Harmful Algae 121 (2023) 102356
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Harmful Algae
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Morphology and phylogeny of Prorocentrum porosum sp. nov.
(Dinophyceae): A new benthic toxic dinoflagellate from the Atlantic and
Pacific Oceans
Edgar Arteaga-Sogamoso a, b, *, Francisco Rodríguez c, d, Alberto Amato e, Begoña Ben-Gigirey d,
Santiago Fraga f, Luiz Laureno Mafra Jr. g, Luciano Felício Fernandes h,
Carlos Eduardo J. de Azevedo Tibiriçá g, Nicolas Chomérat i, Tomohiro Nishimura j, k,
Chiho Homma k, Masao Adachi k, José Ernesto Mancera-Pineda l
a
Instituto de Investigaciones Marinas y Costeras José Benito Vives de Andréis, INVEMAR, Santa Marta, Colombia. Calle 25 No. 2-55, Playa Salguero, Rodadero, Santa
Marta, Colombia
b
Universidad Nacional de Colombia, sede Caribe, Santa Marta, Colombia. Calle 25 No. 2-55, Playa Salguero, Rodadero, Santa Marta, Colombia
c
Centro Nacional Instituto Español de Oceanografía (IEO-CSIC), Centro Oceanográfico de Vigo. Subida a Radio Faro 50, 36390 Vigo, Spain
d
European Union Reference Laboratory for Monitoring of Marine Biotoxins, Citexvi Campus Universitario de Vigo, 36310, Vigo, Spain
e
Laboratoire de Physiologie Cellulaire et Végétale, Université Grenoble-Alpes CEA CNRS INRA IRIG-CEA Grenoble, 17 rue des Martyrs, 38054 Grenoble Cedex 9, France
f
Praza Mestra Manuela 1, 36340 Nigrán, Spain
g
Centro de Estudos do Mar, Universidade Federal do Paraná, P.O. Box 61, 83255-976, Pontal do Paraná, PR, Brazil
h
Departamento de Botânica, SCB, Centro Politécnico, Universidade Federal do Paraná, P.O. Box 19031, 81531-990, Curitiba, Paraná CEP Brazil
i
Station de Biologie Marine, IFREMER, Littoral, LER BO, Place de la Croix, F-29900, Concarneau, France
j
Cawthron Institute, 98 Halifax Street East, 7010 Nelson, New Zealand
k
Laboratory of Aquatic Environmental Science (LAQUES), Faculty of Agriculture and Marine Science, Kochi University, 200 Otsu, Monobe, Nankoku, 783-8502, Kochi
Japan
l
Universidad Nacional de Colombia, sede Bogotá, Carrera 45 No. 26-85, Bogotá D. C. Colombia.
A R T I C L E I N F O
A B S T R A C T
Keywords:
Prorocentrum porosum
Taxonomy
Phylogeny
LSU
ITS
Okadaic acid
A new marine benthic toxic Prorocentrum species is described from the tropical/subtropical regions of the
Atlantic (Colombian Caribbean Sea and Northeast Brazil) and Pacific (Southern Japan) oceans. Morphological
cell structures were examined using light (LM) and scanning electron (SEM) microscopy. Prorocentrum porosum
sp. nov. was characterized by 35.9–50.2 μm long and 25.4–45.7 μm deep cells, covered by broadly ovoid
symmetric thecal plates. The surface of both thecal plates is smooth and covered by randomly scattered kidneyshaped pores (n = 102–149), rounder towards the center, absent in the central part, and surrounded by a
conspicuous marginal ring of about 69–92 evenly spaced pores. Broad V-shaped periflagellar area exhibiting
flagellar and accessory pores. The molecular phylogenetic position of P. porosum sp. nov. was inferred using
partial LSU rRNA gene (rDNA) and rDNA ITS sequences. This new species branched with high support in a
Prorocentrum clade including P. caipirignum, P. hoffmannianum and P. cf. lima (P. lima morphotype 5 sensu Zhang
et al., 2015). Pairwise comparison of ITS1 and ITS2 transcripts with these closest relatives revealed the presence
of compensatory base changes (CBCs), with the exception of P. cf. lima (P. lima morphotype 5), which only
showed in ITS2 a hemi-CBC (HCBC) and two base changes that possibly induce a structural modification. Toxin
analyses performed in two Colombian and Brazilian strains in the present study detected the presence of low
amounts of okadaic acid.
1. Introduction
(Steidinger and Tangen, 1996; Faust et al., 1999), including planktic
and/or epibenthic species inhabiting living (macroalgae, mangrove
roots, seagrasses) or inert substrates (e.g. dead coral, sediments, mollusk
Prorocentrum is a predominantly marine dinoflagellate genus
* Corresponding author at: Universidad Nacional de Colombia sede Caribe-Colciencias-Invemar, Santa Marta, Colombia.
E-mail address: earteagas@unal.edu.co (E. Arteaga-Sogamoso).
https://doi.org/10.1016/j.hal.2022.102356
Received 16 June 2022; Received in revised form 4 November 2022; Accepted 14 November 2022
Available online 7 December 2022
1568-9883/© 2022 Elsevier B.V. All rights reserved.
E. Arteaga-Sogamoso et al.
Harmful Algae 121 (2023) 102356
shells, sandy bottoms, submerged or floating detrital aggregates) (Fraga
et al., 2012; Hoppenrath et al., 2013). Some Prorocentrum species are
causative of harmful algal blooms, and thirteen of them are currently
included in the IOC-UNESCO Taxonomic Reference List of Harmful
Micro Algae (Lundholm et al., 2009 onwards). These are generally
epibenthic species producing biotoxins such as okadaic acid (OA) and its
analogues, dinophysistoxins (DTXs), as well as borbotoxins, fast-acting
and hemolytic toxins (Nakajima et al., 1981; Jackson et al., 1993;
Ten-Hage et al., 2000b; Heredia-Tapia et al., 2002; Pearce et al., 2005;
Amar et al., 2018; Hoppenrath et al., 2013). Some of these compounds
have been implicated in diarrhetic shellfish poisoning (DSP) in humans
(Gayoso et al., 2002). Yet, it has been difficult to establish a clear link
between the presence of these organisms and the occurrence of DSP
episodes (Levasseur et al., 2003; Foden et al., 2005).
The genus Prorocentrum Ehrenberg was first erected by Ehrenberg
(1834) with Prorocentrum micans Ehrenberg as the type species (Till­
mann et al., 2019). It is characterized by laterally compressed cells,
lacking cingulum and sulcus, with round, oval, oblong, ovoid or
heart-shaped cells (Hoppenrath et al., 2013). Specimens are nearly
completely covered by two major thecal plates, presenting the insertion
of its two flagella towards the apical area and not in the ventral area as in
other dinoflagellates (Hoppenrath et al., 2013). The flagella emerge
through one of the two pores surrounded by a cluster made up of several
tiny platelets known as the periflagellar area (Hoppenrath et al., 2013).
Currently, 80 Prorocentrum species are taxonomically accepted accord­
ing to the AlgaeBase (Guiry and Guiry, 2022), though this genus has
been under continuous revision as new species are constantly described
(Herrera-Sepúlveda et al., 2015; Luo et al., 2017; Rodríguez et al., 2018;
Chomérat et al., 2019).
The morphological features often used in the identification of Pro­
rocentrum species are: cell shape including cell symmetry and size
(length and depth); presence or absence of an apical collar, shape and
morphology of the periflagellar area, noticeable in the right thecal plate;
presence of structures such as apical spines, plate depressions; as well as
the shape and distribution of pores (Balech, 1988; Fensome et al., 1993;
Steidinger and Tangen, 1996; Faust et al., 1999; Hoppenrath et al.,
2013). Despite its apparently simple cell morphology, the taxonomy of
Prorocentrum is complex given the plasticity in some of these features
allied to the existence of cryptic species. For such reason, a combination
of morphological and molecular tools (e.g. rDNA sequences, ITS rDNA
secondary structure), is desirable for the characterization of new isolates
at species level within this genus (Hoppenrath and Leander, 2008; Han
et al., 2016; Nascimento et al., 2017; Chomérat et al., 2019; Lim et al.,
2019).
Prorocentrum species are ubiquitous and abundant in tropical and
subtropical regions. The presence of Prorocentrum has been reported
since 1977 in the Colombian Caribbean Sea (Lozano-Duque et al., 2011),
and the number of species has continuously increased to a current total
of eleven species (Rodríguez et al., 2010; Mancera-Pineda et al., 2014;
Arbeláez et al., 2017; Arbeláez et al., 2020). The genus has also been
thoroughly investigated in Japanese Pacific waters, where six species
and two phylotypes have been reported (Fukuyo, 1981; Nishimura et al.,
2020a, 2020b). In Brazil, investigations on Prorocentrum specifically
focusing on benthic habitats have only started recently, with six species
reported so far (Nascimento et al., 2017; Moreira-González et al., 2019).
In the present study, a new marine benthic toxic species, Pro­
rocentrum porosum sp. nov., formerly assigned to Prorocentrum sp. type 2
(Nishimura et al., 2020a) is described based on morphological charac­
ters, phylogenetic analyses [large-subunit (LSU) rRNA gene (rDNA)
(LSU rDNA) and internal transcribed spacer regions (ITS1 and ITS2) and
5.8S rDNA (rDNA ITS)], as well as ITS1 and ITS2 secondary structure
reconstructions, from material collected in the tropical/subtropical re­
gions of the Atlantic (Colombian Caribbean Sea and Northeastern Brazil)
and the Pacific Ocean (Southern Japan). These results allowed differ­
entiating the new species from closely related ones like P. caipirignum
and P. hoffmannianum. Comparisons among genetic distances of the
rDNA sequences from P. porosum and those from closely related spe­
cies/phylotypes support P. porosum as a different entity from the others.
2. Materials and methods
2.1. Sites of isolation and culture conditions
In the Atlantic Ocean, Prorocentrum cells were obtained from mate­
rial collected during independent sampling cruises in Colombia and
Brazil. In Colombia, cells were associated with Thalassia testudinum
leaves collected at Tayrona National Natural Park (PNNT), Colombian
Caribbean (11◦ 19’16.4” N, 74◦ 07’36.5” W) on June, 2018 (Fig. 1A).
