Apoptosis of hemocytes from lions-paw scallop Nodipecten subnodosus induced with paralyzing shellfish poison from Gymnodinium catenatum

Immunobiology 219 (2014) 964–974

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Apoptosis of hemocytes from lions-paw scallop Nodipecten subnodosus induced with paralyzing shellfish poison from Gymnodinium catenatum Norma Estrada a,∗ , Felipe Ascencio a , Liora Shoshani b , Rubén G. Contreras b,∗∗ a b

Centro de Investigaciones Biológicas del Noroeste, S.C. (CIBNOR), Calle IPN #195, La Paz, B.C.S. 23096, Mexico Centro de Investigación y de Estudios Avanzados del IPN (CINVESTAV), Av. Instituto Politécnico Nacional #2508, Mexico City, D.F. 07300, Mexico

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Article history: Received 25 January 2014 Received in revised form 5 May 2014 Accepted 15 July 2014 Available online 22 July 2014 Keywords: Apoptosis Gonyautoxin 2/3 epimers (GTX 2/3 epimers) Gymnodinium catenatum Hemocyte Nodipecten subnodosus Paralyzing shellfish poisons

a b s t r a c t The toxic dinoflagellate Gymnodinium catenatum produces paralyzing shellfish poisons (PSPs) that are consumed and accumulated by bivalves. Previously, we recorded a decrease in hemocytes 24 h after injection of PSPs (gonyautoxin 2/3 epimers, GTX2/3) in the adductor muscle in the lions-paw scallop Nodipecten subnodosus. In this work, qualitative and quantitative analyses, in in vivo and in vitro experiments, revealed that the lower count of hemocytes results from cells undergoing typical apoptosis when exposed to GTX 2/3 epimers. This includes visible morphological alterations of the cytoplasmic membrane, damage to the nuclear membrane, condensation of chromatin, DNA fragmentation, and release of DNA fragments into the cytoplasm. Induction of apoptosis was accompanied by phosphatidylserine exposure to the outer cell membrane and activation of cysteine-aspartic proteases, caspase 3 and caspase 8. Addition of an inhibitor of caspase to the medium suppressed activation in hemocytes exposed to the toxins, suggesting that cell death was induced by a caspase-dependent apoptotic pathway. The results are important for future investigation of the scallop’s immune system and should provide new insights into apoptotic processes in immune cells of scallops exposed to PSPs. © 2014 Elsevier GmbH. All rights reserved.

Introduction Gymnodinium catenatum (Graham, 1943; Pyrrophycophyta: Gymnodiniaceae) is a naked, chain-forming dinoflagellate that produces paralyzing shellfish poisons (PSPs). PSPs consist of more than 20 neurotoxic, hydrophilic, tetrahydropurine derivatives that are part of four subgroups: (1) carbamates (saxitoxin STX, neoSTX)

Abbreviations: Caspase, cysteine-aspartic proteases; DAPI, 4 ,6-diamidino-2phenylindole; DEVD-AFC, Asp-Glu-Val-Asp-7-amino-4-trifluoromethyl coumarin; DMSO, Dimethyl sulfoxide; DNA, Deoxyribonucleic acid; EDTA, Ethylenediaminetetraacetic acid; FITC, Fluorescein isothiocyanate; GTX, Gonyautoxin; GTX 2/3, Gonyautoxin 2 and Gonyautoxin 3 epimers; IETD-AFC, Ac-Ile-Glu-Thr-Asp-7amino-4-trifluoromethyl coumarin; MU, Mouse units; PI, Propidium iodide; PSM, Physiological saline media; PSP, Paralyzing shellfish poison; AU, Arbitrary units; SLS, Sodium lauryl sulfate; STX, Saxitoxin; STX eq, Saxitoxin equivalents; TBE, Tris-borate-EDTA buffer; Z-VAD-FMK, Carbobenzoxy-valyl-alanyl-aspartyl-[Omethyl]-fluoromethylketone. ∗ Corresponding author.Tel.: +52 612 123 8484. ∗∗ Corresponding author.Tel.: +52 55 5747 3800. E-mail addresses: [email protected] (N. Estrada), rcontrer@fisio.cinvestav.mx (R.G. Contreras). http://dx.doi.org/10.1016/j.imbio.2014.07.006 0171-2985/© 2014 Elsevier GmbH. All rights reserved.

and gonyautoxins (GTXs, GTX1–GTX4); (2) N-sulfo-carbamoyls (GTX5, GTX6, C1–C4); (3) decarbamoyls (dcSTX, dcneoSTX, and dcGTX1–dcGTX4); and (4) deoxydecarbamoyls (doSTX, doneoSTX, and doGTX1) (Mons et al., 1998; Quilliam et al., 2001). Commonly, G. catenatum shows a characteristic toxin profile, which consists of N-sulfo-carbamoyl toxins, such as B and C toxins as major components. When heated at low pH, toxins with this N-sulfo-carbamoyl moiety as a side chain, are partly converted to the corresponding carbamate toxins through hydrolysis (Mons et al., 1998). A large proportion of GTXs occur in cultures from Japan, Spain, and Tasmania (Oshima et al., 1993). GTX 2/3 epimers and its analogs bind to the pore of sensitive voltage-gated sodium channels that are expressed in excitable cells of vertebrates and block them. Excitable cells incubated with PSPs are incapable of increasing sodium conductance, generating action potentials, and producing nerve impulses, thereby becoming paralyzed (Mons et al., 1998). Some bivalves are resistant to PSPs because their sodium channels are insensitive to saxitoxins, demonstrated by whole-nerve assays (Bricelj et al., 2005); nothing is known about sodium channels in hemocytes of bivalves nor the effects or targets of these toxins and other biotoxins.

