Harmful Algae 109 (2021) 102120

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Harmful Algae
journal homepage: www.elsevier.com/locate/hal

Original Article

Diversity and toxicity of Pacific strains of the benthic dinoflagellate Coolia(Dinophyceae), with a look at the Coolia canariensis species complex
Yong Heng Phua a, Michael C. Roy b, Sarah Lemer c, Filip Husnik b, Kevin C. Wakeman d, *aSchool of Science, Hokkaido University, North 10, West 8, Sapporo 060-0810, Hokkaido, Japan
Okinawa Institute of Science and Technology Graduate University, 1919-1 Tancha, Onna-son, Okinawa, 904-0495, Japanc
University of Guam Marine Laboratory, 303 University Drive, UOG Station, Mangilao, Guam 96923, USAdInstitute for the Advancement of Higher Education, Hokkaido University, North 10, West 8, Sapporo 060-0810, Hokkaido, Japanb

A R T I C L E I N F O

A B S T R A C T

Keywords:
Cooliatoxin
Cryptic species
Harmful algal bloom
High-resolution MS
Phylogeny
Taxonomy

Coolia Meunier 1919 from benthic assemblages of Hawai’i and Guam were isolated and clonal cultures were
established from single cells. Cultures were identified to species-level based on 28S rRNA and ITS-5.8S rRNA
genes and tested for toxicity. In Hawai’i, two strains of C. malayensis were isolated. In Guam, a high biodiversity
was identified: four strains of C. malayensis, one strain of C. palmyrensis, one strain of C. tropicalis, one strain of
C. canariensis phylogroup III, and two strains forming a new phylogroup (phylogroup IV) of nontoxic
C. canariensis. Morphology of the new C. canariensis phylogroup was described using light microscopy andscanning electron microscopy. Mass cultures and methanol extracts of representative cultures (C. malayensis, C.
palmyrensis, C. canariensis, C. tropicalis) from Guam were prepared for liquid chromatography-mass spectrometry
analysis. Chemical analyses revealed yessotoxin analogue C56H78O18S2 is produced by C. malayensis, C. canar­
iensis phylogroup IV and C. palmyrensis, but other analogues, C57H80O18S2 and C58H86O18S2, were only found in
C. malayensis (Okinawa) and C. canariensis phylogroup IV. Individual toxin profiles were also different over time
for an Okinawa strain of C. malayensis (NIES-3637), highlighting intra and inter-species variation in Yessotoxinanalogue expression. Biological activity was tested using Artemia bioassay and toxicity was observed in Guamand Okinawa strains of C. malayensis. Strong support of four distinct clades within the C. canariensis species
complex was recovered in phylogenetic analyses, despite morphological similarities.

1. Introductionof species complexes within Coolia (Nascimento et al., 2019; David et al.,2020). It is likely that within this genus there is an unexplored diversitythat has yet to be teased apart, and to this end, Coolia is an intriguing
model to study the biodiversity and toxin production of marine benthic
dinoflagellates.
Coolia monotis Meunier 1919 is the type species for this genus. Other
species within this genus have been differentiated by variation in thethecal plate pattern such as the shape of the 1′ and 3′ plates in
C. tropicalis, C. malayensis and C. canariensis (Meunier, 1919; Faust,
1995; Fraga et al., 2008; Leaw et al., 2010), areolation in C. areolata(Ten-Hage et al., 2001), or pore size in C. guanchica and C. palmyrensis(Karafas et al., 2015; David et al., 2020). The more contemporary use of
DNA sequencing, has facilitated the identification of more species thatappear morphologically stagnant, or have phenotypic plasticity (Laz­a-Martinez et al., 2011; Leaw et al., 2016).
Molecular phylogeny of the 28S and ITS-5.8S rRNA gene from

Marine benthic dinoflagellates produce natural products that may be
toxic and can have negative implications for marine organisms andpublic health (Wang, 2008; Moestrup et al., 2014; Longo et al., 2020;
Mudadu et al., 2021). One such genus, Coolia Meunier 1919, has a broad
distribution, ranging from tropical to temperate waters (Gómez et al.,2016; Leaw et al., 2016), and some strains are known to produce toxic
compounds (e.g., cooliatoxin) (Holmes et al., 1995; Leaw et al., 2016).
Currently, eight species of Coolia are recognized (David et al., 2020);
four of these lineages, C. tropicalis, C. malayensis, C. palmyrensis, andC. santacroce are known to be toxic (Holmes et al., 1995; Rhodes andThomas, 1997; Laza-Martinez et al., 2011; Karafas et al., 2015). Still, ourcontemporary understanding of morphology within the genus, and theuse of molecular phylogenetics (mainly based on 28S and ITS-5.8S re­gions of the ribosomal RNA operon), have resulted in the identification* Corresponding author.
E-mail address: wakeman.k@oia.hokudai.ac.jp (K.C. Wakeman).
https://doi.org/10.1016/j.hal.2021.102120
Received 22 June 2021; Received in revised form 26 September 2021; Accepted 2 October 2021
Available online 13 October 2021
1568-9883/© 2021 Elsevier B.V. All rights reserved.