Marine seagrass, Thalassia testudinum leaves were obtained by snor­
keling at a depth of around 2 m, gently cut from their base to avoid
detachment of epiphytes. In Brazil, cells were collected using passive
samplers composed of rectangular pieces of fiberglass screen (10 × 15
cm; ~2 mm mesh size; 174 cm2 surface area), placed ~0.5 m above the
bottom with the aid of an anchor and a small buoy, as described in
Tester et al. (2014). Samplers were deployed by SCUBA diving and left
for 24 h on the metallic structure of two shipwrecks near the city of
Recife (Pernambuco State, Northeast Brazil): a tugboat named “Marte”
(8◦ 21’18.6’’ S, 34◦ 32’49.8’’ W; 26 m depth) and a landing ship tank
named “Gonçalo Coelho” (8◦ 20’60’’ S, 34◦ 32’24’’ W; 24 m depth) on
March 28, 2017 (Fig. 1B).
Both the T. testudinum leaves and the passive samplers were carefully
placed in hermetically sealed plastic bags (Ziploc®) with seawater. In
the laboratory, bags were shaken for two minutes to loosen the adhered
organisms, after which most of the leaves – or the fiberglass screen –
were manually removed. The seawater containing the released organ­
isms was passed through a 250 μm sieve to retain particle debris and
remaining plant material. The resulting filtrate was used to search for
and isolate single dinoflagellate cells in aliquots on excavated slides
under different inverted microscopes. Single cell isolation was carried
out manually using capillary pipettes adapted to a silicone hose to pick
up the organisms of interest. Once isolated, cells were transferred to
another excavated slide with a drop of sterile filtered seawater,
repeating the procedure several times until no contaminants were
observed. The isolated single cells were inoculated in 96-well micro­
plates containing K (Keller et al., 1987) or f/2 (Guillard and Morton,
2004) culture medium, both diluted by half and without addition of
silicates, prepared with sterilized seawater from the source area in
Colombia (salinity: 36) or Brazil (salinity: 32), respectively. Cells were
kept at a temperature of 25 ◦ C and a photoperiod of 12:12 h light/dark
using “cool-white” fluorescent lamps at a photon irradiance (PAR) of
about 90 µmol quanta m− 2 s− 1, as quantified by a radiometer (Bio­
spherical Instruments Inc. San Diego, CA, USA). When active growth was
observed, cells were successively transferred and kept into small glass
vials and Erlenmeyer flasks with increasing volume of culture medium,
under the same conditions described above. Three Prorocentrum mono­
clonal cultures were successfully established and used in this investi­
gation: strain INV MYZ0002, from PNNT (Colombia), and strains
LM-067 and LM-084, from “Gonçalo Coelho” and “Marte” shipwrecks
(Brazil), respectively.
In addition, fixed specimens of a currently lost strain previously re­
ported as Prorocentrum sp. type 2 (Nishimura et al., 2020a), were also
investigated for thecal morphology. Strain OUN248P, originated from
the Pacific Ocean, was isolated from a brown alga (Padina minor) at <3
m depth in Nakagusuku Bay, Uruma, Okinawa, Japan (26◦ 14’47” N,
127◦ 52’59” E) (Fig. 1C), on November 15, 2015 (sampling and culture
conditions were shown in Nishimura et al. (2020a) in Sections 2.1. and
2.5., respectively).
2.2. Light microscopy (LM) and scanning electron microscopy (SEM)
observations
For LM observations, cells of the Colombian Prorocentrum strain INV
2
E. Arteaga-Sogamoso et al.
Harmful Algae 121 (2023) 102356
Fig. 1. Sampling locations of Prorocentrum porosum strains in the present study. A) Bonito Gordo (Bahía Concha Bay), in the Tayrona National Natural Park (PNNT),
Colombia; B) “Gonçalo Coelho” and “Marte” shipwrecks, Brazil; C) Nakagusuku Bay, Uruma, Okinawa, Japan.
MYZ0002 were observed using a Leica DMLA light microscope (Leica
Microsystems GmbH, Wetzlar, Germany) equipped with UV epifluor­
escence system (CoolLED pE-300WHITE, Ex.: 365 nm), coupled to an
AxioCam HR3 photographic camera (Zeiss, Göttingen, Germany). The
nucleus was stained using SYBR Green (Molecular Probes, Eugene, OR,
USA) as described in a method adapted from Figueroa et al. (2010) as
follows: a 5 mL aliquot of culture was fixed with 0.5% paraformaldehyde
for 10 min and washed in PBS pH7.0 (Sigma–Aldrich, St. Louis, USA) by
centrifugation at 1200 × g during 10 min. Chlorophylls were extracted
by resuspending the pellet in 5 mL of cold methanol and then storing the
suspension overnight in the refrigerator. The cells were then washed
twice in PBS (pH 7.0) as described above, and the pellet was stained with
a 1:150 solution of SYBR green in PBS 0.01 M (pH7.4), observed in a
Leica DM LA epifluorescence microscope (Leica Microsystems GmbH,
Wetzlar, Germany) with blue excitation (Ex.: 450/490 nm) and photo­
graphed as mentioned above. In living cells, the autofluorescence of
chloroplasts was registered using the same microscope and imaging
equipment. For the Brazilian Prorocentrum strain LM-084, living,
Lugol-fixed (1%) and NaClO-washed (Steidinger and Tangen, 1996)
cells were observed at 100–400 × magnification under a Zeiss AxioVert
A1 light microscope coupled to a Zeiss AxioCam® ERc 5s digital camera.
For SEM observations, cells of the strain INV MYZ0002 were exam­
ined after treating 2 mL of the culture in exponential phase with 4%
Triton (20 min), and washing with distilled water through 10 μm
membrane filters (TCTP, Isopore, Ireland) to remove cell membranes.
Then, cells were fixed with formaldehyde 4% for 24 h at 5 ◦ C, before
being dehydrated by passing them (twice each step) through increasing
ethanol concentrations: 30, 50, 75, 90, 95 and 100%. Samples were
dried using a critical point dryer equipment (Baltec CPD030). After­
wards, filters containing the specimens were mounted in a stub with a
Leit carbon tab, and osmium tetroxide vapors were applied for 30 min.
Finally, specimens were coated with gold using K550X sputter coater
and observed with a JEOL JSM6700 F (JEOL, Tokyo, Japan) with an
acceleration voltage of 5 Kv, at the CACTI facilities (Universidade de
Vigo, Spain). The lugol-fixed cells of LM-084 were carefully transferred
by disposing small aliquots (2− 5 mL) of the culture onto Nuclepore
polycarbonate filters with 2 μm pores (Whatman ®), and washing ten
times with distilled water. Cells were suspended from the filter and
samples were finally mounted on a stub, dried at 36–40 ⁰C for 3 h and
sputter coated with gold palladium. Cells were observed using a JEOL®
JSM 6360-LV (JEOL, Tokyo, Japan) microscope adjusted to 10 Kv ac­
celeration voltage, at the CME-UFPR (Universidade Federal do Paraná,
Brazil).
Regarding the Japanese strain OUN248P, cells were harvested in
stationary phase as a pellet and stored at − 20 ◦ C for around five years.
Then cells were fixed using glutaraldehyde 2.5% in final concentration:
0.1 mL of 25% glutaraldehyde; 0.9 mL of metals mix SWII medium
(Matsuda et al., 1996; Nishimura et al., 2020a). SEM preparations fol­
lowed the method described in Nishimura et al. (2020b), and the spec­
imens were observed at the CACTI facilities as above.
Cell size measurements were obtained from SEM images using
Irfanview 4.60 freeware for strain OUN248P, ZEISS ZEN lite freeware
for strain INV MYZ0002, and Inkscape freeware for strain LM-084.
The terminology proposed by Hoppenrath et al. (2013) was followed
to describe the cell orientation, and for the description, arrangement and
enumeration of the periflagellar platelets. Therefore, we use the term
anterior or "apical" for the site where the flagella emerge and "antapical"
or posterior for the opposite end, with the right thecal plate being the
most excavated anteriorly relative to the left one (both connected to
each other through a sagittal suture).
2.3. DNA extraction, PCR amplification and sequencing
Three to five cells of strain INV MYZ0002 were isolated by micro­
pipetting, then washed and stored for 24 h at − 40 ◦ C in a 200 μL tube.
Later, samples were cold shocked with liquid nitrogen for 1 min, and
finally heated (94 ◦ C, 1 min) in an Eppendorf Mastercycler EP5345
(Eppendorf AG, New York, USA).
The LSU rDNA D1–D3 regions were amplified using the pair of
primers D1R/LSUB (5´-ACCCGCTAATTTAAG-CATA-3´/5´-ACGAAC­
GATTTGCACGTCAG-3’) (Scholin et al., 1994). The full rDNA ITS was
amplified using ITSF01 (5’-AGGAAGGAGAAGTCGTAACAAGG-3’) (Ki
and
Han,
2007)
and
PERK-ITS-AS
(5’-CTTACTTA­
TATGCTTAAATTCAG-3’) (Kotob et al., 1999) primers. LSU D1–D3 and
rDNA ITS amplifications were performed by (PCR) in an Eppendorf
Mastercycler EP5345, using Horse-Power™ Taq DNA Polymerase Mas­
terMix 2x (Canvax, Spain) following manufacturer’s instructions. PCR
conditions for the LSU rDNA and rDNA ITS amplifications followed
those described in Sunesen et al. (2020). The PCR products of strain INV
MYZ0002 were purified with ExoSAP–IT Express (Applied Biosystems,
Foster City, CA, USA). Sequencing reactions were performed using the
Big Dye Terminator v.3.1 reaction cycle sequencing kit and migrated in a
SeqStudio genetic analyzer (both at Applied Biosystems, Foster City, CA,
USA) at the CACTI sequencing facilities (Universidade de Vigo).