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Bivalve mollusk sometimes consume toxic dinoflagellates (Smayda, 1990; Bricelj and Shumway, 1998) and concentrate the toxins in their tissues. In turn, humans, birds, and other vertebrates consume mollusks and are affected by the toxins (Hallegraeff et al., 1995). Bivalves that are fed toxic algae or exposed to toxin extracts can detect the toxins and respond with a variety behavioral, tissue, and molecular changes (Hallegraeff et al., 1995; Landsberg, 1997). Molluskan hemocytes can mediate physiological responses environmental, biological, and disease stresses, inducing efficient immune responses, including chemotaxis, lectinmediated recognition, encapsulation, phagocytosis, and production of antimicrobial peptides (Girón-Pérez, 2010; Song et al., 2010). Hemocytes are not confined to the hemolymph system, but move freely out of the sinuses into surrounding connective tissue, the mantle cavity, and gut lumen; hence, these cells play important roles in physiological processes, such as gas exchange, osmoregulation, nutrient digestion and transport, excretion, and wound and shell repair (Cheng, 1981; Fisher, 1986). Some studies report that hemocytes has a lesser effect to exposure to biotoxins (Mello et al., 2012; Prado-Alvarez et al., 2012), but others report that bivalves, particularly their immune system, is adversely affected by toxic microalgae, with clear changes in hemato-immunological parameters (Hégaret and Wikfors, 2005a,b; Galimany et al., 2008a,b; da Silva et al., 2008; Estrada et al., 2010; Medhioub et al., 2013). Many marine microalgal toxins induce apoptosis in some vertebrate cell models (Boe et al., 1991; Leira et al., 1992; Lerga et al., 1999; Pérez-Gómez et al., 2004; Cabado et al., 2004; Lago et al., 2005; Xing et al., 2009). Despite decades of studies of mulluskan neurobiology and development, comparatively little is known about the mechanisms of apoptosis in this phylum. Although there are differences in apoptotic mechanisms between invertebrates and vertebrates, crucial biochemical components of the pathways of programmed cell death remain remarkably conserved throughout evolution (Kiss, 2010). Some major stimuli that induce apoptosis include pathogen infection, cell stress, and damage to DNA (Osborne, 1996; Opferman and Korsmeyer, 2003). The characteristic hallmarks of apoptosis in some mollusks include cell shrinkage and blebbing, chromatin condensation, DNA fragmentation, and phospholipid phosphatydilserine translocation into the outer leaflet of the cell membrane, as well as, induction of a family of caspases (cysteine-aspartic proteases) that cleave target proteins at specific sites (Sokolova, 2009; Earnshaw and Lazebnik, 1998; Cryns and Yuan, 1998). Apoptosis is an essential mechanism in the regulation of homeostasis and immune defense. It is a selective process of physiological cell deletion, as indicated by high baseline rates of apoptosis in circulating and resident hemocytes (Terahara and Takahashi, 2008; Sokolova, 2009). The lions-paw scallop Nodipecten subnodosus (Sowerby, 1835; Mollusca: Pectinidae) is a suspension-feeding bivalve mollusk that is widespread along the coast of the Baja California Peninsula to the southern coast of Peru (Keen, 1971). In the State of Baja California Sur, local shellfish aquaculturalists are interested in promoting intensive farming. Its economic value reflects its large size and weight (20 cm; 1000 g total wet mass), appealing flavor, and its large adductor muscle. Toxic algae impact scallop cultivation worldwide and provoke public health concerns regarding the safety of this resource. In a previous study, we demonstrated that injection of GTX 2/3 epimers from G. catenatum in the adductor muscle of N. subnodosus provokes paralysis and a decrease in hemocytes at 24 h, accompanied by negative responses in the scallop. This was determined by visible effects, generation of nitric oxide, peroxidation of lipids, and changes in antioxidant and hydrolytic enzymes in hemocytes and tissues (Estrada et al., 2010). In our present study, qualitative and quantitative analysis of in vivo and in vitro experiments revealed that many hemocytes died through a mechanism

