe et al. 1990, Elbrächter and Schnepf 1996). HoweverIn the present
study, we could not detect any 18S rRNA gene other than that from the G. chlorophorum
nucleus. The set of PCR primers we used should amplify most eukaryotic 18S rRNA genes,
including those from Viridiplantae and the chlorarachniophyte nucleomorphs. If an
endosymbiont 18S rDNA could have been amplified by the PCR, in the subsequent RFLP
analysis, it is unlikely that any other 18S rDNA present it would not contain at least one distinct
restriction site recognized by one of the nine enzymes we used. In addition, genetic
heterogenityheterogeneity would likely have been detected by when performing the direct
sequencing of PCR products. We concludeNevertheless, it We cannotcannot be exclude the
possibilityed that the PCR amplification of a putative endosymbiont 18S rDNA could have been
prevented byfor unknow reasons, for example, (i) mutationsimportant genetic alterations that
would have renderedmade the eukaryotic primer set inappropriate, or (ii) the presence of introns
that would made the gene too large to be amplified with our PCR protocolsconditions, (which
are suitable for most eukaryotic organisms). While it is difficult to prove a negative, Altogether,
however, our negative result suggests the absence of an endosymbiotic 18S rRNA gene
structurally similar to that existing in most eukaryotes, and in cryptophyte and
chlorarachniophyte nucleomorphs (Gilson and McFadden 1996, Douglas et al. 2001). Hence, we
suggest that it is suggested that a nucleomorph genome, at least with a structure similar to that
known in cryptophytes and chlorarachniophytes, which contain three chromosomes capped with
19
a rRNA operon at both ends (Gilson and McFadden 1996, Douglas et al. 2001), is unlikely to
exist in G. chlorophorum probably does not have a nucleomorph genome,; however, but .further
investigations are required to verify this hypothesis. Based on this negative result, it appears that
G. chlorophorum contains true plastids that are not part of an endosymbiont.
The G. chlorophorum chloroplast genome. Our results suggest the existence of a
conventionalare the first report of an intact chloroplast genome from ain green dinoflagellates. In
This contrasts with peridinin-containing dinoflagellates, in for which the plastid genome appears
to be composed of multiple minicircles, each containing one or two genes (Zhang et al. 1999).
Consistently, in these organisms, data from expression libraries indicated massive transfer of
plastid genes to the nuclear genome (Bachvaroff et al. 2004, Hackett et al. 2004). This does not
appear to be the case for green dinoflagellates. The G. chlorophorum chloroplast genome appears
to contain be a single long genome, yet is among the smallest reported for unicellular,
photosynthetic algae (Coleman and Goff 1991). The 100 kb genome is ~20% smaller than the
chloroplast genome of Mesostigma viride, however the latterwhich contains the highest number
(99) of protein coding genes among the green plastid genomes sequenced to date (Lemieux et al.
2000). Compared with Mesostigma, prasinophycean and chlorophycean algae have a reduced
number of chloroplast genes as a result of gene losses (Grzebyk et al. 2003), (Grzebyk et al.
2003); however, the latter contain longer intergenic spacers and inverted repeat regions, resulting
in larger chloroplast genome sizes with lower densities of coding information (Wakasugi et al.
1997, Turmel et al. 1999, Maul et al. 2002). In some respects, a similar pattern of evolution also
occurred in the secondary chloroplast genome of euglenophytes (Hallick et al. 1993). Analysis of
all known plastid genomes reveals that secondary green or red plastids contain fewer genes than
primary algal plastids (Grzebyk et al. 2003). The plastid in G. chlorophorum almost certainly is a
secondary organelle (see discussion below), hence, we predict that it contains fewer genes than in
20
chloroplast genomes of primary symbiotic green algae. In the other phytoplankton species, the
apparent plastome size in I. galbana (112 kb) is similar to that determined by sequencing in the
Isochrysidiale Emiliania huxleyi strain CCMP 373 (105.3 kb, Sánchez Puerta et al. 2004); we
also obtained an apparent size of 112 kb for the strain CCMP 373 using PFGE analysis (data not
shown), suggesting PFGE analysis leads to slight overestimates of that plastome sizes estimated
in this way could be slightly overestimated. Based on PFGE analysis, slightly higher plastome
size have also been reported compared to the actual size subsequently determined by sequencing
in Chlamydomonas reinhardtii and Cyanidioschyzon merolae (Maleska 1993, Maul et al. 2003,
Ohta et al. 2004). Accordingly, the plastome size estimated here for H. akashiwo (~ 165 kb) is
consistent with a previous estimation (~150 kb, Reith and Cattolico 1986). In A. carterae, we
could not detect a plastid genome, possibly because of the minicircle structure of this genome
and/or the high divergence of plastid genes (e.g. psbA) in this dinoflagellate species compared to
other unicellular algae plytoplankters (Zhang et al. 1999, 2000).