For Brazilian strains LM-067 and LM-084, cells were centrifuged
(2332 × g) for 5 min and the supernatant replaced by ethanol (100%)
prior to DNA extraction. Later, cells were isolated with capillary
micropipette and washed six times with deionized water. A single cell
was placed in each PCR tube (four tubes per strain) containing 1–3 μL of
deionized water and stored at − 20 ◦ C. Cells of strains LM-067 and LM3
E. Arteaga-Sogamoso et al.
Harmful Algae 121 (2023) 102356
084 were sequenced following two consecutive PCR reactions (nested
PCR) to amplify the rDNA ITS and LSU D1–D3. For the first PCR reac­
tion, 2.5 µL of each primer - ITSfw (5′ -GTAGGTGAACCTGCGGAAGG-3ʹ;
Adam et al., 2000) and D3B (5′ -TCGGAGGGAACCAGCTACTA-3ʹ; Nunn
et al., 1996), 12.5 µL of the PCR Master Mix 2X reagent (Promega,
Madison®, WI, USA) containing polymerase, dNTPs, MgCl2 and the re­
action buffers, and 6.5 µL of nuclease-free water were added to each
tube. The PCR was performed in a Biometra TOne thermocycler (Ana­
lytik Jena) as follows: denaturation at 95 ◦ C for 2 min, then 35 cycles
composed of 30 s at 95 ◦ C, 1 min at 62 ◦ C (melting temperature; MT) and
1 min at 72 ◦ C, and a final elongation step of 5 min at 72 ◦ C. For the
second PCR reaction, 1 µL of the first reaction product was added to a
tube containing 2.5 µL of each primer – ITSfw and 28S364r
(5′ -CTCTCTTTTCAAAGTCCTTTTC-3′ ; Tibiriçá et al., 2020) for the rDNA
ITS; D1R (5′ -ACCCGCTGAATTTAAGCATA-3ʹ; Scholin et al., 1994) and
D3B for the LSU D1–D3, 12.5 µL of the GoTaq® G2 Hot Start Green
Master Mix reagent (Promega®, Madison, WI, USA) and 6.5 µL of
nuclease-free water. The second PCR was carried out as the first one,
changing the MT to 50 ◦ C for the rDNA ITS and to 56 ◦ C for the LSU
D1–D3 rDNA. Positive samples were purified and sequenced at
IFREMER (Concarneau, France) as described in Chomérat et al. (2019).
Folding and hybridization package’ (UNAFold) web server (Zuker,
2003). Nucleotides were annotated with probability (color code is re­
ported on the figure). The structure characterized by minimum free
energy was reported. The comparison was performed manually.
2.5. Toxin analyses by liquid chromatography – mass spectrometry
2.5.1. Sample preparation
2.5.1.1. Culturing and harvesting. Prior to toxin analysis, Prorocentrum
cells of strains INV MYZ0002 and LM-084 were grown in 500 mL or 200
mL of medium, respectively, under the culture conditions described in
Section 2.1 for each strain. Culture aliquots were taken for cell counts
and fixed with Lugol’s Iodine solution. Subsamples were collected from
each aliquot and cells were diluted with filtered seawater and enumer­
ated (average from 2 to 3 samples) using a Sedgwick-Rafter chamber.
After that, culture aliquots were harvested for toxin analysis at the early
exponential (“culture 1”: 9 days; 3067 cells mL− 1; n = 2) and late sta­
tionary culture phases (“culture 2”: 58 days; 51,744 cells mL− 1; n = 2)
for strain INV MYZ0002, and at early stationary phase (21 days; 29,050
cells mL− 1; n = 3) for strain LM-084.
2.4. Phylogenetic analyses and secondary structure predictions
2.5.1.2. Toxin extraction for the Colombian strain INV MYZ0002. Four
46 mL aliquots from cultures 1 and 2 at the growth phases already
described were harvested for toxin analysis and 3 mL aliquots were
taken for cell counts. An additional 25 mL sample was taken to check the
time required for completely breaking the cells during sonication. Sub­
aliquots of this sample were sonicated for 2, 4 or 6 min and the resulting
extracts were visually inspected under an inverted microscope (Eclipse
TE2000-S, Nikon Corporation, Tokyo, Japan) to determine the most
efficient sonication time.
Following a modified Quilliam et al. (1996) procedure, all four ali­
quots from each culture (corresponding to 141 × 103 cells for culture 1
and 2.38 × 106 cells for culture 2), were transferred to 50 mL conical
polypropylene tubes and centrifuged at 5240 × g for 10 min, at 10 ◦ C.
Then supernatants were carefully decanted without disturbing the cell
pellets. In order to determine the concentration of extracellular toxins,
each 46 mL supernatant aliquot was concentrated and purified by solid
phase extraction (SPE; Waters SPE-PAK tC18, 500 mg, 3cc), following
the procedure described by Riobó et al. (2013). The purified extracts
were filtered through 0.22 µm prior to analysis of extracellular toxins by
LC-HRMS.
Intracellular toxins from aliquots containing the cell pellets were
extracted following a modified version of Method 1 described in Quil­
liam et al. (1996). Cell pellets were transferred to Eppendorf tubes and
extracted first with 1 mL of 100% methanol and sonication for 6 min at
50 W while keeping the sample container in ice. This was followed by
centrifugation at 20,600 × g. Supernatants were recovered in individual
Eppendorf tubes and pellets were re-extracted with 1 mL of 100%
methanol and vortex mixed for 2 min. After centrifugation both super­
natants from each aliquot were combined (final volume 2000 µL). Each
cellular extract was subdivided into three subsamples. The first one
(750 µL) was filtered through 0.22 µm (PTFE syringe filter, Filter-Lab®,
Barcelona, Spain), prior to analysis by LC-HRMS. The second one (750
µL) was cleaned-up by SPE (see above), while the third subsample of
each extract (500 µL) was hydrolyzed (Villar-González et al., 2007) to
check for OA and DTXs esters and subsequently cleaned-up by SPE (see
above). Purified extracts were then filtered through 0.22 µm (PTFE sy­
ringe filter, Filter-Lab®, Barcelona, Spain), prior to LC-HRMS analysis.
The LSU D1–D3 and rDNA ITS sequences had final lengths of 811 and
602 bp, respectively for strain INV MYZ0002, 845 and 879 bp for the
LSU D1–D3 sequences of strains LM-067 and LM-084, and 797 and 761
bp for their corresponding rDNA ITS sequences. The alignment was
elaborated using MAFFT version 7 online (Katoh et al., 2019), with the
iterative refinement method L-INS-i. Sequences belonging to the most
similar species/phylotypes of Prorocentrum were selected. Final LSU
D1–D3 and rDNA ITS alignments included 62 and 54 sequences, with
992 and 922 positions in the final dataset, respectively. Phylogenetic
model selection for Maximum Likelihood (ML) method was performed
on MEGA 10.2.2 (Kumar et al., 2018). A Kimura-2 parameter (K2+G, γ
= 0.71) model was selected in LSU, same as in ITS phylogeny (K2+G, γ
= 0.95). Sequences from genera Karlodinium, Takayama and Scrippsiella
were used to root the LSU, while ITS tree was unrooted. Maximum
Likelihood phylogenetic analyses were conducted in MEGA 10.2.2.
Bootstrap values were estimated from 1000 replicates. The phylogenetic
relationships were also determined using Bayesian Inference (BI)
method and, in this case, the substitution models were obtained by
sampling across the entire GTR model space following the procedure
described in Mr. Bayes v3.2 manual. Bayesian trees were performed with
Mr. Bayes v3.2 (Huelsenbeck and Ronquist, 2001) and the program
parameters were statefreqpr = dirichlet (1,1,1,1), nst = mixed, rates =
gamma. The phylogenetic analyses involved two parallel analyses, each
with four chains. Starting trees for each chain were selected randomly
using the default values for the Mr. Bayes program. The corresponding
number of unique site patterns for the LSU D1–D3 and rDNA ITS
alignments were 305 and 295, respectively. The number of generations
used in these analyses was 1,000,000. Posterior probabilities were
calculated from every 100th tree sampled after log-likelihood stabili­
zation and 25% ‘‘burn-in’’ phase. Overall topologies by ML and BI
methods were very similar. The phylogenetic trees were represented
using the ML method with bootstrap values and posterior probabilities
from the BI. Uncorrected p-distances [proportion (p) of nucleotide sites
at which two sequences are different (Transitions + Transversions), i.e.
proportion of nucleotide sites that are different] were calculated for the
LSU D1–D3 and rDNA ITS2 sequences using MEGA 10.2.2. Thus, no
corrections for multiple substitutions at the same site, substitution rate
biases (e.g., differences in the transitional and transversional rates), or
differences in evolutionary rates among sites are considered (Nei and
Kumar, 2000).
ITS1 and ITS2 secondary structures were predicted using the Mfold
RNA Folding Form V2.3 implemented in the ‘Unified Nucleic Acid
2.5.1.3. Toxin extraction for the Brazilian strain LM-084. Toxin extrac­
tion for strain LM-084 followed the general procedures described in
Quilliam et al. (1996). A 20 mL culture aliquot, corresponding to 1.45 ×
106 cells, was transferred to a 50 mL conical polypropylene tube and
centrifuged for 10 min, at 2050 × g. Then, the supernatant was carefully
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E. Arteaga-Sogamoso et al.
Harmful Algae 121 (2023) 102356
collected without disturbing the cell pellet. The cell pellet was frozen at
− 20 ◦ C and then lyophilized prior to toxin extraction. Subsequently, 2
mL of 90% methanol were added and the cells were disrupted using a
sonic dismembrator (Cole Parmer CPX130, USA) at 105 W 80% for 5
min, followed by centrifugation at 1200 × g for 10 min and removal of
the supernatant. The procedure was repeated and the supernatant
fractions from both steps were combined and evaporated to dryness with
nitrogen gas at 40 ◦ C. The extract was suspended in 1 mL of 100%
methanol and filtered using 0.22 µm PTFE syringe filter (Analitica, São
Paulo, Brazil).
The following MRM precursor and product ions (m/z) were moni­
tored at optimized collision energy (CE) and collision cell entrance po­
tential conditions: 817 > 255.0 (CE: − 62.0 eV; CEP: − 40.6 V) and 817.5
> 113.0 (CE: − 82.0 eV; CEP: − 41.5 V) for dinophysistoxin-1 (DTX1),
803.5 > 255.0 (CE: − 64.0 eV; CEP: − 40.1 V) and 803.5 > 113.0 (CE:
− 82.0 eV; CEP: − 41.5 V) for OA and dinophysistoxin-2 (DTX2), in
negative ionization mode, 876.5 > 823.5 (CE: 39.0 eV; CEP: 56.5 V) and
876.5 > 213.0 (CE: 54.0 eV; CEP: 56.0 V) for PTX2, in positive mode.