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typical of cells undergoing apoptosis 24 h after injection of the toxin. Materials and methods General experimental design Firstly, we performed in vivo experiments by injecting GTX 2/3 epimers to confirm hemocyte death and for determining if toxins lead to apoptotic death. Secondly, we confirmed and demonstrated that GTX 2/3 epimers provoke apoptosis in hemocytes in vitro tests. We used several techniques to highlight apoptotic events when hemocytes were exposed to GTX 2/3 epimers (histological observations, structural alterations with cell death, exposure of phosphatidylserine on the outer leaflet of the plasma membrane, damage to DNA, and activation of caspases). Algal culture and source of scallops The dinoflagellate Gymnodinium catenatum (strain GCQM2) was isolated from vegetative cells collected in Bahía de Mazatlán, Mexico and deposited in the collection of the Marine Dinoflagellate Collection (CODIMAR; http://www.cibnor.gob.mx/ es/investigacion/colecciones-biologicas/codimar). The cells were cultured in GSe medium (Blackburn et al., 1989) with filtered (0.45 ␮m) seawater and grown in monoalgal cultures in 20 L glass flasks for a 16-h light:8-h dark photocycle at 21 ◦ C under 70 W fluorescent lamps. During acclimation (14–21 days), scallops were fed a mixture of microalgae (Chaetoceros calcitrans, C. muelleri, and Isochrysis galbana; 1:1:1). C. calcitrans (CHCAL-7) and C. muelleri (CHM-8) were cultivated in 20 L plastic bags in F/2 growth medium at 22 ◦ C under constant illumination at salinity of 32 psu. I. galbana (ISG-1) was grown in MA-F/2 medium under the same conditions. Cultures of G. catenatum were harvested in the late exponential growth phase and the others in the stationary growth phase. Juvenile scallops cultivated in suspended cages at Rancho Bueno, Mexico (24◦ 32 N, 111◦ 42 W) were collected and transported to CIBNOR. The specimens were placed in 40 L plastic tanks containing filtered (1 ␮m) seawater (35 psu) pumped from the lagoon. The water was maintained at 22 ◦ C with constant aeration through air stones. The water was replaced every 2 days. Extraction, identification, and quantification of toxins Cells of G. catenatum were harvested at the late exponential growth phase. Briefly, 50 L of G. catenatum culture were harvested for analysis by extracting the biomass by centrifugation at 3000 × g, suspended in 30–50 mL 0.1 N HCl and homogenized with glass beads (5 mm dia). Examination under an optical microscope of the cell debris, after homogenization, showed that the cells were completely disrupted. The starting material was a largescale laboratory culture of G. catenatum, and its principal toxin was N-sulfo-carbamoyl (C1, C2), as previously described (Estrada et al., 2007; Escobedo-Lozano et al., 2012). As described by Laycock et al. (1994), these N-sulfo-carbamoyl toxins were extracted and chemically converted to their analog carbamates, a mixture of GTX 2/3 (∼3:1). The cell extract was heated to 85 ◦ C for 15 min. After cooling the vial to room temperature, the pH was adjusted to 3.4–3.6. The extract was centrifuged for 10 min at 2000 × g and filtered with a single-use syringe filter (0.45 ␮m). After filtration, the supernatant was mixed with activated charcoal (1 g 10 mL−1 sample), agitated, and let stand for 24 h at 4 ◦ C. Activated charcoal with toxins were centrifuged at 2000 × g for 30 min and washed three times with deionized water. Then the toxins adsorbed on the charcoal were eluted with 200 mL 25% ethanol containing 1% acetic acid (1 g 10 mL−1 ). Many aliquots of toxin were made and

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the ethanol was removed by evaporation to dryness in vacuo. The dried toxin extracts were stored at −80 ◦ C. Dried toxin was dissolved in filtered 0.1 N HCl to verify identity of gonyautoxins epimers (GTX2 and GTX3) and subjected to chromatographic analysis, using the post-column oxidative fluorescence method (Franco and Fernández-Villa, 1993). Toxins were identified by reversed-phase chromatography using a 5 ␮m LiChrospher 100 RP-18 cartridge (12.5 cm × 4 mm id; #150477, EMD Millapore, Billerica, MA) at 33 ◦ C with a flow rate of 0.8 mL min−1 . An ion-pair buffer gradient, composed of a solution of 1.5 mM octansulfonic acid in 10 mM ammonia phosphate at pH 7, was used. The post-column reaction was performed in a teflon coil (10 m × 0.5 mm i.d.), held in a glass tube (at 65 ◦ C), and immersed in a water bath connected to a circulating temperature control unit. After post-column oxidation with 7 mM alkaline periodic acid and acidifying with 0.5 M acetic acid, the resulting products were detected with fluorescence detection set at 330 nm excitation and 390 nm emission. PSPs were identified and quantified by comparing chromatograms obtained from sample extracts with samples of certified toxin standards (PSP-1B; National Research Council Canada, Institute for Marine Biosciences, Certified Reference Material Program, Halifax, NS, Canada). Quantification of the toxin was based on comparing peak areas in chromatograms of sample extracts with the corresponding response factor. The biological activity was performed by mouse bioassay according to AOAC (1990) standards. Activity is expressed in mouse units (MU), where 1 MU is defined as the amount of toxin required to kill a 20 g mouse in 15 min after intraperitoneal injection, which is 0.18 ␮g saxitoxin equivalent (␮g STX eq). In vivo experiments – injecting GTX 2/3 epimers into adductor muscle Experimental design Before the experiments, specimens were placed in filtered (1 ␮m) seawater for 48 h to clear the gut. Adductor muscles of 12 live lions-paw scallops (shell height = 6.01 ± 2.61 cm, group 1) were injected with 0.2 mL of toxin at concentration of 140 ␮g STX eq 0.2 mL−1 in HCl 0.01 N with a 26 gauge needle attached to a 1 mL syringe. Another 12 scallops, used as controls (group 2), were injected with 0.2 mL HCl 0.01 N and had no adverse effects. After 24 h, the 12 scallops in group 1 were separated into four subgroups. Hemolymph of each subgroup of three scallops was drawn and pooled, and immediately put on ice; total hemolymph for each subgroup was ∼5 mL. The same was done for the controls. No scallop was subjected to more than one sampling. Samples were used for total and viability count, toxin analysis, and resin histological sections. To extract hemolymph, the valves were kept separated with a knife blade, and the hemolymph was withdrawn from the adductor muscle with a 26 gauge needle attached to a 1 mL syringe. Identification of GTX 2/3 epimers in hemolymph and hemocytes Two milliliter of hemolymph from each subgroup was used to identify the toxin; the same was done for the controls. Hemolymph was centrifuged at 1500 × g at 4 ◦ C for 10 min; the supernatant was lyophilized and re-suspended in 50 ␮L 0.1 N HCl. The hemocyte pellet was re-suspended in 50 ␮L 0.1 N HCl and sonicated for 10 min, then centrifuged at 5000 × g at 4 ◦ C for 10 min. Both extracts were then filtered with a single-use syringe filter (0.45 ␮m). The extracts were then subject to the chromatographic analysis. Hemocyte viability after injection of GTX 2/3 epimers Cell viability was measured by the propidium iodide (PI) exclusion method (Yeh et al., 1981), which is used as a DNA stain to determine cell cycle analysis. The ability of PI to enter a cell is dependent on the permeability of the membrane; it does not