What is the origin of green dinoflagellate plastids? The two G. chlorophorum plastid genes
we sequenced diverge significantly relative to all primary plastids. This situation is similar to
that observed in the two other secondary green plastid taxa (euglenophytes and
chlorarachniophytes). None of the phylogenetic reconstructions inferred from the plastid gene
sequences (rbcL and psbA) indicated a likely relationship between the G. chlorophorum
chloroplast and those from extant Prasinophyceae. Instead, phylogenetic trees based on rbcL
sequence analysis suggest the G. chlorophorum chloroplasts are derived from the
Trebouxiophyceae (rbcL gene), while the psbA gene tree suggests that these plastids are derived
from a ChlorophyteChlorophyceae. Hence, given that Chlorophyceae and Trebouxiophyceae are
likely closely related clades (Courties et al. 1998, Marin and Melkonian 1999, Fawley et al. 2000,
Maul et al. 2002), one possible explanation for the two different gene trees is that the plastid in
21
the green dinoflagellate was acquired prior to the divergence of Trebouxiophyceae and
Chlorophyceaethese two clades. Alternatively, the apparent ambiguity of origin of the plastid in
the green dinoflagellate may simply reflect an increased tempo of evolution in the plastid
genome. Regardless, based on gene sequence analysis, it would appear that the source of the
plastid in G. chlorophorum was either a Trebouxiophyte or a Chlorophyte alga, but not a
prasinophyte.
The plastid genetic data of the G. chlorophorum strain are consistent with its pigment
composition, which resembles those present in the vast majority of green algae including the
Chlorophyceae, the Trebouxiophyceae, and the prasinophyceaen order Chlorodendrales
(including the genus Tetraselmis). This composition corresponds to the type 1 carotenoid series, ;
however, G. chlorophorum lacks lutein, which is generally abundant in greenthis type of algae
(Egeland et al. 1997, Jeffrey et al. 1997). Pigment composition was primarily used to infer on the
algal origin of dinoflagellate green plastids. The report of prasinoxanthin in the type strain was a
strong indication in favor of a prasinophyte origin of G. chlorophorum plastids (Elbrächter and
Schnepf 1996). However, this carotenoid is absent in our strain, as well the chlorophyllide
derivative MgDVP, which is present in most prasinophycean taxa (Latasa et al. 2004).
Loroxanthin and/or siphonaxanthin and ester derivatives, present in prasinophyte taxa that do not
contain prasinoxanthin (Latasa et al. 2004), were not detected as well. For the other green
dinoflagellate, L. viride, the prasinophycean origin of the endosymbiont was initially inferred
from the low chlorophyll a/b ratio value (Watanabe et al. 1987), which was in the range of those
measured in our strain (1.2-1.6). Watanabe et al. (1991) further speculated in favor of a
prasinophycean origin, considering that the absence of lutein is a characteristic of the
Prasinophyceae that possess siphonaxanthin or praxinoxanthin (i.e. the type-2 and type-3
carotenoid series, respectively; Egeland et al. 1997). Lutein, along with -carotene, are indeed
22
generally not detected, or rare, in algae containing the characteristic signature type-2 or type-3
pigments (Sasa et al. 1992, Egeland et al. 1995, Egeland et al. 1997, Yoshii et al. 2002, Latasa et
al. 2004). In G. chlorophorum DIN3, the absence of lutein and other -carotene derived
xanthophylls, regardless of the intensity of light used for growth, suggests that the transformation
steps between -carotene and lutein may have been lost. The genes responsible for the synthesis
of carotenoid pigments (as well as most genes for the synthesis of chlorophyllic pigments) are
located in the algal nucleus. Hence, the genes coding the enzymes involved in the synthesis of
lutein from -carotene appear to have been lost during the green plastid endosymbiosis process;
i.e. these genes were not transferred from the green algal nucleus to the dinoflagellate host cell
nucleus. While lutein is not required for photosynthetic function, the pigment (which does not
transfer light efficiently to the reaction center) does confer a tolerance to very high irradiance
(Sukenik et al. 1987).