Collision cell exit potential (CXP) was set at − 2 V and declustering po­
tential (DP) − 129 V for OA, DTX1 and DTX2, and at 8 V and 70 V for
PTX2, respectively.
Using the software Analyst®, OA concentrations were calculated
from calibration curves made of serial dilutions of the CRM-OAd reference material (National Research Council, Halifax, Canada).
The limits of detection (LOD), statistically calculated from repeated runs
(n = 4) of the calibration solution at the minimum measurable con­
centrations, were equivalent to 0.17 pg OA mL− 1, 1.8 pg DTX1 mL− 1,
0.81 pg DTX2 mL− 1 and 0.45 pg PTX2 mL− 1.
2.5.2. Instrumental analysis
2.5.2.1. Liquid chromatography-high resolution mass spectrometry (LCHRMS) for the Colombian strain. LC-HRMS analyses of the extracts from
strain INV MYZ0002 were performed using a Dionex High-Speed LC
coupled to an Exactive mass spectrometer, equipped with an Orbitrap
mass analyzer and an ESI probe for electrospray ionization (Thermo
Fisher Scientific, Waltham, MA, USA). The software used for MS analysis
was Xcalibur 4.1 (Thermo Fisher Scientific). Analyses were conducted
under alkaline conditions (Gerssen et al., 2009; van den Top et al.,
2011). The specific toxin analogues tested were OA, dinophysistoxin-1
(DTX1), dinophysistoxin-2 (DTX2) and pectenotoxin-2 (PTX2). The
column used for separations was a Gemini® NX-C18 110 Å (3 µm, 2.0
mm × 100 mm) (Phenomenex Inc., Torrance, CA, USA). Mobile phases,
gradient conditions and column temperature were as described in Reg­
ueiro et al. (2011). OA, DTX1 and DTX2 standards were from CIFGA
(Lugo, Spain). PTX2 standard was from National Research Council
Canada. Toxins were quantified by external standard calibrations.
The mass spectrometer was operated both in positive and negative
ESI (ESI+, ESI− ) with polarity switching. Source conditions were as
follows: spray voltage − 4000 V and +3700 V, capillary temperature
320 ◦ C, sheath gas 40 arbitrary units (au), and aux gas 0 au. The in­
strument was set in Full MS mode with the following parameters: scan
range 100 – 1000 m/z, mass resolution setting of 140,000, automatic
gain control (AGC) target of 3 × 106, maximum injection time of 200 ms.
For OA and DTX2 identification, the extracted ion chromatograms
within the 803–804 m/z range in negative mode were selected, while for
DTX1 and PTX2 the positive mode was chosen with extracted ion
chromatograms within the ranges 836–837 m/z and 876–877 m/z,
respectively.
3. Results
Prorocentrum porosum E. Arteaga-Sogamoso, F. Rodríguez, A. Amato,
B. Ben-Gigirey, C. Tibiriçá, N. Chomérat, L. Mafra, T. Nishimura, M. Adachi,
et J.E. Mancera-Pineda sp. nov.
Description: Prorocentrum porosum cells are symmetrical, broadly
ovoid, 35.9–50.2 μm long (L; mean 44.1 ± SD: 2.1, n = 114) and
25.4–45.7 μm deep (D; mean 38.2 ± 2.7, n = 114), and L/D = 1.02–1.41
(mean 1.16 ± 0.07, n = 114). The surfaces of both thecal plates and the
intercalary bands are smooth. Both thecal plates have a ring of 69–92
marginal pores (mean 77 ± 6.6, n = 24) and 102–149 thecal pores
(mean 128 ± 11, n = 25) randomly scattered except in the center. The
pores are rounded, elongated bordering the periflagellar area, but
mainly kidney-shaped in the rest of both thecal plates, 0.28–0.68 μm
long (L; mean 0.51 ± 0.10, n = 28) and 0.22–0.39 μm deep (D; mean
0.31 ± 0.04, n = 28). Wide V-shaped periflagellar area with no spines,
composed of eight platelets that leave place for a flagellar and an
accessory pore. One or more depressions are present on the platelets.
Photosynthetic cells with pyrenoids.
Type locality: Tayrona National Natural Park, Magdalena, Colombia,
Colombian Caribbean (11◦ 19´16.4’’ N; 74◦ 07´36.5’’ W, 2 m depth)
(Fig. 1).
Holotype: Specimen from strain INV MYZ0002 mounted on a SEM
stub deposited under the code (C-A-99709) at The Natural History
Museum of Denmark (Copenhagen) is indicated as the holotype.
Isotype: formalin-fixed (4%) material for strain INV MYZ0002 is
maintained at Marine Natural History Museum of Colombia (MHNMC)Makuriwa, from INVEMAR, Santa Marta, Colombia.
Molecular characterization: sequences of INV MYZ0002 were depos­
ited in GenBank under Acc. Nos. MW251881 (LSU rDNA) and
MW251880 (ITS rDNA).
Etymology: the species name refers to the large number of both thecal
and marginal pores that this organism presents, which differentiates it
from closely related species/phylotypes of the genus Prorocentrum.
Habitat: Prorocentrum porosum can be found in shallow to relatively
deep waters (2–26 m) with high transparency and salinity between 32
and 36, living as epiphyte on seagrass (Thalassia testudinum) leaves,
brown algae (Padina minor) or artificial substrates (metallic structure of
shipwrecks) in tropical/subtropical regions.
Morphology: symmetrical broadly ovoid cells in lateral view, with the
apical (or anterior) edge slightly tapering and the posterior rounded
(Figs. 2–5). The morphological and morphometric comparison between
the studied strains from the three locations is summarized in Table 1.
Length of cells, as measured from all strains, varied between 35.9 and
50.2 μm, (mean ± SD: 44.1 ± 2.1, n = 114), depth between 25.4 and
45.7 μm (38.2 ± 2.7, n = 114) and length/depth ratios between 1.02 and
1.41 (1.2 ± 0.1, n = 114). One central pyrenoid is remarkably visible in
2.5.2.2. Liquid chromatography-tandem mass spectrometry (LC-MS/MS)
for the Brazilian strain LM-084. Intracellular toxin contents of strain LM084 were determined as described in the harmonized protocol for
determination of lipophilic toxins of the European Reference Laboratory
for Monitoring of Marine Biotoxins (EURLMB, 2015) with adaptations.
Cell pellet extracts were analyzed by LC-MS/MS using an Agilent 1200
series (USA) LC system coupled to a 3200 AB Sciex® QTRAP triple
quadruple mass spectrometer (Applied Biosystems; USA) equipped with
a Turbo Spray interface. Chromatographic separations were achieved on
a Zorbax Eclipse Plus C18 column (50 mm × 4.6 mm I.D., 1.8 μm, 95 Å;
Agilent®) at 20 ◦ C, following the injection of 5 μL. Mobile phases were
composed of (A) 100% water and (B) 95% acetonitrile + 5% water, both
containing 2 mM ammonium formate and 50 mM formic acid (LC-MS
grade; Merck®). A gradient elution (0.3 mL min− 1) was applied as fol­
lows: 20% to 100% of B in 5 min, maintained at 100% B for another 4
min, and returned to the initial condition until the end of the analysis
(13 min in total). Toxins were detected using multiple reaction moni­
toring (MRM) with a turbo ion spray (ESI) source heated at 500 ◦ C.
During the MRM scanning, curtain gas (nitrogen) was set at 25 psi, the
nebulizer gas and auxiliary gas (air) at 40 psi, and the collision gas
(nitrogen) adjusted to the medium position. Ionization voltage and
entrance potential (EP) were 4500 V and 10 V, respectively, with a MRM
dwell time of 36 ms and a 5 ms pause between the mass range.
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E. Arteaga-Sogamoso et al.
Harmful Algae 121 (2023) 102356
right thecal plate. The connection between the periflagellar area and the
left theca has variable shape, from straight to curved (Fig. 6). Platelet 1
is triangular and excavated, with three to four depressions (Fig. 6). This
platelet connects with platelets 2, 6, 7 and externally with the accessory
pore. Platelet 2 is small and square to pentagonal and has one or two
depressions (Figs. 4F; 5C and 6), connecting with platelet 7. Platelet 2 is
also in contact with platelets 1, 3 and 8, and forms the left side of the
accessory pore. Platelet 3 is square to trapezoidal and excavated in four
depressions. It is large and in contact with platelets 2, 4, 5, and 8,
forming one edge of the flagellar pore. Platelet 4 has an elongated
triangular shape of variable length, with several depressions (between
one and four, usually three). It is in contact with platelets 3 and 5,
forming with the latter the ventral side of V-shaped periflagellar area
(Fig. 6). Platelet 5, characterized as elongated and thin, with a wide
curved arc shape, is in contact with platelets 3, 4, 6 and forms most of
the right side of the flagellar pore (Figs. 4G, 5C, 6). It generally presents
an elongated depression that gives it a channel-like appearance,
although in some cases it may exhibit depressions (Fig. 4F). Platelet 6 is
characteristically square and short, in contact with platelets 1, 5, 7, 8,
and with part of the flagellar and the accessory pores. It also forms a list
sometimes overlapping platelet 7 and the accessory pore (Figs. 4E, G; 5C
and 6). Platelet 7, which is internal and surrounds part of the accessory
pore, is adjacent to 1 and it is also in contact with platelet 2. Platelet 8
separates flagellar and accessory pores, forming a bridge connecting
platelet 6 in one side and platelets 2 and 3 on the other side (Fig. 6).