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C. canariensis recovered three separate phylogroups with strong nodal
support (David et al., 2014; Nascimento et al., 2019). Phylogroup I has
only been found in the Canary Islands (Fraga et al., 2008); phylogroup II
only in Trinidad Island, Brazil (Nascimento et al., 2019); phylogroup III
has been found in multiple locations including Hong Kong (Leung et al.,2017), South Korea (Jeong et al., 2012), Spain (Fraga et al., 2008;
Laza-Martinez et al., 2011; David et al., 2014) and Australia (Momi­
gliano et al., 2013). However, these geographic boundaries may be theresult of limited sampling effort. The ITS sequence for phylogroup I is
not available, however, pairwise comparison (p-distance) between
phylogroup II and III is ≥ 0.04, which is greater than the genetic dif­ferences between other recognized Coolia species (i.e., C. monotis, C.
malayensis and C. tropicalis) (Litaker et al., 2007; Nascimento et al.,2019). Despite this genetic variation, there is no consistent and reliablemorphological character unique to each C. canariensis phylogroup.
Certain strains of C. malayensis, C. palmyrensis, C. santacroce andC. tropicalis produce natural products that are toxic such as cooliatoxin, apolyether toxin analogous to yessotoxin (Holmes et al., 1995), andgambierone (Tibiriçá et al., 2020), which can be toxic to mice, in vitro
human cell lines and invertebrates (Rhodes and Thomas, 1997; Rhodes
et al., 2010; Karafas et al., 2015; Leung et al., 2017). One strain of
C. malayensis produces several analogues of yessotoxin (Wakeman et al.,2015). While C. malayensis, C. palmyrensis, C. santacroce and C. tropicalis
are generally known to produce toxins, some intraspecies variation ex­
ists. For example some strains of C. malayensis, and C. tropicalis are
non-toxic (Rhodes et al., 2014; Leaw et al., 2016).
The aim of this study was to understand the diversity and toxicity ofCoolia from Guam and Hawai’i; two under-sampled regions in the Pa­
cific Ocean. Coolia was isolated into clonal cultures from shallow reefs
from both locations and identified to species level based on 28S and ITS5.8S rRNA genes. Their chemical profiles and toxicity were also evalu­
ated using HPLC and Artemia bioassays, respectively. An existing cultureof C. malayensis (NIES-3637) was also examined, in order to gain aperspective into the intra-species/intra-strain variation of the chemical
profiles.