Although we cannot conclusively rule out the possibility that prasinophyte-specific
photosynthetic pigments are absentwere lost as a result of gene losses during the plastid
aquisitionacquisition, both plastid gene phylogenies and pigment patterns clearly suggest that the
green plastids in G. chlorophorum originated from the modern
cChlorophytes.ceae/Trebouxiophyceae lineage.
Biochemical impact of acquiring green plastids for dinoflagellate cells. The acquisition of a
green plastid was accompanied by the gain of the green type I RuBisCO, replacing the type II
RuBisCO present in the peridinin-containing dinoflagellates (Whitney et al. 1995). In
fucoxanthin-derivative containing dinoflagellates, the replacement of the type II RuBisCO by the
type 1D also occurred as part of the plastid replacement process (Takishita et al. 2000, Yoon et
al. 2005). Although the type I RuBisCO is more efficient in terms of CO2 fixation than its type II
23
counterpart, this change did not increase the growth rate of this species compared to other
dinoflagellates.
In contrast, the plastid replacement had a significant impact on the elemental composition of
the green dinoflagellate (Ho et al. 2003, Quigg et al. 2003). The overall trace elemental profile of
G. chlorophorum is similar to that of green algae, showing a higher requirement for Fe and Cu
than that displayed by most secondary red plastid organisms (i.e. diatoms, haptophytes and
peridinin-containing dinoflagellates). In contrast, the C:N:P composition of G. chlorophorum
does not differ from that of peridinin-containing dinoflagellates, and displays a lower N:P ratio
than that which is found in green algae. Quigg et al. (2003) suggested that the differences in the
trace element composition between algal phyla were due to their plastids, which would in turn
reflect the redox state of the ocean. Accordingly, suboxicthe anoxic oceans favored the green
algae, whereas the oxidizeded oceans favored the secondary red plastid taxa. If this
hypothesissupposition is true, then one might ask how this would have affected the evolution of
green dinoflagellates. One possible scenariohypothesis is that green plastids were acquired by
dinoflagellates during an ocean anoxic event (OAE) that would have selectively favored the
predominence of green algae in the oceans during this period. The last significant OAE occurred
during the Cretaceous, about 93 million years ago, and was followed by a modest diversification
of dinoflagellates, as indicated by the increase in the number of species-specific cysts in the
microfossil record (Katz et al. 2004). Interestingly, according to the fossil record, this OAE did
not yield abundant prasinophycean cysts. In fact, the most recent occurrence of high
prasinophycean cyst abundance was during the Jurassic Toarcian OAE, about 185 million years
ago (Van de Schootbrugge et al. 2005).