Phylogenetic relationships: The LSU D1–D3 phylogeny (Fig. 7) placed
P. porosum in a distinct branch, supported with high bootstrap and
probability values, together with former sequences of strains PL1-11
(labeled as P. lima; Murray et al., unpubl.) and OUN248P [Pro­
rocentrum sp. type 2; Nishimura et al. (2020a)]. The closely related
Prorocentrum species/phylotypes were P. caipirignum Fraga, Menezes &
Nascimento, P. cf. lima [P. lima morphotype 5 by Zhang et al. (2015)],
Prorocentrum sp. type 1 [Nishimura et al. (2020a)], and
P. hoffmannianum Faust.
Regarding the rDNA ITS phylogeny, the only sequences available for
P. porosum are those arising from the present study (Fig. 8). Overall, the
resulting topology was very similar to that of the LSU D1–D3, though a
higher resolution was obtained for the different Prorocentrum clades
considered. The pairwise comparisons of LSU and ITS2 (uncorrected p-
Fig. 2. Light and epifluorescence microscopy images of Colombian Pro­
rocentrum porosum strain INV MYZ0002. A) right side view of living cell. Scale
bar: 10 μm; B) epifluorescence image showing the pyrenoid (dark arrow) and
nucleus (white arrow). Scale bar: 20 μm; C) epifluorescence image of chloro­
plasts. Scale bar: 20 μm.
living cells but two overlapping ones could not be discarded (Fig. 2A, B).
The kidney-shaped nucleus located in the posterior region (Fig. 2B).
Cells have branched and reticulated chloroplasts surrounding the central
area (Fig. 2C). Both thecal plates have smooth surface with 102–149
pores (128 ± 11, n = 25). Pores were rounded or elongated around the
periflagellar area, but generally kidney-shaped in the rest of thecal
plates. They measured 0.28 and 0.68 μm in length (0.51 ± 0.1, n = 28)
and 0.22–0.39 μm in depth (0.31 ± 0.04, n = 28) (Figs. 3A, G, H; 4D, E;
5D), and were randomly distributed in most of the thecal surface, except
in the central area (Fig. 3B, C; 4A–C; 5A, B). The margins of the thecal
plate edges were completely surrounded by approximately 69–92 (77 ±
6.6, n = 24) evenly spaced marginal pores (Figs. 3A, D and E). A smooth
or horizontally striated intercalary band could be observed (Figs. 3A, D
and E), with variable thickness depending on the cell age (Fig. 3F). Wide
V-shaped periflagellar area frequently exhibiting collar (Fig. 2A, B; 3A;
4F-G; 5C). Thecal plates and platelets lacking structures like ridges,
protrusions, curved projections, wings or spines (Fig. 6A, B). Thick
flange surrounding the periflagellar area was not observed. Sometimes
platelet lists surrounded flagellar and accessory pores (e.g. platelets 1, 5
and 6).
Following the nomenclature proposed by Hoppenrath et al. (2013),
eight periflagellar platelets are observed in this species (Figs. 4F; 5C and
6). A schematic drawing is shown in Fig. 6C. Platelets 1 – 4 in contact to
the border of left thecal plate, with platelets 1, 4, 5 and 6 touching the
Fig. 3. Scanning electron microscopy images of Colombian P. porosum strain INV MYZ0002. A) apical view with the periflagellar area. Scale bar: 10 μm; B, C) left and
right side of cells. Scale bars: 10 μm; D, E) intercalary band. Scale bars: 10 μm; F) megacytic cell with wide intercalary bands. Scale bar: 10 μm; G, H) detail on pores,
left side of cell. Scale bars: 2 and 0.5 μm, respectively.
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E. Arteaga-Sogamoso et al.
Harmful Algae 121 (2023) 102356
Fig. 5. Scanning electron microscopy images of Japanese P. porosum strain
OUN-248P. A, B) right and left side of cells. Scale bars: 10 μm; C) periflagellar
area. Scale bar 1 μm; D) detail on pores, left side of cell. Scale bar: 0.5 μm.
Fig. 4. Scanning electron microscopy images of Brazilian P. porosum strain LM084. A, B) right side and C) left side of cells. Scale bars 10 µm; D, E) detail on
cell surface, intercalary bands and pores, right side of the cell. Scale bars: 2 µm;
F, G) Images of periflagellar area. Scale bars: 2 µm.
thermodynamic support. ITS2 secondary structures had good support for
all four helices.
Comparing ITS1 secondary structures of P. porosum and that of its
closest sibling P. cf. lima (P. lima morphotype 5) (Fig. 9A), two
compensatory base changes (CBCs) (one at the base of helix b C-G → UA; and one at the base of helix II C-G → G-C), three hemicompensatory
BC (HCBCs) (one at the base of helix II A-U → G•U; one at the tip of helix
II C-G → U•G; one at the base of helix III G•U → G-C) and four base
changes that produce a bond loss were recorded. The comparison be­
tween P. porosum ITS1 secondary structure with those of P. caipirignum
and P. hoffmannianum showed more pronounced differences. Namely,
the P. porosum vs P. caipirignum ITS1 secondary structure comparison
(Fig. 9B) revealed four CBCs (three at the base of helix II G-C→A-U, AU→G-C, C-G→G-G; one in helix III C-G→G-G), three HCBCs (one at the
base of helix II A-U→G•U; two in helix III G•U→ G-G, U-A→U•G) and
several other structural differences. Insertions were recorded as well
(reported as orange triangles in Fig. 9B). Noteworthy are the size and
structure of helices a and b which exhibited substantial structural and
compositional differences, although these helices are less supported. A
CBC analysis of the ITS1 secondary structures from P. porosum and
P. hoffmannianum (Fig. 9C) revealed two CBCs (one at the base of helix b
A-U→ U-A and one at the base of helix II C-G→G-G) and one HCBC (at
the base of helix II A-U→G•U). A UGUG insertion was also recorded at
the tip of helix b in P. porosum.
ITS2 secondary structures showed robust support (Fig. 9D–F). The
comparison among the ITS2 secondary structures from P. porosum, P. cf.
lima (P. lima morphotype 5), P. caipirignum and P. hoffmannianum
showed that P. cf. lima (P. lima morphotype 5) shared the same sec­
ondary structure with P. porosum with two exceptions, namely, a HCBC
on the 3’ side of the helix I (U•G → U-A), and two base changes possibly
inducing a structural modification. The first located at the tip of helix I
on the 5’ side (G•U → U•U). The other at the tip of helix IV on the 3’ side
(U-A → A). The latter base change induces a considerable variation in
the size of the bulge at the tip of the helix IV. The ITS2 secondary
distances) between P. porosum and its closest species/phylotypes are
described below.
Regarding LSU, the mean distances within groups for P. caipirignum,
P. cf. lima (P. lima morphotype 5) and P. hoffmannianum were 0.0061,
0.0021 and 0.0107, respectively. In turn, mean p-distances between
P. porosum and these species/phylotypes were higher than the former
ones (0.0124, 0.0113 and 0.0301, respectively), as well as that
regarding the single sequence of Prorocentrum sp. type 1 (0.0134). These
values were slightly more divergent than the corresponding compari­
sons calculated between P. caipirignum vs. P. cf. lima, P. hoffmannianum
and Prorocentrum sp. type 1 (0.0095, 0.0261, and 0.0188, respectively).
In the case of ITS2, within group mean distances were 0.0077 and
0.0163 for P. caipirignum and P. hoffmannianum. These values were
several times lower than p-distances between P. porosum and its closest
species/phylotypes [0.0879, 0.0665 and 0.0749, for P. caipirignum, P. cf.
lima (P. lima morphotype 5) and P. hoffmannianum, respectively],
somewhat lower than the corresponding values between P. caipirignum
vs. P. cf. lima and P. hoffmannianum (0.0785 and 0.0780, respectively).
ITS1 and ITS2 secondary structure predictions and comparison: The
rDNA ITS sequences of P. porosum strain INV MYZ002 (Ac. No:
MW251880), P. cf. lima (P. lima morphotype 5) strain DS4G4 (Ac. No:
KM266625), P. caipirignum strain UFBA (Ac. No: KY039500), and
P. hoffmannianum strain CCMP683 (Ac. No: KF885225) were used for
the prediction and comparison. Folding the P. porosum ITS1 region, a
five helix-secondary structure characterized by a ΔG = − 66.30 kcal/mol
was obtained (Fig. 9A–C). The ITS2 region secondary structure (ΔG =
− 50.40 kcal/mol) was composed of four helices, named helix I through
IV (Fig. 9D–F). Both structures are consistent with those predicted by
Gottschling and Plötner (2004) using 150 dinoflagellate sequences.
Helices I, II and III of the ITS1 are more robustly supported, while two
more helices (named a and b) received a weaker statistical and
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Harmful Algae 121 (2023) 102356
Table 1
Morphological and morphometric comparison between strains of Prorocentrum porosum from Colombia (INV MYZ0002), Brazil (LM-084) and Japan (OUN248P).
Strains
Cell shape
Cell size
Length
Depth
Length/Depth
Periflagellar area
Shape
Collar on left plate
Wing-shaped spine
Protrusions
Platelet list(s)
Platelet with depressions
No. of platelets
Flagellar pore
Accessory pore
Theca ornamentation
Thecal Pores
No. of pores
Round pores
Oblong pores
Ovoid
Kidney-shaped
Pores pattern
Plate center
Pores length
Pores width
Marginal pores
No. of pores
INV MYZ0002
LM-084
OUN248P
Total
Broadly ovoid
Broadly ovoid
Broadly ovoid - oval
Broadly ovoid - oval
40.2–50.2
33.8–45.7
1.02–1.29
40.1–46.8
33.5–40.4
1.02–1.37
35.9–43.5
25.4–35
1.15–1.41
35.9–50.2
25.4–45.7
1.02–1.41
Wide V-shaped
Sometimes
No
No
Yes
Yes, several in some platelets
8
Yes
Yes
Smooth
Wide V-shaped
Sometimes
No
No
Yes
Yes, several in some platelets
8
Yes
Yes
Smooth
Wide V-shaped
Sometimes
No
No
Yes
Yes, several in some platelets
8
Yes
Yes
Smooth
Wide V-shaped
Sometimes
No
No
Yes
Yes, several in some platelets
8
Yes
Yes
Smooth
114–149
Yes
Yes
No
Yes (mainly)
No, scattered
Devoid
0.55–0.68
0.27–0.39
112–139
Yes
Yes
No
Yes (mainly)
No, scattered
Devoid
0.52–0.57
0.32–0.52
102–120
Yes
Yes
No
Yes
No, scattered
Devoid
0.28–0.43
0.22–0.28
102–149
Yes
Yes
No
Yes
No, scattered
Devoid
0.28–0.68
0.22–0.39
70–92
71–74
69–70
69–92
resolution) in Brazilian strain LM-084 was 1.68 pg free OA cell− 1.