stain live or early-stage apoptotic cells because the plasma membrane is intact. Pooled hemolymph (at a final concentration of 4 × 105 cells mL−1 is adjusted with hemolymph without hemocytes) was placed in a Neubauer chamber and observed under a phase-contrast microscope coupled with fluorescence for characterization. The number of cells in the central large square of the chamber was counted (cells mm−2 ). To obtain the number of cells in this square, the numbers of cells in each of the 25 medium-sized squares were counted and recorded as live or dead cells. Then the average of all measured squares was calculated, using two hematocytometer chambers per sample. Microscopy observation of hemocytes after injection of GTX 2/3 epimers Histological sectioning and contingency tables were performed for qualitative and semiquantitative analysis. For histology, 2 mL hemolymph of each of the four treated pooled subgroups and four control groups were mixed with Karnovsky’s fixative 1:1 (v:v) (Karnovsky, 1965) for 24 h at 4 ◦ C, then washed in several changes of Karnovsky’s buffer, dehydrated through an ethanol series, and embedded in catalyzed acrylic monomer (JB-4 plus embedding kit, Polysciences, Warrington, PA). Histological samples were sectioned from 0.5 to 1 ␮m. Histological sections were incubated with the lipophilic styryl dye (FM1-43, #T-3163, membrane, green; Invitrogen, Carlsbad, CA) for 15 min at room temperature, washed with physiological saline media (PSM) containing NaCl (20 g L−1 ), KCl (0.3 g L−1 ) CaCl2 (2.2 g L−1 ) and MgCl2 (1.9 g L−1 ) and then counterstained for 3 min with DAPI, blue (#10236276001, Roche Diagnostics, Basel, Switzerland), a fluorescent stain that binds strongly to A–T-rich regions in DNA. The slides were washed three times in the physiological saline medium for 5 min and mounted with mounting media (Vectashield #H-1000, Vector Laboratories, Burlingame, CA) and observed under an epifluorescence microscope. To understand the relationship between cytological responses to GTX 2/3 epimers, we made contingency tables with the following characteristics: (1) alterations in membrane structure, condensed chromatin with hyperchromasia, and fragmented DNA with release of DNA fragments into the cytoplasm; (2) positive or negative response; and (3) number of hemocytes of a given cytological type and response category. With these data, we used the 2 (chisquare) test (JMP 7, SAS Institute, Cary, NC). Data came from five random histological sections of each pooled subgroup, cut longitudinally with a difference of approximately 10–20 ␮m between sections (100 hemocytes were analyzed in each section). In vitro experiments with hemocytes exposed to GTX 2/3 epimers Dried GTX 2/3 epimers was diluted to a stock concentration of 1000 ␮g STX eq mL−1 in 0.01 N HCl. For the different experiments, we used three groups of five scallops (15 scallops, shell height = 5.34 ± 2.3 cm). Another three groups of five scallops were used as controls. To extract hemolymph, the valves of the scallops were kept separated with a knife blade, and the hemolymph was withdrawn from the adductor muscle with a 26 gauge needle attached to a 1 mL syringe. Hemolymph was extracted from five scallops and pooled in equal volumes in a total of three pooled subgroups. Hemocytes were counted with an electronic particle counter (Coulter Multisizer, Beckman Coulter, Brea, CA), to adjust cell concentration to densities of 1 × 106 cell mL−1 with hemolymph without hemocytes for the next experiments. Induction of apoptosis and hemocyte viability Each of the pooled subgroups of hemocytes were subdivided into three aliquots (each 100 ␮L at densities of 1 × 106 cell mL−1 ) for each treatment at each time interval and put on ice before the