Physiological ecology of G. chlorophorum. The maximum in vitro growth rate estimated
obtained in the strain DIN3 (~0.9-1.0 div•d-1) is in the upper range of that reported for the
24
dinoflagellates, including a variety of Gymnodinium species (Tang 1995, Doblin et al. 1999,
Band-Schmidt et al. 2004). However, the temperature-light gradient experiment revealed the
extremely narrow optimum growth conditions for G. chlorophorum. The narrow optimum
temperature range observed in vitro, 18 ± 2.0° C, is consistent with the seasonal occurrence of
blooms, which are mostly observed during late summer. The range of irradiances supporting
optimum growth (~150-200 µmol quantaE•m-2•s-1) corresponds to approximately the 10% light
depth in the open ocean at mid-day in the summer. This low light selection is unusual for many
green algae (Sukenik et al. 1990), and. This characteristic may, in part, be a reflection of be a
consequence of the pigment profile of this strain (i.e. a low carotenoid content and the absence of
lutein). For the green algae and land plants, these pigments have photoprotective and light-
harvesting functions, respectively, at high and low growth irradiances. This study documents that
G. chlorophorum decreases its cellular concentrations of Chls a+b and carotenoids at high growth
irradiances (Table 3). Since the functional cross-section of PSII (PSII) does not vary with
irradiance, we suggest that this alga photo-acclimates by decreasing its number of photosynthetic
units at high growth irradiances (Falkowski and Raven 1997). The carotenoid cell content,
however, varied with a lower slope of variation than the chlorophyll content, resulting in a 2-fold
higher carotenoid-to-Chl a ratio at low vs. high light. Nevertheless, this ratio remained low
relative to the values reported for chlorophytes. Under low light, coincident with the increase in
carotenoid content, this alga modifies its carotenoid composition by enhancing the xanthophyll
cycle to its maximum potential by increasing cellular violaxanthin concentrations. Under high
light, while G. chlorophorum is unable to produce lutein or to increase its absolute zeaxanthin
content, this alga may receive a photo-protective benefit from the doubling of zeaxanthin-to-Chl
a ratio.
25
Ecological considerations. G. chlorophorum blooms during summertime or in early autumn,
when the narrow temperature and light conditions supporting optimum growth are met in turbid
coastal waters (Mouritsen and Richardson 2003). These environmental conditions may also
decrease the exposure to UV radiation, which has been shown experimentally to decrease
motility and photosynthesis (Schäfer et al. 1993, Tirlapur et al. 1993). Despite its narrow
optimum growth conditions, G. chlorophorum exhibits specific physiological characteristics that
may account for its ability to form blooms. Due to the production of an abundant polysaccharide
mucilage, the viscosity of its surrounding medium would be expected to increase. The
degradation of this mucilage by bacteria may also lead to subanoxic conditions during bloom
events. Both conditions would be expected to reduce the mortality associated with zooplankton
grazing. In addition, during the late bloom phase, in concert with mucilage production, G.
chlorophorum cells are capable of forming dense jelly aggregates. G. chlorophorum is also
capable of producing temporary cysts, which may play a role in reseeding the water column with
regenerated cells in coastal areas.
Although in the contemporary ocean, green dinoflagellates are nowadays relatively rare
components of modern phytoplankton communities, on occasion, they can form visibly dense
blooms (“green tides”). The occurrence of local dinoflagellate green tides seems to have
increased during the last two decades (Sournia et al. 1992, Paulmier et al. 1995, Elbrächter and
Schnepf 1996). While theThis increased frequency of green tides may be an artefact relatedfor
encountering this phenomenon may be coincident with to the installation of phytoplankton
monitoring networks, however, one can ask how such bloomsit could have actually remained
previously unnoticed previously. The question is open as to whether or not this expansion may be
related to changes in coastal environments over the two last decades.
26
Acknowledgments
We thank E. Erard-LeDenn (IFREMER, Centre de Brest, Brest, France) for kindly providing the
strain of Gymnodinium chlorophorum DIN3, on behalf of Dr. Chantal Billard (Laboratoire de
Biologie et Biotechnologies Marines, Université de Caen, Caen, France), and Dr. Malte
Elbrächter (Wadden Sea Station Sylt, List, Germany) for giving a sample from the type strain
(BAH ME 100) of G. chlorophorum. We thank Dr. Steve Mayfield (The Scripps Research
Institute, La Jolla, California, USA) for kindly provinding the tobacco RuBisCO antiserum. We
are very grateful to Valentin Starovoytov (Division of Life Sciences, Electron Imaging Facility,
Rutgers University, Piscataway, New Jersey, USA) for preparing the transmission electron
micrographs, to Stephanie Christensen (Department of Oceanography, University of Hawai`i at
Manoa, Honolulu, Hawai`i, USA) for assistance with the HPLC pigment analyses, and to Joana
Barcelos E Ramos for assistance with the temperature/light growth experiment. We thank two
anonymous reviewers for constructive comments. This work was funded by the National Science
Foundation through the Biocomplexity program (NSF Grant OCE-0084032 EREUPT).