(60%, 70%, 80%, 90%, and three times 100%) for 5 min at each step andleft to air-dry overnight. They were then mounted on stubs and sputtercoated for 4 min with 5 µm gold and viewed using Hitachi N-3000
scanning electron microscope (Hitachi, Tokyo, Japan).
2.3. DNA extraction, PCR amplification and sequencing
Extraction of genomic DNA was performed on 5 cells in PCR tubes
containing 10 µl QuickExtract™ FFPE DNA Extraction Kit (Lucigen
corp., USA) and incubated at 56 ◦ C for 1 h then 98 ◦ C for 2 min. To
identify cultures at the species level, 28S rRNA (28S) and ITS1-5.8S-ITS2(ITS) genes were amplified and sequenced. Initial amplification of 18S,
ITS, and 28S rRNA genes was done using primers PF1 ~ 28-1483R(Daugbjerg et al., 2000; Leander et al., 2003) with MyFi™ Mix andthe following program on a thermocycler: initial denaturation 95 ◦ C for
1 min; 35 cycles of 95 ◦ C for 15 s, 52 ◦ C for 15 s, 72 ◦ C for 3 min 30 s;
final extension 72 ◦ C for 7 min. Then 1:100 dilution of the initial PCR
reaction was used as template for a nested reaction using primer set
D1RF1 ~ 25R1 (D1 region) and 305-27F ~ 852R -70 (D2 region)(Yamada et al., 2013) using EconoTaq 2 × Master Mix (Lucigen corp.,
USA), with the following program on a thermocycler: initial denatur­
ation 94 ◦ C for 2 min; 25 cycles of 94 ◦ C for 15 s, 52 ◦ C for 15 s, 72 ◦ C for
1 min 40 s; final extension 72 ◦ C for 7 min. To amplify the ITS region,
initial reaction using primer set SR4 ~ 25R1 (Takano and Horiguchi,
2006; Yamaguchi et al., 2006) with MyFi™ Mix and the following
program on a thermocycler: initial denaturation 95 ◦ C for 1 min; 35
cycles of 95 ◦ C for 15 s, 52 ◦ C for 15 s, 72 ◦ C for 3 min; final extension
72 ◦ C for 7 min. Then 1:100 dilution of that reaction was used as tem­plate for a nested reaction using primer set Lp1F1 ~ 25F1R (Howard
et al., 2009; Takano and Horiguchi, 2006) using EconoTaq 2 × MasterMix (Lucigen corp., USA), with the following program on a thermo­
cycler: initial denaturation 94 ◦ C for 2 min; 25 cycles of 94 ◦ C for 15 s,
52 ◦ C for 15 s, 72 ◦ C for 1 min 40 s; final extension 72 ◦ C for 7 min.
Supplementary table 1 lists the primer sequences.
Size of PCR products was checked on 1% agarose gel, purified with
Polyethylene Glycol (PEG), dye-labeled with BrilliantDye™ Terminatorv1.1 (Nimagen, Netherlands) according to manufacturer’s protocol, andsequenced on a 3130 genetic analyzer (Applied Biosystems, Massachu­
setts, USA). The sequences were checked using the National Center for
Biotechnology Information’s (NCBI) Basic Local Alignment Search Tool
(BLAST). Sequences generated in this study were deposited in the NCBI
under the following accession numbers: MZ801746-MZ801769.

2. Materials and methods
2.1. Sample collection and treatment
Seawater samples were collected from macroalgae and turf algae oncoral reefs in Pago Bay, Guam (13◦ 25′ 39.6" N 144◦ 47′ 54.1" E) and
Waimanalo, Hawai’i (21◦ 19′ 52.29′ ’ N 157◦ 41′ 12.40′ ’ W). Samples werebottled and brought back to the laboratory then separated into plastic
containers with half-concentration DAIGO IMK, and incubated for two
weeks at 25 ◦ C. Single cells of Coolia were isolated and grown in 48-wellplates in half-concentration DAIGO IMK growth medium. After thecultures became established (and cell density was visible by eye), they
were transferred to a Petri dish containing DAIGO IMK.
Coolia malayensis from Okinawa is the strain referenced in Wakeman
et al. (2015), acquired from NIES microbial culture collection(NIES-3637).

2.4. Phylogeny
The 28S and ITS rDNA sequences obtained in this study were aligned
using the MAFFT E-INS-I algorithm online (Katoh et al., 2002) alongwith Coolia sequences retrieved from GenBank and the alignments weremanually checked in Mesquite (Maddison and Maddison, 2019).
Alignments were trimmed using Gblocks, Version 0.91b (Castresana,
2000; Talavera and Castresana, 2007) with less stringent selection op­
tions; allow smaller final blocks, allow gap positions within the final
blocks, allow less strict flanking positions. Each locus was aligned
separately and Ostreopsis ovata was used as an outgroup. The final 28Salignment contained 100 sequences with 741 aligned nucleotides, andthe ITS rRNA alignment contained 72 sequences with 311 aligned nu­
cleotides (Supplementary file 1 and 2).
Iqtree 1.6.12 software was used to choose the best substitution modelaccording to Akaike Information Criterion with correction (Nguyen
et al., 2014). Substitution model TIM+F+G4 and TN+F+G4 were usedfor 28S and ITS alignments, respectively. Maximum likelihood (ML) tree
inference was run in Iqtree with 1000 bootstrap pseudoreplicates.
Bayesian analysis was performed with MrBayes v3.2.7 (Ronquist andHuelsenbeck, 2003). The General Time Reverable model with a gamma
distribution of site-rate heterogeneity was specified. FourMonte-Carlo-Markov chains were run for 1,000,000 generations and2.2. Morphological analysis
Cultured cells were fixed with 2.5% glutaraldehyde then washedwith seawater. The cells were then permeabilized at 20 ◦ C for 30 min
with an enzyme mix of 1% hemicellulose (Sigma-Aldrich, USA), 1%
cellulase (MP Biomedicals, California, USA) and 0.4 M mannitol (Wako
Pure Chemical Industries, Osaka, Japan), adjusted to pH 5.8 and stainedwith 0.1 µg/mL DAPI (Roche Diagnostics GmbH, Germany) then imaged
with Nikon eclipse Ti2 (Nikon, Tokyo, Japan). For scanning electron
microscopy (SEM), 1 ml of cell culture was fixed with 6% glutaraldehyde
overnight then filtered through a 10 μm MF-Millipore™ (Merck, USA)
membrane filter. Cells were then washed twice with distilled water anddehydrated through five steps with increasing ethanol concentration
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Harmful Algae 109 (2021) 102120