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experiments. Cells were exposed in triplicate to staurosporine and camptothecin for 24 h at concentrations of 2.5 or 5 ␮M. The data were compared with GTX 2/3 epimers (10 and 20 ␮g STX eq mL−1 ) at 25 ◦ C. Control hemocytes were exposed to 0.01 N HCl (2 ␮L 0.01 N HCl 100 ␮L−1 culture). Campthotecin (#C-9911, Sigma–Aldrich, St. Louis, MO) was dissolved in dimethyl sulfoxide (DMSO) and made into a stock 1 mM solution. Staurosporine (#81590, Cayman Chemical, Ann Harbor, MI) was dissolved in DMSO and made into a 1 mM stock solution.The number of dead hemocytes was determined at 3, 6, 12, and 24 h at 25 ◦ C. Cell viability was measured with the PI exclusion method (Yeh et al., 1981), as described in Section Hemocyte viability after injection of GTX 2/3 epimers. Microscopic observation of hemocytes Histological sections and contingency tables were used to determine the alterations in cells in vitro. We used 3 mL of each of the pooled subgroups to perform histological sections using fluorescence microscopy. We tested staurosporine at concentration of 5 ␮M and GTX 2/3 epimers (20 ␮g STX eq mL−1 ) for 24 h at 25 ◦ C. Another pooled subgroup of hemolymph exposed to 0.01 N HCl (2 ␮L 0.01 N HCl 100 ␮L−1 culture) was used as a control. Pooled hemolymph was fixed, sectioned, and stained as described in Microscopy observation of hemocytes after injection of GTX 2/3 epimer. To study the relationship between cytological responses to GTX 2/3 epimers, we made contingency tables (explained in Microscopy observation of hemocytes after injection of GTX 2/3 epimer) with hemocytes exposed to GTX 2/3 epimers, compared with the controls. Data came from five random histological sections of each sample (pooled hemolymph of 15 scallops in three treated subgroups and three control subgroups), cut longitudinally with a difference of ∼10–20 ␮m between sections (100 hemocytes were analyzed in each section). Exposure of phosphatidylserine on the outer leaflet of the plasma membrane To identify the exposure of phosphatidylserine in the plasma membrane, we used the annexin V-fluorescein isothiocyanate (FITC) assay. Each of the pooled subgroups were subdivided into three aliquots (aliquots of 150 ␮L at densities of 1 × 106 cell mL−1 ) for each treatment. Cells were exposed in triplicate samples to staurosporine or camptothecin for 24 h at a concentration of 5 ␮M, and data was compared with GTX 2/3 epimers (20 ␮g STX eq mL−1 ) at 25 ◦ C. Control hemocytes were exposed to 0.01 N HCl (2 ␮L HCl 0.01 N 100 ␮L−1 culture). Cells were harvested (100 ␮L of each aliquot), washed with filtered (0.2 ␮m) physiological saline medium, resuspended in 100·␮L of the medium, and stained for annexin V exposure of the plasma membrane, using the annexin VFITC apoptosis detection kit (#K101, BD Biosciences Pharmingen, San Diego, CA). Exposure of phosphatidylserine on the outer leaflet of the plasma membrane is a surface change common to many apoptotic cells and is determined by annexin V-FITC staining. The hemocytes were spread in a Neubauer chamber and observed under a phase contrast microscope coupled with fluorescence for characterization. Viable cells, cells udergoing apoptosis and dead cells were analyzed at 20×. At least 100 cells were counted in each sample, five replicates for each treatment. Categories were assigned, based on the total number of hemocytes. Viable cells with intact membranes exclude PI, whereas the membranes of dead and damaged cells are permeable to PI. Cells viable are FITC, annexin V, and negative to PI. Cells that are in early apoptosis are FITC, annexin V positive and PI negative. Cells that are in late apoptosis or already dead are both FITC, annexin V, and PI positive. Breakage of double-stranded DNA Neutral comet assay was used to detect apoptosis. Aliquots of 150 ␮L cell suspension (1 × 106 cell mL−1 ) of each pooled subgroup

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were exposed in triplicate to four concentrations of camptothecin or staurosporine (2.5, 5, 10, and 20 ␮M) and two concentrations of GTX 2/3 epimers (10 or 20 ␮g STX eq mL−1 ) for 24 h at 25 ◦ C. Control hemocytes were exposed to 0.01 N HCl (2 ␮L HCl 0.01 N 100 ␮L−1 culture). To detect DNA fragmentation associated with apoptosis, we performed neutral comet assay (Fairbairn and O’Neill, 1996). This assay can detect various forms of breakage of DNA strands, dependent on the pH during electrophoresis (Collins, 2004). Under neutral conditions, it mainly detects breakage of double-stranded DNA and is therefore suitable for detection of apoptosis (Olive et al., 1991, 1993; Godard et al., 1999). Hemocytes at a concentration of 1 × 106 cell mL−1 were added to 0.75% low-temperature melting agarose at a ratio of 1:10 (v/v) and spread on glass slides that were pre-coated with 0.7% regular agarose and then air-dried. Slides with double-layered agarose were submerged in pre-cooled lysis solution (154 mM NaC1, 10 mM Tris, 10 mM EDTA, and 0.5% SLS at pH 10) at 4 ◦ C for 30 min, washed briefly to remove detergent and salt, and electrophoresed at approximately 7 V cm−1 for 3 min in TBE solution (40 mM Tris-boric acid, 2 mM EDTA at pH 8.3) and then stained for 10 min in PI (10 ␮g mL−1 ). DNA damage was quantified by measuring displacement between the genetic material of the nucleus (comet head) and the resulting tail; the intensity of the comet tail, relative to the head, reflects the number of DNA breaks. This was followed by visual analysis with staining of DNA and calculating fluorescence to determine the extent of the DNA damage. DNA damage was analyzed at 20× by fluorescence microscopy. Four random pictures per slide were taken (100 cells were counted in each slide). To rank each comet, we followed the microscopic scoring approach (Collins, 2004), assigning each comet to a class according to the degree of development of the tail (arbitrary unit scores and comet percentage). Conventionally, five classes are distinguished, from 0 to 4 (no visible tail to almost all DNA transferred to tail and indicating severe damage). A score based on visual assessment was computed for 400 comets per slide by giving each comet a value representing its class, so that the total ranges from 0 to 400 arbitrary units. Activation of caspase 3 and caspase 8 Activity of caspase 3- and caspase 8 were quantified by fluorometric detection of AFC (AFC; 7-amino-4-trifluoromethyl coumarin) after cleavage from DEVD-AFC (Asp-Glu-Val-Asp-7amino-4-trifluoromethyl coumarin) and IETD-AFC (Ac-Ile-Glu-ThrAsp-7-amino-4-trifluoromethyl coumarin), respectively, according to manufacturer’s instructions (K105, K112, BioVision, Milpitas, CA). Aliquots of 150 ␮L cell suspension (1 × 106 cell mL−1 ) of each pooled group were exposed in triplicate to camptothecin or staurosporine (5 ␮M) and GTX 2/3 epimers (20 ␮g STX eq mL−1 ) for 24 h at 25 ◦ C. Control hemocytes were exposed to 0.01 N HCl (2 ␮L HCl 0.01 N 100 ␮L−1 culture). Aliquots of hemolymph were placed in a microplate and hemocytes were lysed in the cell lysis buffer for 10 min on ice; then samples were incubated for 1 h at 37 ◦ C in the presence of the reaction buffer optimized for caspase activity assays, which is 10 mM dithiothreitol and 50 M DEVD-AFC for measurement of caspase 3-like activity or 50 M IETD-AFC for measurement of caspase 8-like activity. The samples were read by a fluorometer at max = 400 nm for excitation and 505 nm for emission and the amount of product cleaved under linear conditions was determined. Comparison of the fluorescence of AFC from an apoptotic sample with an un-induced control allows determination of the fold increase in caspase activity. To confirm the role of caspase activation, we treated hemocytes with the pan-caspase inhibitor Z-VAD-FMK (#G7231, Promega, Madision, WI) at three concentrations (0.8, 1.6, and 3.2 ␮M) for 1 h before adding either of the two commercial inductors of apoptosis (10 ␮M) or GTX 2/3 epimers (20 ␮g STX eq mL−1 ) for an additional 24 h. Z-VAD-FMK was dissolved in DMSO, aliquoted, and stored at 40 ◦ C. Cells were harvested