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36
Table 1: Primers and PCR conditions for cloning the selected DNA fragments, and additional
sequencing primers, that have been used in this study.
DNA fragments Forward primers Reverse primers
18S rDNA
Cloning 18S-F 5’-TCC TGC CAG TAG TCA TAT GC-3’ 18S-R 5’-TGA TCC TTC YGC AGG TTC AC-3’
PCR 94° C, 5 min; 30x (94° C, 1 min; 55° C, 30 s; 68° C, 2 min 30 s); 68° C, 7 min
Sequencing 18S-I1F
18S-I3F
5’-TCT AAG GAA GCC AGC AGG-3’
5’-GGG AGT ATG GTC GCA AGG-3’
18S-I2R
18S-I4R
5’-ACC AGA CTT GCC CTC CAA-3’
5’-CAA AGT CCC TCT AAG AAG-3’
ITSs-28S(D3) rDNA
Cloning 18S-I3F 5’-GGG AGT ATG GTC GCA AGG-3’ 28S-D3B 5’-TCG GAG GGA ACC AGC TAC TA-3’
PCR 94° C, 5 min; 30x (94° C, 1 min; 55° C, 30 s; 68° C, 2 min 30 s); 68° C, 7 min
Sequencing 18S-ITS1-F
28S-D1S-F
28S-D3S-F
5’-CTT AGA GGA AGG AGA AGT CG-3’
5’-TTA AGC ATA TWA GTA RGC GG-3’
5’-AGR RCT TTG RAA AGA GAG-3’
ITSs-R
28S-D2S-R
5’-STT CAY TCG CCR TTA C-3’
5’-TTG GTC CGT GTT TCA AGA CG-3’
RbcL
Cloning RBCLG-1F 5’-ATG KCW CCA MAA ACW GAR AC-3’ RBCLG-1389R 5’-TTC CAA ACT TCR CAN GCD GC-3’
PCR 94° C, 5 min; 35x (94° C, 1 min; 52° C, 30 s; 68° C, 2 min); 68° C, 7 min
Sequencing rbcL-GcIF 5’-AGC TCT TCG TAG ACT TCG TC-3’ rbcL-GcIR 5’-GAT GAC ACC ATG CAT CRC TC-3’
PsbA
Cloning PSBA2Fm 5’-YTW TAY ATI GGW TGG TTY GG-3’ PSBA4R 5’-GGR AAG TTR TGI GCR TTH CG-3’
PCR 94° C, 5 min; 30x (94° C, 1 min; 45° C, 1 min; 68° C, 2 min); 68° C, 7 min
37
Table 2. Pigment composition, accessory pigment-to-chl a ratios (w:w, using the violaxanthin response factor for neoxanthin and
antheraxanthin), and carotenoid percentages (w:w) of total carotenoid content, in G. chlorophorum (strain DIN3), D. tertiolecta (strain
CCMP1320) and P. provasolii (strain CCMP1203).
Pigments Chl a Chl b Neo Viola Anthera Zea Lutein Prasino -car -car T-Car
G. chlorophorum
Ratio vs. chl a 1.000 0.624 0.082 0.071 0.082 0.012 ND ND 0.027 0.021 0.294
Carotenoid % _ _ 28.00 24.10 27.73 4.08 - - 9.13 6.97 100
D. tertiolecta
Ratio vs. chl a 1.000 0.589 0.072 0.073 0.069 0.132 0.433 ND 0.049 0.194 1.023
Carotenoid % _ _ 7.06 7.18 6.75 12.85 42.35 - 4.81 19.00 100
P. provasolii
Ratio vs. chl a 1.000 0.742 0.065 0.023 0.056 0.224 0.053 0.280 0.003 0.076 0.780
Carotenoid % _ _ 8.28 2.97 7.24 28.74 6.74 35.93 0.39 9.71 100
Pigment abbreviations: Chl a, chlorophyll a; Chl b, chlorophyll b; Neo, neoxanthin; Viola, violaxanthin; Anthera, antheraxanthin; Zea,
zeaxanthin; Lut, lutein; Prasino, Prasinoxanthin; -car, -carotene; -car, -carotene; T-Car, total carotenoids.