sampled every 100th generation. Tracer 1.7 (Rambaut et al., 2018) wasused to check convergence and posterior probability values were ob­tained from the consensus tree, after the first 25% was discarded asburn-in.
MEGA-X (Kumar et al., 2018) was used to calculate the pairwise
distances between the phylogroups for each the 28S and ITS genes. The
model selected was p-distance with transition and transversions;evolutionary rates among sites were set as gamma-distributed. Other
parameters were left at default settings.

3. Results
3.1. Morphological analyses C. canariensis phylogroup III and IV from
Guam
Coolia canariensis phylogroup IV isolated from Guam have an average
dorsal-ventral (DV) width of 28.78 ± 3.4 µm (range = 22 – 34 µm, n =28) and DV depth of 30.8 ± 3.3 µm (range = 23 – 37 µm, n = 21). The
nucleus is bean-shaped, central in the cell (Fig. 1). The plate formula isPo, 1′ , 7′′ , ?c, 5′′ , ?s, 2′′′′ . The surface of the cell is smooth, compared to
the rough, ornamented hypotheca (Fig. 2A). The apical pore is elongatedand slightly curved, average length was 6.80 ± 0.83 µm (range = 5.21 –7.75 µm, n = 9) and width is 1.53 ± 0.44 µm (range = 0.843 – 2.17 µm, n= 8), connected to plates 1′ , 2′ and 3′ (Fig. 2B). The 1′ plate is the largest
epithecal plate, hexagonal and connected to Po, 2′ , 3′ , 1′′ , 2′′ , 6′′ and 7′′
plates. Plate 2′ is thin rectangular, touching the Po, 1′ , 2′′ , 3′′ and 4′′
plates. Plate 3′ is pentagonal, contacting Po, 1′ , 2′ , 4′′ , 5′′ and 6′′ plates(Fig. 2B). Plate 1′′′ is trapezoidal, touching 1′ , 2′′ and 7′′ (Fig. 2B). Plate
2′′ is pentagonal, elongated at the cingulum and is contacting 1′ , 2′ , 1′′
and 3′′ plates (Fig. 2C). Plate 3′′ is rectangular, touching 2′ , 2′′ and 4′′
plates (Fig. 2D). Plate 4′′ is square, touching 2′ , 3′ , 3′′ and 5′′ plates(Fig. 2D). Plate 5′′ is trapezoidal, contacting 3′ , 4′′ and 6′′ plates (Fig. 2D,E). Plate 6′′ is pentagonal, touching 1′ , 3′ , 5′′ and 7′′ plates, the shortestside is contacting 3′ plate (Fig. 2E). Plate 7′′ is trapezoidal, contacting 1′ ,1′′ and 6′′ plates; it also extends further than plate 1′′ making an unevencingulum (Fig. 2A, E). Postcingulum plate 1′′′ is triangular, contacting
plate 2′′′ and 1′′′′ (Fig. 2A). Plate 2′′′ is trapezoidal, touching plates 1′′′ ,3′′′ and 2′′′′ (Fig. 2A, B, F). Plates 3′′′ and 4′′′ are trapezoidal touchingplates 2′′′ , 4′′′ , 2′′′′ and 3′′′ , 5′′′ , 2′′′′ respectively (Fig. 2F). Plate 5′′′ istriangular contacting plate 4′′′ (Fig. 2F). Of the post cingulum plates,
Plate 3′′′ is smoothest, 1′′′ and 5′′′ have the roughest surface followed byplates 2′′′ and 4′′′ (Fig. 2F). Plate 1′′′′ is protruding over the sulcal plates
contacting plate 1′′′ (Fig. 2F). Plate 2′′′′ is a small square surrounded byplates 1′′′ , 2′′′ , 3′′′ , 4′′′ and Sp (Fig. 2F).
Coolia canariensis phylogroup III from Guam and C. canariensis phy­logroup IV show the same plate morphology (Fig. 3A - F). It has roughly
spherical cells with a plate formula Po, 1′ ,7′′ , ?c, 5′′ , ?s, 2′′′′ , average DV
width 28.17 ± 1.72 µm (range = 25 – 38 µm, n = 6) average DV depth =28.0 ± 2.65 µm (range = 25 – 31 µm, n =7) (Fig. 3A - F). Apical pore issurrounded by plates 2′ , 3′ , 1′ , and is elongated and slightly curved withaverage length = 7.22 ± 0.91 µm (range = 8.46 – 5.92, n = 5) andaverage width = 1.31 ± 0.39 µm (range = 0.86 – 1.81, n = 4) (Fig. 3B).