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and the assay for caspase 3 and caspase 8 were performed, as described earlier. Statistical analysis For all experiments, means and SD were calculated, and results are expressed as the means ± SD of three or four independent experiments. Student’s t-tests were performed to detect significant statistical differences between treated cells and control cells. The chi-square analysis (2 test) was used for histological observations. For caspase activity, we performed one-way ANOVA among compounds tested to induce activation if caspase. Statistical significance was set at P < 0.05. All analyses were performed with the JMP 7 statistical software (SAS Institute, Cary, NC). Results GTX 2/3 epimers concentration in hemolymph and hemocytes after injection Gymnodinium catenatum is composed of gonyautoxins (GTXs) epimers GTX2 and GTX3 (GTX 2/3 epimers) obtained after chemical hydrolysis and identification by reverse phase chromatography (Fig. 1B) and compared with standards (Fig. 1A). To investigate whether GTX 2/3 epimers in contact with hemocytes after injection and cause damage, we quantified GTX2/3 epimers. We found that GTX 2/3 epimers were present in hemolymph, but in hemocytes only GTX 3 was present (Fig. 1C).

Fig. 1. HPLC chromatograms showing fluorescence intensity of GTXs against retention time. (A) Toxin GTXs standard references. (B) GTX 2/3 epimers cell extract from Gymnodinium catenatum chromatogram. (C) Presence of GTX 2/3 epimers in the hemocytes of Nodipecten subnodosus 24 h after injection of GTX 2/3 in the adductor muscle.

volume and pyknotic-reduced nucleus (Fig. 2F). Most hemocytes in the control scallops had intact membranes and intact nuclei (Fig. 2B). Table 1 shows the statistical differences of the cytological features in hemocytes of scallops exposed to GTX 2/3 epimers, of which the most significant differences were condensed chromatin and fragmented nuclei.

Apoptotic hemocytes after injection of GTX 2/3 epimers in the adductor muscle

In vitro induction of apoptosis by incubation of hemocytes with GTX 2/3 epimers

We measured viability of the hemocytes after injection of GTX 2/3 epimers (140 ␮g STX eq 0.2 mL−1 ). Fig. 2A shows that GTX 2/3 epimers increased the number of dead hemocytes by ∼40%. Therefore, we investigated the cellular mechanism involved in hemocyte death induced by GTX. We observed histological sections of hemocytes with nuclei and membrane fluorescent staining. The membrane structure of cells were altered, hyperchromasia occurred with condensation of chromatin, nuclei were fragmented, and some chromatin fragments were released into the cytoplasm, all occurred concomitant with the apoptotic process (Fig. 2C–E). Additionally, there were some necrotic cells with a greater cell

To test whether in vivo and in vitro experiments had the same effects and determine whether the cause was apoptosis, we incubated hemocytes with GTX 2/3 epimers and measured hemocyte death and morphological alterations, and the potential induction of two very well-known inducers of apoptosis in vertebrate cells, staurosporine and camptothecin. Cell death was detected within 3 h after treatments; however, the kinetics differed slightly. Staurosporine induced cell death when used at concentrations as low as 2.5 ␮M. With GTX 2/3 epimers, the percentage of dead hemocytes were similar to death caused by camptothecin (Fig. 3A and B). Staurosporine and camptothecin produced condensation of chromatin

Fig. 2. In vivo hemocyte viability at 24 h after injection of GTX 2/3 epimers in the adductor muscle. (A) Percentage of live and dead hemocytes in control and GTX 2/3 epimers (140 ␮g STX eq 0.2 mL−1 ) treated cells. Results are expressed as the means ± SD of four independent experiments. (B) Fluorescence images from control hemocytes displaying nuclei (blue) and cell membranes (green) with normal morphology. (C) Hemocytes exposed to GTX 2/3 epimers, have chromatin condensation with hyperchromasia (arrow) and alteration in membrane structure (arrow head). (D) Chromatin condensation in blocks in the nuclei (arrows). (E) Fragmented chromatin was observed in the cytoplasm (arrows). (F) Some necrotic cells with increased cell volume and with a pyknotic-reduced nucleus. c = cytoplasm, n = nuclei. Scale bar = 2 ␮m. (For interpretation of the color information in this figure legend, the reader is referred to the web version of the article.)

