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 grzebyk@imcsmarine.rutgers.edu? Current address: Muséum National d’Histoire Naturelle, USM0505, Systématique et Ecotoxicologie des Microalgues, 12 rue Buffon, Case 39, 75231 Paris Cedex 053

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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

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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

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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.

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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

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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

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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,

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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.

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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

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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

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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

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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).

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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.

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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