2.5. Methanol extraction and desalting of crude extracts
Representative strains (NIES-3637, G10, F6-G23, G6, G11, G8) weregrown in mass cultures for toxin analysis. Cells were filtered throughWhatman® (Merck, USA) filter paper grade 1 and stored in 50 ml falcon
tubes in -20 ◦ C until ample material had been collected. Frozen tubes
were thawed for 30 min at room temperature then 25 ml methanol wasadded to each falcon tube, vortexed and sonicated for 5 min. After which
the samples were left at room temperature for 30 min with occasional
shaking then transferred into another flacon tube. Extraction was
repeated another two more times, the last extraction was left overnightat room temperature. The methanol extractions were centrifuged, dec­
anted and concentrated using SP Genevac EZ-2 Plus (SP Scientific, UK)
until each sample contained about 5 ml solution. The extract was
defatted by adding 5 ml ethyl acetate and vortexed for 10 min before
separating water-soluble and ethyl-acetate soluble parts. Defatting byaddition of ethyl acetate and separation water-soluble and ethyl-acetatesoluble parts was repeated two more times. A portion of water-soluble
part was desalted using ZipTip NIES-36378/P10 (Millipore) and theremaining dried using the evaporator and stored at -20 ◦ C and later
desalted using Supel™-Select HLB 1ml tubes (Merck, USA).
Zip tips were activated using aqueous-methanol-formic acid
(1:9:0.05) solution. Concentration of methanol was decreased with eachstep (90%, 50%, 10%), waiting 2 min between each step. Sample waspassed through the tip 10-15 times then washed with aqueousmethanol-formic acid (9:1:0.05). The retained sample was eluted with
10 μl aqueous-methanol-formic acid (1:9:0.05) into HPLC sample tubes
and diluted by adding 20 μl aqueous-methanol-formic acid (9:1:0.05).
Solid phase extraction tubes, 1 ml HLB tubes (30 mg/ml) wereactivated using 1 ml MeOH then 1 ml aqueous-methanol-formic acid
(9:1:0.05). Dried water-soluble part was dissolved using aqueousmethanol-formic acid (9:1:0.05) then passed over the HLB column andwashed with 1 ml aqueous-methanol-formic acid (9:1:0.05) two times.
Finally, the retained sample was eluted with 1 ml MeOH (30 mg/ml),
dried in the evaporator and stored in -20 ◦ C.

3.2. Phylogenetic analyses
Both analyses of 28S and ITS regions recovered seven distinct clades
representing seven species (Fig. 4 and 5). Strains G4, G10, E6-G27,
E2G25, NIES-3637, C12, C14 from Guam and Hawai’i grouped with
C. malayensis. Strains F6-G23 and G8 from Guam grouped with
C. palmyrensis and C. tropicalis, respectively. Based on the 28S

2.6. Nanoliquid chromatography mass spectrometry
Water-soluble part was dissolved in 200 μl MeOH and diluted 1:10 inaqueous-methanol-formic acid (9:1:0.05) and loaded into a
microcolumn-Liquid Chromatograph (Paradigm MS4, Microm Bio­
resources) connected to a Thermo Scientific Hybride Mass Spectrometer(LTQ Orbitrap) for mass spectrometry (MS) data collection. The mass
spectrometer parameter was set up following Wakeman et al. (2015).
2.7. Artemia lethality bioassay
Artemia eggs were placed in 0.22 μm-filtered seawater for 24 h tohatch. Ten nauplius were transferred into a 24 well plate containing 475
μl filtered seawater and 25 μl Coolia culture extract water-soluble frac­tion. Three wells (n = 30) were made for each Coolia extract. Watersoluble part from solid phase extraction HLB tubes was dissolved in
100 μl distilled water (Invitrogen, USA) and 25 μl was added to eachwell. Negative controls were seawater and 25 μl distilled water. Viabilityof Artemia nauplius was checked at intervals of 30 min, 1 h, 2 h, 4 h, 8 h,
12 h, 24 h, 30 h, and 48 h.