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969

Table 1 Bivariate chi-square analysis (2 test) of cellular and nuclear variables observed in hemocytes obtained from control and GTX 2/3 epimers-injected scallops (140 ␮g STX eq 0.2 mL−1 ). Variable

Negative

Positive

n

(%)

n

(%)

Control Treated

1823 1103

91 55

177 897

9 45

Control Treated

1791 543

90 27

209 1457

10 73

Control Treated

1907 1598

95 80

93 402

5 20

Alteration in membrane structure

Condensed chromatin with hyperchromasia

Release of DNA to cytoplasm

in blocks or in the periphery of the nuclei, hyperchromasia and membrane alterations (Fig. 3F and G). Hemocytes treated with GTX 2/3 epimers at 10 and 20 ␮g STX eq mL−1 had damaged cell membranes, condensed chromatin, and hyperchromasia (Fig. 3D and E and Table 2). Hemocytes from the control sample maintained intact membranes and nuclei (Fig. 3C). There was no release of chromatin fragments into the cytoplasm, as observed during in vivo experiments when injected with GTX 2/3 epimers. Phosphatidylserine translocation to the extracellular leaflet of the plasma membrane Fig. 4 shows the percentage of: (1) viable or no measurable apoptotic cells, (2) early apoptotic cells with intact membrane, and (3)

2

P

660

<0.0001

1602

0.0000

724

<0.0001

cells in the last stage of apoptosis or dead. Counts were made 24 h after exposure to 20 ␮g STX eq mL−1 of GTX 2/3 epimers or 5 ␮M staurosporine or 5 ␮M camptothecin. GTX 2/3 epimers induced 31% apoptosis, staurosporine 55%, and camptothecin 45%. Under control conditions, the rate was 9%. Apoptosis and breakage of double-stranded DNA To measure damage and breakage of double-stranded DNA when hemocytes were subject to camptothecin and staurosporine at 2.5, 5, 10, and 20 ␮M and GTX 2/3 epimers at 10 and 20 ␮g STX eq mL−1 , we assayed the hemocytes with neutral, single cell gel electrophoresis after 24 h at 25 ◦ C. Breakage was quantified by measuring the electrophoretic displacement of nuclear

Fig. 3. In vitro hemocyte viability at 24 h after exposure of GTX 2/3 epimers and bona fide inductors of apoptosis. (A and B) Cell death of hemocytes, expressed as percentage of hemocytes exposed to: (A) 2.5 or 5 ␮M camptothecin or 2.5 or 5 ␮M stausosporine, (B) 10 or 20 ␮g STX eq mL−1 of GTX 2/3 epimers. Results are expressed as the means ± SD of three independent experiments. (C) Control hemocytes displaying nuclei (blue) and cell membranes (green) with normal morphology. (D and E) Hemocytes exposed 24 h to GTX 2/3 epimers (20 ␮g STX eq mL−1 ) with chromatin condensation and hyperchromasia (arrowhead) and chromatin condensation in blocks (arrows). (F and G) Hemocytes incubated with 5 ␮M of staurosporine with hyperchromasia (arrow head) and chromatin condensation in blocks (arrows). c = cytoplasm, n = nuclei. Scale bar = 2 ␮m. (For interpretation of the color information in this figure legend, the reader is referred to the web version of the article.)

970

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Table 2 Chi-square analysis (2 test) of GTX 2/3 epimers-induced effects (20 ␮g STX eq mL−1 ) in hemocytes exposed in vitro. Hemocytes were processed for histochemistry to determine alterations of cellular membranes and nuclei. Variable

Negative

Positive

n

(%)

n

(%)

Control Treated

977 794

98 79

23 206

2 21

Control Treated

917 129

92 13

83 871

8 87

Alteration in membrane structure

Condensed chromatin with hyperchromasia

genetic material resulting in a “tail”. Greater DNA breakage produces a longer tail (Fig. 5A) Fig. 5B shows that GTX 2/3 epimers, staurosporine, and camptothecin induce DNA damage in a concentration-dependent manner. In vitro activation of caspase in hemocytes caused by GTX 2/3 epimers To measure if activation of caspases induced death from GTX 2/3 epimers (20 ␮g STX eq mL−1 ), camptothecin (5 ␮M), and staurosporine (5 ␮M), we incubated hemocytes for 24 h at 35 ◦ C with these substances and examined caspase 3 and caspase 8 activity. Fig. 6A and B shows that GTX 2/3 epimers, camptothecin, and staurosporine activate caspase 3 and caspase 8, with higher amounts of caspase 3 than caspase 8. These are blocked by the pan-caspase inhibitor Z-VAD-FMK at 3.2 ␮M.