38
Table 3. Pigment composition, accessory pigment-to-chl a ratios (w:w, using the violaxanthin response factor for neoxanthin and
antheraxanthin), and carotenoid percentages (w:w) of total carotenoid content, in G. chlorophorum (strain DIN3), in cells exposed to
three light levels. Data are the average ± range determined for duplicate analyses.
Pigments Chl a Chl b Neo Viola Anthera Zea -car -car T-Car
High Light
Content (fg cell-1)
Ratio vs. chl a
Carotenoid %
1238.8 ± 15.6
1.000
-
1070.6 ± 11.0
0.864 ± 0.002
-
73.7 ± 0.7
0.060 ± 0.001
14.76 ± 0.24
127.2 ± 1.7
0.103 ± 0.000
25.46 ± 0.16
59.0 ± 0.5
0.048 ± 0.000
11.80 ± 0.01
156.0 ± 1.4
0.126 ± 0.000
31.21 ± 0.07
4.3 ± 0.2
0.003 ± 0.000
0.86 ± 0.04
79.5 ± 0.3
0.064 ± 0.002
15.91 ± 0.05
499.7 ± 3.3
0.403 ± 0.002
100.00
Medium Light
Content (fg cell-1)
Ratio vs. chl a
Carotenoid %
2987.4 ± 110.4
1.000
-
2649.0 ± 49.0
0.887 ± 0.016
-
225.0 ± 2.97
0.075 ± 0.002
23.08 ± 0.03
319.1 ± 7.4
0.107 ± 0.001
32.73 ± 0.38
112.3± 2.0
0.038 ± 0.002
11.53 ± 0.34
156.9 ± 1.5
0.053 ± 0.001
16.10 ± 0.04
31.6 ± 0.5
0.011 ± 0.000
3.24 ± 0.01
129.9 ± 1.2
0.04 ± 0.001
13.32 ± 0.04
974.7 ± 11.5
0.327 ± 0.008
100.00
Low Light
Content (fg cell-1)
Ratio vs. chl a
Carotenoid %
6549.3 ± 82.8
1.000
-
5971.5 ± 88.7
0.912 ± 0.002
-
482.4 ± 0.8
0.074 ± 0.001
31.25 ± 0.13
567.1 ± 12.4
0.087 ± 0.001
36.73 ± 0.70
47.6 ± 1.7
0.048 ± 0.000
3.08 ± 0.10
99.4 ± 6.1
0.015 ± 0.001
6.44 ± 0.41
238.0 ± 1.1
0.036 ± 0.000
15.41 ± 0.03
109.9 ± 4.2
0.017 ± 0.002
7.09 ± 0.29
1543.9 ± 4.2
0.236 ± 0.002
100.00
Pigment abbreviations as provided in Table 2.
39
Table 4. Photosynthetic parameters measured using the FRR fluorescence method for G.
chlorophorum cells growing under various light and temperature conditions.
Culture conditions Fv/Fm PSII
(2•quantum-1)
Qa
(µs)
Light
(µE•m-2•s-1)
Temperature
(° C)
99 17 0.473 338.6 237
187 17 0.543 354.0 424
216 17 0.538 289.4 351
522 21 0.461 353.7 253
40
Figure captions
Fig 1: TEM micrographs of Gymnodinium chlorophorum. A. Section showing the nucleus (n),
chloroplasts (c) and mitochondria (m); note that no scales are visible inside or outside of the cell.
B. Group of chloroplasts with pyrenoid (p). Endosymbiont cytoplasm contains a double-
membraned vesicle (arrowhead) similar to that previously described as a nucleus-like structure
(Watanabe et al. 1987; Elbrächter and Schnepf 1996).
Fig. 2: RFLP analysis of 18S rDNA amplified by PCR from G. chlorophorum using nine selected
restriction enzymes cutting once the dinoflagellate gene. For all enzymes, digestion patterns
correspond to those predicted from the dinoflagellate nucleotide sequence.