Fig. 1. A) Light microscopy and B) epifluorescence image of Coolia canariensis
phylogroup IV. Scale bar–10 μm.
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Fig. 2. Scanning electron micrographs of Coolia canariensis phylogroup IV. A) Ventral view. B) Dorsal view. C) Antapical view. D) Apical view. E) Apical right-lateralview. F) Right lateral view. Apical pore (Po), Sulcal plate (Sp). Scale bars–5 μm.

Fig. 3. Scanning electron micrographs of Coolia canariensis phylogroup III isolated from Guam. A) Ventral view. B) Dorsal view. C) Antapical view. D) Apical view. E)
Left leteral view. F) Right leteral view. Apical pore (Po), Sulcal plate (Sp). Scale bars–5 μm.

phylogeny, the C. canariensis clade contains four phylogroups withmoderate-to-high bootstrap and Bayesian posterior probability (≥70 /≥0.95). Phylogroup IV consists of two strains from Guam (G6 and G17),
and was sister to phylogroup III (Fig. 4). The ITS phylogeny of
C. canariensis clade consisted of six sequences and did not include

phylogroup I (Fig. 5). Strain G6 and G17 were grouped with highbootstrap values and Bayesian posterior probability (100/1) in phy­logroup IV, sister to phylogroup III. Strain G11 from Guam did not show
strong support as a member of phylogroup III (Fig. 5). Pairwise com­
parison of 28S rRNA gene showed the average p-distance within each4

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Fig. 4. Maximum likelihood phylogenetic tree inferred from 28S sequences using TIM F G4 substitution model. Bootstrap support lower than 60% and Bayesianposterior values lower than 95% are not displayed. Sequences published from this study are written in white, bold and highlighted.

species was highest in C. canariensis (p = 0.079 substitutions per site),followed by C. palmyrensis (p = 0.0718); the remaining Coolia species
ranged between p = 0.0016 - 0.0138 (Supplementary table 2). Mean
distance between species was shown to be less between C. monotis, C.
malayensis and C. santacroce (p = 0.106 - 0.148), than phylogroups of
C. canariensis and strains G6 and G17 (p > 0.15) (Supplementary table3). This was also matching in p-distance calculated for ITS, where

C. canariensis strain G6 and G17 was p > 0.3 in relation to strain G11,
and previously studied phylogroups II and III (Supplementary table 4).
3.3. Mass profile of Coolia culture extract
High-resolution mass (HRMS) was used to identify cooliatoxin inwater-soluble part which were previously identified toxins by Wakeman
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Fig. 5. Maximum likelihood phylogenetic tree inferred from ITS sequences using TN F G4 substitution model. Bootstrap support lower than 60% and Bayesianposterior values lower than 95% are not displayed. Sequences published from this study are written in white, bold and highlighted.

et al. (2015). Coolia malayensis strain NIES-3637 used was the samestrain as the previous study, cooliatoxin 1 and 2 at m/z 1101.46
C56H78O18S2 [M− H]− Δ 5.14 mmu, and 1115.47 [M− H]− Δ 2.43 mmu
respectively were identified (Supplementary fig. 2A, B). MS2 fragmen­
tation of toxin compound 1 showed the sulphonate loss [M− SO3− H]− at
m/z 1021.41, MS3 fragmentation showed water loss [M‒SO3‒H2O‒H] −at m/z 1003.50 (Supplementary fig. 2D - G). Due to low signal intensity,
the peak for toxic compound 2 at m/z 1115.47 was not selected for MS2
fragmentation (Supplementary fig. 2G).
From strain G10 from Guam, compound 1 at m/z 1101.45,
C56H78O18S2 [M− H]− Δ 1.42 mmu and compound 5 m/z 1133.5213
C58H86O18S2 [M− H]− Δ 4.08 mmu were detected (Fig. 6). In
C. palmyrensis strain F6-G23, a peak at m/z 1101.45 C56H78O18S2[M− H]− Δ 1.467 mmu (Fig. 7) and C. canariensis phylogroup IV strain
G6 m/z 1101.45, C56H78O18S2 [M− H]− Δ 2.66 mmu, m/z 1133.52
C58H86O18S2 [M− H]− Δ 9.14 mmu were detected (Fig. 8). In these threestrains, MS2 fragmentation was not observed due to less intense signal.
Peaks matching the toxin compounds were not detected in C. canariensis
phylogroup III (Strain G11) nor C. tropicalis (Strain G8).