Interferences were also recorded during the analysis of the hydro­
lyzed raw intracellular extracts from strain INV MYZ002. Therefore, the
SPE clean-up procedure was also employed to allow the quantification of
total OA (sum of free OA and the hydrolyzed OA esters). Hydrolysis
revealed the presence of higher OA levels in the intracellular extract
with mean values (n = 4) of 2.66 ± 0.33 pg total OA cell− 1) for “culture
1” and 5.08 ± 1.09 pg total OA cell− 1 for “culture 2”. DTX2 and DTX1
were not detected in the hydrolyzed intracellular extracts either.
structures from P. porosum and P. caipirignum (Fig. 9E) besides a single
HCBC in helix I (U•G → U-A), showed a 8 bp-insertion in P. caipirignum
at the tip of helix I and structural as well as compositional differences in
helices III and IV. The comparison with P. hoffmannianum ITS2 sec­
ondary structure (Fig. 9F) showed two HCBCs in helix I (U•G → U-A and
C-G → U•G), a base change that induced a bond loss in P. porosum (G-C
→ A) and two indels. Also, in P. hoffmannianum, as in the case of
P. caipirignum, the structure and composition of helices III and IV are
substantially different compared to P. porosum.
Toxin profile and content: In LC-HRMS for the extracts of Colombian
P. porosum strain INV MYZ0002, extracted ion chromatograms from the
full-scan (m/z 100–1000) were selected as follows: in negative mode m/
z: 803–804 for OA and DTX2, in positive mode 836–837 m/z and
876–877 m/z, for DTX1 and PTX2 respectively. Retention times were
2.07 min for OA, 3.36 min for DTX2, 4.19 min for DTX1 and 8.21 min for
PTX2. OA and DTX2 are isomers and their spectra corresponded to [M H]− at 803.4560 m/z (Fig. 10). By the use of certified reference stan­
dards, we also checked the DTX1 and PTX2 spectrums. DTX1 spectrum
corresponded to [M + NH4]+ at 836.5146 m/z and [M + Na]+ at
841.4699 m/z. PTX2 spectrum corresponded to [M + NH4]+ at
876.5095 m/z and [M + Na]+ at 881.4647 m/z (data not shown).
Analysis of the four 46 mL extracellular (clarified medium) aliquots
from P. porosum strain INV MYZ0002 in “culture 1” revealed the pres­
ence of OA at a mean concentration of 8.76 ± 1.23 ng mL− 1 of extract
(equivalent to 2.86 ± 0.40 pg cell− 1), while in “culture 2” the mean
value was 9.65 ± 2.52 ng mL− 1 extract (0.19 ± 0.05 pg eq. cell− 1).
DTX2, DTX1 and PTX2 were not detected.
Similarly, DTX2, DTX1 and PTX2 were not detected in the P. porosum
raw intracellular extracts of the Brazilian and Colombian strains. How­
ever, during the analysis of the extracts from strain INV MYZ0002, in­
terferences around the OA retention time were detected. This prompted
us to implement the same SPE clean-up step as the one used for the
extracellular toxins, but purifying only 750 µL of extract. Further anal­
ysis of the SPE cleaned-up extract revealed the presence of OA in the
aliquots tested, yielding a mean value (n = 4) of 0.50 ± 0.09 pg free OA
cell− 1 for culture 1 and 0.73 ± 0.21 pg free OA cell− 1 (n = 4) for “culture
2”. The intracellular OA content measured by LC-MS/MS (low
4. Discussion
4.1. Morphological and morphometric comparison among P. porosum
strains
Cells of the Colombian P. porosum strain (INV MYZ0002) and Bra­
zilian (LM-084) strain exhibited a rounded or broadly ovoid shape,
while Japanese (strain OUN248P) cells tended to be more elliptical.
Additionally, cells of the Japanese strain were generally smaller, with
slightly fewer thecal and marginal pores (Table 1). Even though, the
numbers of thecal and marginal pores in cells of the Japanese strain
were still very high compared to those of other Prorocentrum species,
allowing their distinction, as discussed later. Regarding the character­
istics of the periflagellar area, including the number and arrangement of
platelets (Table 1), no differences were observed among the three
strains: all exhibited eight platelets (Table 1) some of them usually
presenting more than one depression, which could be an additional
feature assisting in species distinction.
4.2. Morphological and morphometric comparison between P. porosum
and other phylogenetically similar species
The diagnostic characters of the morphotypes of P. lima include the
surface morphology of the thecal plates and the periflagellar area, but
their variability has hampered the species identification within the socalled “Prorocentrum lima complex” (Nagahama et al., 2011; Nasci­
mento et al., 2016; Chomérat et al., 2019). As a result, this complex
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Harmful Algae 121 (2023) 102356
distinguishing P. porosum from P. lima and P. hoffmannianum, which
display round, oblong or ovoid pores (Table 2). However, this pore
shape is also present in P. caipirignum [Zhang et al. (2015) (as P. lima
morphotype 4 and P. cf. lima morphotype 5; Luo et al. (2017) (as P. cf.
maculosum); Nascimento et al. (2017)]. The depressions found in the
periflagellar platelets of P. porosum are also present in P. hoffmannianum
and P. caipirignum, but not explicitly reported in P. cf. lima [as P. lima
morphotype 5 in Zhang et al., (2015)] (Table 2), nor in P. lima (Naga­
hama et al., 2011).
The L/D values of P. porosum cells (1.02–1.41) are different from
those reported for P. cf. lima (P. lima morphotype 5), but are similar to
those of P. hoffmannianum, and overlap, total or partially the typical
values for P. caipirignum (Table 2). No L/D data were provided for P. lima
strain UNR-01 in Nascimento et al. (2017) but P. porosum displays
generally lower L/D ratios than those reported for other P. lima strains
(1.27–1.77; n = 25) in Nagahama et al. (2011). In fact, the strains in
Nagahama et al. (2011) belonged to P. lima complex [clades A and B, the
latter containing also UNR-01, as classified in Zhang et al. (2015)], and
included P. lima morphotypes 2 and 3, which are ovate and similar to the
epitype of P. lima (Nagahama et al., 2011), with a L/D ratio of 1.37 [after
Fig. 2A in Nagahama and Fukuyo (2005)].
The cell shape of P. porosum resembles that of P. arenarium, though
cells of the latter were originally described as round or slightly oval
(Faust, 1994). P. arenarium is considered by several authors as synonym
of P. lima (Grzebyk et al., 1998), but as a different species by Nascimento
et al. (2017), who would ascribe it to P. lima morphotype 1 sensu Zhang
et al. (2015). Moreover, thecal pores are typically round in P. arenarium,
which also exhibits fewer thecal and marginal pores [42–84 and 28–77
(Zhang et al., 2015); 65–73 and 50–57 (Faust 1994), respectively] than
P. porosum. Finally, the two species are clearly distant from each other
genetically (Figs. 7 and 8).
Phylogenetic analyses place P. cf. lima [P. lima morphotype 5 by
Zhang et al., (2015)] as the closest taxon relative to P. porosum. They
share several common features, such as smooth thecal plates; scattered
kidney-shaped pores (absent towards the center); a marginal ring of
large pores in both thecal plates; V-shaped periflagellar area, with a
conspicuous pyrenoid towards the center of the cell. However, in P. cf.
lima [P. lima morphotype 5 by Zhang et al., (2015)], cell shape is oblong
to oval, displaying L/D ratios much greater than P. porosum (Table 2).
The number of thecal and marginal pores is lower in P. cf. lima (P. lima
morphotype 5) and this appears to be the main difference in relation to
P. porosum (Table 2). In addition, cells of P. cf. lima (P. lima morphotype
5) tend to be smaller than those of P. porosum (Table 2).
In Chengue Bay, close to the study area in Colombia, Arbeláez et al.
(2017) reported the presence of P. lima-like cells, which were smaller (L:
40.4 ± 2.9 μm) and more elongated than those of P. porosum. The same
observation occurred with P. lima-like cells found together with
P. porosum in Bonito Gordo during the present study. In our case, the
markedly smaller P. lima-like cells (no genetic sequences available yet)
varied in length from 33.0 to 40.9 μm, with a L/D ratio between 1.2 and
1.5 (n = 38).
Fig. 6. Scanning electron microscopy images of periflagellar area in Colombian
P. porosum strain INV MYZ0002. A) numbering of platelets in the periflagellar
area; B) an example of morphological variation in the periflagellar area. Scale
bars: 2 μm. C) Schematic drawing of the periflagellar area in P. porosum.
includes several clades and morphotypes (e.g. Zhang et al., 2015;
Chomérat et al., 2019) but only two recognized species: P. caipirignum
and P. arenarium (and the latter only by some authors – see Nascimento
et al., 2017).
For comparative purposes, the morphological features of P. porosum
and the other species/phylotypes within the P. lima complex closely
related to P. porosum are shown in Table 2. Some of these features were
proposed by Hoppenrath et al. (2013) as the basis to circumscribe Pro­
rocentrum species. Overall, P. porosum mainly differs from closely related
species in the number of thecal and marginal pores, which do not
overlap with the typical ranges characterizing the compared taxa
(although this has not been determined for P. hoffmannianum, due to the
lack of sufficient information). In turn, some characteristics are shared
by all – or most – species/phylotypes analyzed (Table 2). These include
the wide V-shape of the periflagellar area; the smooth left and right
thecal plates [except in P. hoffmannianum, being reticulate-foveate near
the margins and smooth in the middle (Rodríguez et al., 2018)], the
absence of structures like wing-shaped spine, thick flange and pro­
trusions; and the presence of a theca covered by pores except towards
the central area, as well as accessory and flagellar pores, marginal pores,
pyrenoids and eight periflagellar platelets, except in P. caipirignum,
which has only 6–7 platelets according to Nascimento et al. (2017).