Fig. 3: Elution profile of photosynthetic pigments from (A) G. chlorophorum, (B) Pycnococcus
provasolii CCMP1203, and (C) Dunaliella teriolecta CCMP1320. Peak numbers are: 0, solvent
front; 1, MgDVP; 2, neoxanthin; 3, prasinoxanthin; 4, violaxanthin; 5, antheraxanthin; 6, lutein;
7, zeaxanthin; 8, chlorophyll b; 9, chlorophyll a; 10, -carotene; 11, -carotene.
Fig. 4: Western blot analysis using anti-tobacco RuBisCO antibody. Lane 1: protein size marker;
Lane 2: Dunaliella tertiolecta CCMP1320; Lane 3: Pyramimonas parkeae CCMP724; Lanes 4
and 5: Gymnodinium chlorophorum (samples obtained from two different cultures and protein
extractions); Lane 6: Isochrysis galbana CCMP1323; Lane 7: Amphidinium carterae
CCMP1314; Lane 8: Skeletonema costatum CCMP1332.
41
Fig. 5: Pulsed-field gel electrophoresis and southern blot analysis by probing the psbA gene for
the visualization of G. chlorophorum chloroplast genome. (A) Ethidium bromide stained DNA.
(B) psbA gene probed DNA. Lanes 1 and 11: kb DNA weight ladder (MidRange I PFG Marker
from New England Biolabs, MA); Lanes 2 and 3: Heterosigma hakashiwo CCMP1680; Lanes 4
and 5: Nannochloropsis oculata CCMP525; Lane 6: Dunaliella tertiolecta CCMP1320; Lane 7:
Amphidinium carterae CCMP 1314; Lanes 8 and 9: Gymnodinium chlorophorum; Lane 10:
Isochrysis galbana CCMP1323. In the panel (A), In both panels, the black arrows points the I.
galbana plastid DNA with an apparent size of about 112 kb. In the panel (B), white arrowheads
point the G. chlorophorum chloroplast DNA sizing about 100 kb. Plastid DNA is indicated in N.
oculata by either white (panel A) or black (panel B) stars (*), and in H. hakashivo by the symbol
(<).
Fig. 6: Inferred phylogenetic tree from partial rbcL gene sequences (1070 nucleotides) from
unicellular green algae (Cyanobacteria were used as an outgroup). (A) Phylogeny based on first
and second codon positions of amino acid codons, as obtained from a Bayesian likelihood
analysis (program MrBayes v.3B4). Values Numbers at nodes are posterior probability values >
0.5. (B) Phylogeny based on threeall codon nucleotide positions as obtained from Bayesian
(MrBayes) or maximum likelihood (program PHYML) analyses. Main taxa or clusters only are
displayed. Values Numbers at nodes are bayesian posterior probability values > 0.5 (left), or
consensus values > 50 from 100 bootstrapped ML analyses (right when displayed). According to
this variety of analyses, the G. chlorophorum rbcL sequence appeared derived from the
Trebouxiophyceae cluster with moderate to high support. The DNA sequence accession numbers
are provided next to each species’ name.
42
Fig. 7: Inferred phylogenetic tree from partial psbA gene sequences (987 nucleotide positions)
from unicellular green algae (Cyanobacteria were used as an outgroup). (A)This phylogeny
Phylogenies was obtained using the first and second nucleotide positions of amino acid codons,
(A) from Bayesian likelihood analysis (MrBayes v.3B4) and, (B) from ML analysis (fastDNAml,
consensus tree from 100 bootstrapped analyses) using the first and second nucleotide positions
from encoded amino acids. (C) Phylogeny obtained from the all nucleotide positions; numbers at
nodes are bootstrap values from PHYML (left) and fastDNAml (middle) analyses, or posterior
probability values (right) from MrBayes analysis applying different gamma–distributed rates of
evolution for each codon position. The G. chlorophorum psbA sequence was consistently related
to the Chlorophyceae/Trebouxiophyceae cluster, which also included the Chlorodendrale
prasinophyte T. marina. Values Numbers at nodes are bootstrap values > 50% or posterior
probability values > 0.5; dashes (-) indicate lower values. The DNA sequence accession numbers
are provided next to each species’ name in the tree (A).
Fig. 8: Influence of light and temperature on the maximum growth rate kmax (div•d-1, estimated
during the exponential growth phase) of G. chlorophorum.
43