3.4. Artemia lethality bioassay
There was no visible change for the first 8 h after addition of thewater-soluble part. After 12 h, Artemia with the addition of C. malayensis
extracts (NIES-3637, Okinawa and G10, Guam) had impaired mobility.
After 24 h, Artemia with NIES-3637 extracts were alive however motility
severely diminished. Artemia nauplius in wells with G10 extracts had
10% mortality and severely diminished mobility (Supplementary video1). After 30 h, Artemia with extracts from NIES-3637 were no longer
capable of swimming, and similar to G10, had a 33% mortality rate.
After 48 h, mortality was detected for NIES-3637 (avg. death = 36%),
majority of Artemia with G10 extracts were dead (avg. death = 93%).
Toxic effects of the water-soluble fraction on Artemia for C. palmyrensis(F6-G23) and C. canariensis (G6, G11) were not observed.
4. Discussion
4.1. Global distribution of Coolia
In the present study, 11 strains of Coolia from Guam and Hawai’i
were established. Despite our limited geographical sampling, a larger
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Fig. 6. NanoLCMS (negative ion) profile of water-soluble extract of Coolia malayensis strain G10. A) Chromatogram; B) extract ion (m/z 1101.45); C) extract ion (m/z1101.45); D) compound 1 (m/z 1101.4560, RT–15.96-16.22 min); E) compound 5 (m/z 1133.5213, RT–14.34 – 14.85 min).

than expected diversity of Coolia was found. Phylogenetic analysis of
28S and ITS rRNA genes of strains from Pago Bay, Guam, grouped into
four of the eight known species (C. malayensis C. palmyrensis, C. tropicalis
and C. canariensis). The high diversity of Coolia species is similar toprevious studies in Hong Kong (Leung et al., 2017), Great Barrier Reef
(Momigliano et al., 2013), and Brazil waters (Tibiriçá et al., 2020),
indicating a broad distribution of Coolia. This supports the idea thatmany species within the Coolia genus have overlapping geographic
ranges, and suggests a continuous distribution, and rich diversity ofCoolia, throughout the Western Pacific. Conversely, the distribution of
C. monotis has only been isolated from Europe, the Mediterranean Sea
and North Atlantic, which is evidence that some species do haverestricted distributions, possibly due to habitat restrictions like tem­
perature (Karafas et al., 2015; Lewis et al., 2018).

have been supported by molecular data (Faust, 1995; Fraga et al., 2008;
Leaw et al., 2010; Mohammad-Noor et al., 2013; Karafas et al., 2015).
Coolia tropicalis has a smooth thecal surface, differentiating it from
C. areolata which is heavily areolated (Ten-Hage et al., 2001). The epi­
theca of C. canariensis is smooth but the hypotheca is ornamented
prominently in postcingular plates 1′′′ , 2′′′ , 5′′′ and 1′′′′ (Fraga et al.,2008; Fig. 2), distinguishing it from others. Coolia guanchica has fewer
hypothecal ornamentations but contains larger and more numerous
pores, compared to C. canariensis and a smoother surface than C. areolata(David et al., 2020; Supplementary Fig. 1).
However, throughout the C. canariensis phylogroups, this morpho­
logical approach has been problematic. When using the 28S rRNA geneas a molecular marker, newly isolated strains and other isolates fromprevious studies, form four well-supported clades: phylogroup I, II, III,and IV. Phylogroup IV is a sister to phylogroup III, isolated from bothAtlantic and Pacific Oceans, while phylogroup I and II have only been
isolated from the Atlantic Ocean. In addition, the ITS p-distance values
between strains in phylogroup IV and other phylogroups (II, III) is morethan 0.04 which is often able to separate most free living dinoflagellate
species (Litaker et al., 2007). The secondary structure of ITS2 also beenused for delimitation of species boundaries, however, this was not per­formed in this study (Leaw et al., 2016). The differences in genetics andgeographical location would indicate that phylogroup IV represents a
new species, distinct from other known phylogroups in the C. canariensis
clade (Jeong et al., 2012; Nascimento et al., 2019; David et al., 2020).

4.2. Examining the phylogeny and diversity of C. canariensis species
complex
This study also found a new non-toxic phylogroup within the
C. canariensis species complex (phylogroup IV). The morphology ofstrains G6 and G17, which formed phylogroup IV, was similar to strain
G11 (phylogroup II) and other C. canariensis phylogroups. Generally
speaking, Coolia have been morphologically characterized by their
thecal plate patterns and other ornamentations of the thecal plates.
Coolia monotis, for example, has a 6′′ plate which is the largest epithecaplate, shifting the 1′ plate to the left of center. This is distinct fromC. tropicalis, C. areolata, C. guanchica and C. canariensis where the 1′ plate
is located centrally on the epicone. These morphological characteristics
7

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Harmful Algae 109 (2021) 102120

Fig. 7. NanoLCMS (negative ion) profile of water-soluble extract of Coolia palmyrensis strain F6-G23. A) Chromatogram; B) extract ion (m/z 1101.45); C) compound 1(m/z 1101.4561, RT–15.98 min).