Noteworthy, the number of periflagellar platelets may vary in
P. caipirignum [see Luo et al., 2017 (designated as P. cf. maculosum)].
The presence of kidney-shaped thecal pores is another feature
4.3. Phylogenetic relationships
Phylogenetic relationships of Colombian and Brazilian strains of
P. porosum were inferred from two nuclear-encoded ribosomal RNA gene
markers (LSU rDNA D1–D3 and ITS regions) to address species delimi­
tation. These results confirm available phylogenies of Prorocentrum,
depicting P. porosum within the so-called “Prorocentrum clade 2” (Mur­
ray et al., 2009), which includes other benthic species (Figs. 7 and 8).
The most closely related species in both phylogenies were P. caipirignum,
P. hoffmannianum and two phylotypes reported as P. cf. lima – one
designated as P. lima morphotype 5 (sensu Zhang et al., 2015), and the
other as Prorocentrum sp. type 1 (Nishimura et al., 2020a). However,
only LSU rDNA D1–D2 sequence is available for the latter, precluding
any further assessment of ITS phylogeny or secondary structures in that
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Harmful Algae 121 (2023) 102356
Fig. 7. Phylogenetic tree inferred by Maximum Likelihood of LSU rDNA (D1–D3 regions), showing the relationships between Prorocentrum porosum sp. nov and other
closely related Prorocentrum species/phylotypes. Internal node supports are bootstrap values (Maximum likelihood) and posterior probabilities (Bayesian analyses).
Bootstrap values <60 and posterior probabilities <.6 are not shown (as hyphen). Asterisks indicate maximal support.
region.
Recently, Nishimura et al. (2020a) proposed a new phylotype, Pro­
rocentrum sp. type 2, including two strains, PL1-11 (not available) and
the Japanese strain OUN248P (a lost strain included in the present study
for morphological examination), which matched the clade of P. porosum.
The average p-distances calculated in the present work between strains
of P. porosum and its closely related species, P. caipirignum, were
equivalent to the range of the values obtained when the latter was
compared to other similar species/phylotypes in both trees. Regarding
the rDNA ITS, the p-distances between P. porosum and its closely related
species/phylotypes (0.066–0.088), are significantly larger than 0.04, a
boundary value that may be used to delineate most free-living dinofla­
gellate species (Litaker et al., 2007). These authors warned that recently
evolved species could display p-values below that limit, requiring
additional morphological and genetic data to resolve their position.
Nevertheless, this is not the case in P. porosum, in which rDNA ITS se­
quences can be easily obtained and used as species barcodes.
Given the lack of previous morphological information for Pro­
rocentrum sp. type 2 strains PL1-11 and OUN248P, their genetic simi­
larity left unclear in Nishimura et al. (2020a) whether they would
represent a new species. Previously, Chomérat et al. (2019) depicted
also a partial LSU rDNA-based phylogeny of Prorocentrum, including
P. lima complex, and ascribed every sequence according to the five
morphotypes designated by Zhang et al. (2015), except that of strain
PL1-11 which remained unlabeled at that opportunity. Recently, Nish­
imura et al. (2020a) designated P. lima complex clades 1–4 based on the
LSU D1–D3 sequences. The authors also ascribed P. lima morphotypes 1,
2, 3, 4, and 5 as P. lima complex clade 3, P. lima complex subclade 1a,
P. lima complex subclade 1b, P. caipirignum subclade c, and P. cf. lima,
respectively (Fig. 3 in Nishimura et al., 2020a). In the present study,
morphological and molecular characteristics of the Colombian, Brazilian
and Japanese strains were investigated and compared to those of the
closely related species/phylotypes, and these results supported the
erection of P. porosum as a new species.
4.4. CBC and HCBC analyses
The ITS2 secondary structure comparisons have been used in several
systems to set different species apart (Zhang et al., 2020). Since the first
report of ITS2 secondary structure for (biological) species boundary
identification (Coleman, 2000, 2003), the number of studies applying
secondary structure comparisons increased considerably, especially
when dealing with cryptic or pseudo-cryptic species (Coleman, 2007;
Amato et al., 2019). The secondary structures are usually analyzed by
comparing the stem (double-stranded) regions of the primary transcripts
(deduced from the genomic ITS1 or ITS2 regions). Two main variations
can be identified: compensatory base changes (CBCs) and hemi-CBCs
(HCBCs). The former occurs when two nucleotides facing each other
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Harmful Algae 121 (2023) 102356
Fig. 8. Phylogenetic tree inferred by maximum likelihood of ITS region sequences, showing the relationships between Prorocentrum porosum sp. nov. and other
closely related Prorocentrum species/phylotypes. Internal node supports are bootstrap values (Maximum Likelihood) and posterior probabilities (Bayesian Inference).
Bootstrap values <60 and posterior probabilities <0.6 are not shown (as hyphen). Asterisks indicate maximal support.
11
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Harmful Algae 121 (2023) 102356
Fig. 9. Prorocentrum porosum ITS1 (A–C) and ITS2 (D–F) secondary structure comparisons with P. cf. lima (P. lima morphotype 5) strain DS4G4 (A, D), P. caipirignum
(B, E), and P. hoffmannianum CCMP683 (C, F). The nucleotides are annotated in the probability of being in a double stranded area. The color code is reported on the
figure. Compensatory base changes (CBC) and Hemi-CBCs (HCBCs) are indicated with dark blue circles or ellipses. Insertions are indicated with orange triangles. In
light blue full circles, the nucleotides present in the compared structure at the same position. On the compared structure, smaller light blue full circles indicate the
changed nucleotide. Rectangles indicate homologous areas of the structure.
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Harmful Algae 121 (2023) 102356
Fig. 10. LC-HRMS toxin analyses of Colombian P. porosum strain INV MYZ0002. Snapshots of extracted ion chromatograms: A) m/z: 803–804 in negative polarity
and B) m/z: 805–806 in positive polarity, showing OA peak in the hydrolyzed cellular extract. OA spectrums: C) m/z: 803–804 in negative polarity and D) m/z:
822–823 positive polarity. The ammonium adduct [M + NH4]+ (m/z: 822.4975), together with the less intense [M + H]+ m/z: 805.4696 and the sodium adduct [M
+ Na]+ m/z: 827.4524 can be observed.
in a stem region (a helix) vary in such a way that the bond between them
is conserved. As an example, a CBC would be recorded if in one sequence
an A-U bond is present and in the compared sequence both nucleotides
change to produce a G-C bond. In turn, HCBCs occur when the nucleo­
tide change occurs only on one side of the helix. Noteworthy, in RNA
secondary structure, non-canonical base pairing can occur.
Non-canonical base pairing occurs when nucleotides pair following a
scheme other than that suggested by Watson and Crick. A comprehen­
sive analysis of ITS2 sequences in land plants (Müller et al., 2007)
estimated that, in the presence of a CBC, there is a ~93% probability
that two sequences belong to different species, while no CBCs indicate a
~76% probability of being the same species. The CBC species concept
and its correlation with biological species has been experimentally
validated in protists such as the marine pennate diatom genus Pseudo-­
nitzschia, for instance. For this diatom, entities bearing sequences with
no CBCs are potentially inter-fertile and allow gene flow (Amato et al.,
2007).
The biological systems benefitting from the implementation of ITS2
secondary structure comparisons include dinoflagellates (Leaw et al.,
2010, 2016), diatoms (Teng et al., 2015; Amato et al., 2007), green algae
and higher plants (Mai and Coleman, 1997; Caisová et al., 2013), co­
pepods (Di Capua et al., 2017), insects (Verma et al., 2020), tardigrads
(Schill et al., 2010), among others. Conversely, ITS1 secondary structure
has been far less explored, although it can be used to distinguish species
in some systems (Gottschling et al., 2001; Hoshina, 2010; Koetschan
et al., 2014; Ghosh et al., 2017), including dinoflagellates (Gottschling
and Plötner, 2004; Tornhill and Lord, 2010).
ITS2 secondary structure in dinoflagellates (e.g. Alexandrium, Coolia
and Prorocentrum), follows the canonical eukaryote “four domains” ar­
chitecture, except in Ostreopsis (Ramos et al. 2015). For species within
the genus Prorocentrum, only the recent descriptions of P. koreanum
(Han et al., 2016) and P. malayense (Lim et al., 2019) included secondary
structural information of ITS2 and CBCs. Both studies reported CBCs and
HCBCs (or at least a single CBC) between these two species and their
closest relatives. HCBCs can be observed among variants of the same
species, as described in the genus Coolia (Leaw et al., 2016), but the
probability of a CBC in variant ITS2 copies, at least in land plants, is
~99% (Wolf et al., 2013).
Prorocentrum porosum is supported as a different taxonomic entity in
relation to both P. caipirignum and P. hoffmannianum based on molecular
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Harmful Algae 121 (2023) 102356
Table 2
Morphological and morphometric comparison between P. porosum and other phylogenetically closely related Prorocentrum species/phylotype.
P. porosum sp. nov. (formerly P.
sp. type 2)1
P. cf. lima (P. lima morphotype
5)2
P. caipirignum3
P. caipirignum (as P. cf.
maculosum)4
P. hoffmannianum5
Cell shape
Symmetry
Cell size
Length (μm)
Depth (μm)
Length/Depth
Periflagellar area
Shape
Collar on left plate
Thick flange
Wing-shaped spine
Protrusions
Platelet list(s)
Broadly ovoid to oval
Symmetrical
Oblong to oval
Symmetrical
Elliptical
Symmetrical
Oval to ovoid
Symmetrical
Ovoid
Symmetrical
35.9–50.2
25.4–45.7
1.02–1.41
39.7–44.0
25.2–28.9
1.48–1.61
37–44
29–36
1.17–1.37
38.7–51.7
26.1–39.9
1.18–1.55
33.2–49.2
28.7–44.0
1.02–1.35
Wide V-shaped
Sometimes
No
No
No
Yes
Wide V-shaped
Yes (curved or flat)
No (according to photos)
No (according to photos)
No (according to photos)
Yes (according to photos)
Wide V-shaped
No (according to photos).
No (according to photos).
No (according to photos).
Yes (according to photos).
Wide V-shaped
Yes
Yes
No
No
Yes
Platelet with
depressions
No. of platelets
Flagellar pore
Accessory pore
Theca ornamentation
Thecal pores
No. pores
Round pores
Oblong pores
Ovoid
Kidney-shaped
Pores pattern
Yes, several in some platelets
Yes
8
Yes
Yes
Smooth
8
Yes
Yes
Smooth
Wide V-shaped
Sometimes
No
No
No
Yes, (according to
photos)
Yes, generally single per
platelet
6-7
Yes
Yes
Smooth
Yes, several in some
platelets
8
Yes
Yes
Smooth
Yes, several in some
platelets
8
Yes
Yes
Reticulate-foveate
102–149
Yes
Yes
No
Yes (mainly)
No, scattered
62–76
Yes
Yes
No
Yes
Many cases concentric
Devoid
0.28–0.68
0.22–039
Devoid
0.50–0.90
-
59–79
No
Yes
No
Yes
No, scattered (according
to photos)
Devoid
0.39–0.86
0.18–051
Yes
No
Yes
No
No, scattered
Plate center
Pores length (μm)
Pores width (μm)
Marginal pores
No. pores
Type pores
(according to photos)
56–78
No
Yes
No
Yes
No, scattered (according to
photos)
Devoid (according to photos)
-
69–92
Oblong? or kidney-shaped
Oblong or kidney-shaped
(according to photos)
51–69
Oblong or kidney-shaped
Oblong or kidney-shaped
(according to photos)
53–59
51–66
-
-
-
-
-
855–1013
Yes
Yes
No
No
-
-
-
-
0.75
-
-
-
-
0.40
Smooth or horiz.
Yes
Kidney
Posterior
Yes
-
Smooth
Yes
Posterior
Transv. Str.
Yes
Elongated
Posterior
Smooth
Yes
Oval
Posterior
Thecal depressions
No. depressions
Round depressions
Oblong depressions
Ovoid depressions
Kidney-shaped
depressions
Depressions length
(μm)
Depressions width
(μm)
Intercalary band
Pyrenoid
Nucleus form
Nucleus position
1
2
3
4
5
Devoid
0.39–075
-
Round to ovoid
This work (strains: INV MYZ002, LM-084 and OUN248P)
Zhang et al. (2015) (strain: SE10 and strains: AS4F8, DS4G4 and DS4D9)
Nascimento et al. (2017) (strains: LCA-B4, UFBA064)
Luo et al. (2017) (strains: TIO11, TIO102, TIO138, TIO179, TIO180)
Herrera-Sepúlveda et al. (2015) (strains: CCMP2804, CCMP683).
results (phylogeny and CBC analyses of ITS1 and ITS2 secondary
structures). However, it appears closely related to P. cf. lima (P. lima
morphotype 5) as demonstrated after the inspection of ITS secondary
structures. In this case, besides a reduced reliability of the folding, the
ITS2 structures presented one CBC and three HCBCs, while ITS1 only
three HCBCs (Fig. 9A). This weaker support could be interpreted as a
lack of resolution for CBCs as proxies for species boundaries in this
specific case, which would not allow separating P. cf. lima (P. lima
morphotype 5) from P. porosum based solely on this criterion.
In the case of Prorocentrum, its homothallic nature has been
confirmed recently in P. minimum (Berdieva et al., 2020). This obser­
vation, if applicable to other species such as those in the “P. lima
complex” would preclude the use of crossing experiments to identify
species boundaries and their correlation (or not) with CBCs.
Taking altogether, the results reported herein (morphological,
phylogenetic and ITS secondary structure analyses) support the erection
of P. porosum as a new species with the holotypic strain INV MYZ0002.
In our opinion, it is also highly likely that P. lima morphotype 5 deserves
a formal description as well, but this goes beyond the aims of the present
investigation.
4.5. Toxin profile and content
As reported for the Japanese P. porosum strain OUN248P in
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Harmful Algae 121 (2023) 102356
Nishimura et al. (2020a) our LC-HRMS analyses indicated that Colom­
bian and Brazilian strains of P. porosum were toxigenic (OA-producers),
making the proposed species potentially causative of DSP episodes or
other negative effects attributed to OA in these areas. However, whereas
only limited amounts of OA (<2 pg free OA cell− 1 and ≤5 pg total OA
cell− 1) were detected in the intracellular and extracellular extracts from
the Colombian and Brazilian strains, much higher OA levels (39.3 pg
free OA cell− 1) were reported in the Japanese strain (Nishimura et al.,
2020a). No detectable amounts of other diarrhetic shellfish toxins
(DSTs) (i.e., DTX1 and DTX2) and other lipophilic toxins (i.e., PTX1,
PTX2, PTX6 and YTX) have been reported in any P. porosum strains,
either by these authors or in the present study. In any case, the toxin
analysis results obtained in this work point to the production of OA in its
free and ester forms by P. porosum. This toxin has been previously iso­
lated from several Prorocentrum species, including P. lima (Murakami
et al., 1982), P. hoffmannianum (Aikman et al., 1993), P. concavum
(Dickey et al., 1990), P. arenarium (Ten-Hage et al., 2000a),
P. rhathymum (Caillaud et al., 2010), P. caipirignum (Nascimento et al.,
2017), P. cf. fukuyoi (Nishimura et al. 2020b), and P. foraminosum
(Kameneva et al., 2015) [re-assigned as P. aff. foraminosum by Chomérat
et al. (2019)].
Okadaic acid and its analogues are highly-specific inhibitors of
serine/threonine protein phosphatases PP1 and PP2A. Alterations in
DNA and cellular components, effects on immune, nervous system, and
embryonic development, and the potential role as a carcinogenic agent
have been reported (Prego-Faraldo et al., 2013; Valdiglesias et al., 2013;
Louzao et al., 2015; Amar et al., 2018). Our findings highlight the
importance of investigating DSTs and the related harmful algae species
(such as P. porosum) to prevent adverse health effects to humans and
marine organisms.
Distinct DST profiles are produced by different Prorocentrum species
(Lee et al., 2020) and even by different strains of the same species. For
P. lima strains, for instance, Bravo et al. (2001) reported the detection of
free OA, DTX1 and DTX2, together with OA and DTX2 esters, while
Rhodes et al. (2006) detected OA (free and esters) and low free DTX1
levels, and Kilcoyne et al. (2020) detected OA, DTX1 and their esters.
Similarly, Moreira-González et al. (2019) detected OA and DTX1 in
P. lima strains from Cuba and southern Brazil, but only OA in another
strain from northeastern Brazil. Okadaic acid was also the only DST
detected in two strains of P. caipirignum from Brazil (Nascimento et al.,
2017). In a much more comprehensive evaluation, Nishimura et al.
(2020a) reported that all 242 tested strains from the P. lima complex and
P. caipirignum produced OA at markedly varying levels, and some strains
of P. lima complex and P. caipirignum produced DTX1 as well, although
at very low levels in the case of P. caipirignum.
Rhodes et al. (2006) analyzed both free and total OA in extracts from
a P. lima strain over 34 days (stationary phase reached after 18 days).
They reported that algal cells initially released free OA into the culture
medium, but the proportion of free OA (both extracellular and cellular)
to OA esters (cellular) decreased as culture approached the stationary
phase. Rhodes et al. (2006) did not detect OA esters in the extracellular
medium. For P. porosum strain INV MYZ0002 in the present study, toxin
analyses were performed on cells at either early exponential phase,
when no cell lysis was observed, or stationary growth phase. We were
able to quantify intracellular (free and esters forms) and free extracel­
lular OA in all aliquots tested. Levels of OA in both raw and hydrolyzed
cellular extracts were higher at the late exponential phase. The opposite
happened in the extracellular medium.
In summary, based on morphological and molecular data, a new
benthic, OA-producing Prorocentrum species was characterized from
isolates obtained from the tropical/subtropical regions of the Atlantic
(Colombian Caribbean Sea and Northeast Brazil) and Pacific Oceans
(Southern Japan). Prorocentrum porosum is phylogenetically close to
other toxic species (P. caipirignum and P. hoffmannianum) and phylotypes
described in the literature (P. cf. lima = P. lima morphotype 5 and Pro­
rocentrum sp. type 1). Based on the few available LSU rDNA sequences,
P. porosum seems to be distributed in tropical/subtropical areas of both
Atlantic and Pacific basins. Its relative contribution within benthic
dinoflagellate assemblages is yet unknown and should be considered in
further studies.
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.
Acknowledgements
This study was supported by the Ministry of science, technology and
innovation of Colombia - MINCIENCIAS (COLCIENCIAS); the Uni­
versidad Nacional de Colombia-UNAL and CECIMAR (contribution
number: 547, within the project “Influencia de recursos y reguladores en
la abundancia de dinoflagelados bentónicos del Caribe”, Hermes Code:
40410); the Marine and Coastal Research Institute “Jose Benito Vives de
Andreis” (INVEMAR, contribution number: 1343) and Environmental
and Sustainable Development Ministry-MINAMBIENTE through BPIN
project; the International Atomic Energy Agency (IAEA) through the
Research Contract RC#18827 (“Bentox Project”) and the Technical
Cooperation projects RLA/7/014 and RLA/7/020; project DIANAS
(CTM2017-86066-R, MICINN), CCVIEO-10 (IEO, Spanish Institute of
Oceanography) and the Axencia Galega de Innovacion (agreement
GAIN-IEO). E.A.S. was supported by a COLCIENCIAS grant (call No.
727–2015). We thank I. Pazos and D. Cernadas for technical assistance
with SEM and S. Comesaña and V. Outeiriño for sequence analyses at
CACTI (Universidade de Vigo). We also thank P. Riobó for technical
support with MS operation, and the Center of Electron Microscopy at
UFPR (Brazil) for kindly providing lab supplies and making its equip­
ment available to this study. This study was also supported by the Crossministerial Strategic Innovation Promotion Program, Grants from the
Project of the NARO Bio-oriented Technology Research Advancement
Institution (the special scheme project on vitalizing management en­
tities of agriculture, forestry and fisheries), and Japan Society for the
Promotion of Science (JSPS) Bilateral Joint Research Projects (JSPSRSNZ Joint Research Project: JPJSBP120211002).
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