Fig. 8. NanoLCMS (negative ion) profile of water-soluble extract of Coolia canariensis strain G6. A) Chromatogram; B) extract ion (m/z 1133.51); C) compound 1 (m/z 1101.4589); D) compound 5 (m/z 1133.5263, RT–14.17 min).

4.3. Variability and diversity of toxic profiles of Coolia

to Artemia in our bioassays. And due to low abundance, confirmation of
a YTX analogue in LCMS was not achieved. Similarly, the Guam strain of
C. canariensis phylogroup III tested negative in both Artemia assay and

The new lineage of C. canariensis from Guam appeared to be nontoxic
8

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Harmful Algae 109 (2021) 102120

LCMS profile which matches with previous studies where C. canariensis
phylogroup III appears to be a nontoxic lineage (Fraga et al., 2008;
Laza-Martinez et al., 2011; Leung et al., 2017).
Both C. malayensis strains NIES-3637 and G10 were toxic to Artemia
in our bioassays. This was confirmed with LCMS showing YTX analogues
with characteristic MS2 fragmentation pattern of sulphonate loss, – SO3(80 dalton). Differences in LCMS profile from strains G10 and NIES-3637
show intraspecies variation on the type and amount of toxins being
produced. This result supports previous bioassay-based reports
observing intraspecies variation in toxicity (Leung et al., 2017). Toxicity
of each strain could be due to a mixture of different chemical compounds
being produced by the dinoflagellates. Furthermore, in this study the
LCMS profile did not detect several compounds (compounds 3 and 4)
from of C. malayensis strain identified by Wakeman et al. (2015). This
indicates that the variation in toxicity of clones may be influenced by not
only toxin concentration but also by the type of toxic analogues pro­
duced (Neely and Campbell, 2006).
Coolia palmyrensis strain F6-G23 tested negative in the Artemia
bioassay showing variability compared to strains from the central Pacificand West Atlantic (Karafas et al., 2015). This may be attributed to the
low abundance of the potentially toxic compounds, or variation insensitivity of organism used in the assay (Leung et al., 2017). Coolia
tropicalis (strain G6), tested negative in the bioassay as well as LCMS.
However other strains have been shown to be toxic (Faust, 1995; Rho­des et al., 2014; Tibiriçá et al., 2020). This particular strain was unable
to form a dense culture and eventually died which could have influenced
toxin production, due to sub-optimal growth conditions. Abundance ofdetected toxins did not directly relate to the lethality of each strain. The
overall lethality could be therefore influenced by other compounds suchas gambierones (Tibiriçá et al., 2020) or other compounds yet to be
identified.

for the Advancement of Higher Education at Hokkaido University.
Sampling for this project was supported by the University of GuamMarine Laboratory. We would also like to thank the Hokkaido University
School of Agriculture for providing access to their EM Facilities, andOkinawa Institute of Science and Technology, (Instrumental Analysis
Section) for usage of their facilities.
Supplementary materials
Supplementary material associated with this article can be found, inthe online version, at doi:10.1016/j.hal.2021.102120.
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5. Conclusion
Tropical and subtropical regions of the Pacific Ocean harbor a large
biodiversity of the genus Coolia. In this study, four of the eight known
species, C. malayensis, C. palmyrensis, C. canariensis, C. tropicalis, were
found in Guam. A new non-toxic (exibits minimal toxic effect) phy­logroup (phylogroup IV) belonging to the Coolia canariensis species
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Author’s contribution
PYH and KCW designed the project; KCW conducted field sampling;
PYH and KCW isolated and establish clonal strains of Coolia. PYH and
MCR conducted experiments and analysis; PYH, KCW, MCR, SL, FH
interpreted the data, drafted and approved the manuscript.
Declaration of Competing Interest
The authors declare that they have no known competing financial
interests or personal relationships that could have appeared to influence
the work reported in this paper.
Acknowledgments
This work was supported by Japanese Society for the Promotion of
Science (JSPS) grants 18K14774 and PG6R180004 to KCW; additional
funds and administrative support were provided to KCW by the Institute9

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