Fig. 4. In vitro phosphatidylserine translocation to the extracellular leaflet in hemocytes exposed to GTX 2/3 epimers at 24 h. (A) Hemocytes observed by phase contrast microscopy and fluorescence, to detect viable or no measurable apoptotic cells (green and red staining negative), apoptotic cells (green, anexin V-bound), and cells in end stage of apoptosis and dead (red, propidium iodide stained cells, and green anexin V-bound cells). (B) The graph shows percentages of different stages of cells of (A) after treatment with 5 ␮M staurosporine, 5 ␮M camptothecin, or GTX 2/3 epimers (20 ␮g STX eq mL−1 ). Control hemocytes were exposed to 0.01 N of HCl (2 ␮L HCl 0.01 N 100 ␮L−1 culture). Results are expressed as the means ± SD of three independent experiments. a = apoptotic, d = dead, v = viable. Scale bar = 6 ␮m. (For interpretation of the color information in this figure legend, the reader is referred to the web version of the article.)

2

P

165

<0.001

1245

<0.0001

Discussion By ingesting microalgae, bivalves are exposed to a variety of toxic components that may cause pathology and mortality. The effect of biotoxins in bivalves is of growing concern and many studies conclude that toxic dinoflagellates can significantly affect neurological, physiological, and behavioral responses of bivalves (Gainey and Shumway, 1988; Hallegraeff et al., 1995; Bricelj et al., 2005). Paralyzing shellfish poisons, such as GTX 2/3 epimers, bind to the pores of sensitive voltage-gated sodium channels, expressed in excitable cells of vertebrates, and block them (Mons et al., 1998). Some bivalves are resistant to PSP because they express saxitoxin-insensitive sodium channels in tissues, where resistance is caused by natural mutation of a single amino acid residue, which causes a 1000-fold decrease in affinity at the saxitoxin-binding site in the sodium channel pore (Bricelj et al., 2005). Hégaret et al. (2007) found that the dinoflagellate Alexandrium spp. do not exert a toxic effect in oysters hemocytes, which is consistent with the knowledge that PSP toxins interfere specifically with sodiumchannel function in neural tissues and supports the expectation that sodium-channel physiology has no importance in hemocyte functions in oysters. However, nothing is known about sodium channels in hemocytes of bivalves, neither the effects or targets of PSP toxins in these cells at the molecular level. Here, we quantified and identified the toxins in hemolymph and hemocytes 24 h after injection of the toxin in vivo, indicating that the toxin was in direct contact with lymph and hemocytes. Note that in hemocytes, the GTX-2 epimer is not present after 24 h on chromatograms, only a low quantity of the GTX-3 epimer was detected. GTX-2 is a sulfate ester of STX, whereas GTX-3 is the 11-epimer of GTX-2, and these two compounds are always found concurrently and form a 7:3 equilibrium mixture in a solution of neutral or higher pH (Shimuzu, 1988). Many factors could account for the absence of the GTX-2 epimer, such as an unknown differential elimination process, preferencial affinity of GTX-3 over the GTX-2, differences in distribution of toxins after exposure, and/or low content of the toxin in the sample. More studies must be done with different sampling times and different concentrations of toxins. Recent studies show that the immune system of bivalves is adversely affected by toxic microalgae, with a clear modulation of hemato-immunological parameters (Hégaret and Wikfors, 2005a,b; Galimany et al., 2008a,b; da Silva et al., 2008; Estrada et al., 2010; Medhioub et al., 2013). Other studies show that bivalve hemocytes remain unchanged from exposure to biotoxins (Mello et al., 2012; Prado-Alvarez et al., 2012). Still other works report apoptosis from exposure to toxic microalgae and their biotoxins (Galimany et al., 2008b; da Silva et al., 2008; Medhioub et al. 2013). Some biotoxins from microalgae induce apoptosis in some vertebrate cells (Boe et al., 1991; Leira et al., 1992; Lerga et al., 1999; Pérez-Gómez et al., 2004; Cabado et al., 2004; Lago et al., 2005; Xing et al., 2009). Previously, we showed that injection of GTX

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Fig. 5. In vitro DNA double strand breakage in hemocytes exposed to GTX 2/3 epimers (10 and 20 ␮g STX eq mL−1 ), and camptothecin and staurosporine (2.5, 5, 10, and 20 ␮M) for 24 h at 25 ◦ C. (A) Hemocyte nuclei showing images of representative grades of DNA damage, assessed by neutral comet assay, stained red with propidium iodide. DNA damage categories: undamaged, low damaged, medium damage, high damage, and complete damage, using scale of 0–4, respectively. Scale bar = 2 ␮m. (B) Frequency distribution of DNA damage in hemocytes. Control hemocytes were exposed to 0.01 N of HCl (2 ␮L HCl 0.01 N 100 ␮L−1 culture). Data were obtained from 400 scored nuclei. Results are expressed as the means ± SD of three independent experiments. Broken line represents damage level in control hemocytes. *P < 0.05.

2/3 epimers from G. catenatum into the adductor muscle of N. subnodosus provokes a decline in the concentration of hemocytes (Estrada et al., 2010). The scallops did not die when injected with a high concentration of toxin, but were permanently paralyzed. Here, we show through in vivo and in vitro qualitative and quantitative analysis that many hemocytes died by undergoing apoptosis when the scallops are injected in the adductor muscle or when hemocytes are exposed to GTX2/3 epimers. Death of hemocytes showed visible morphological alterations of the cytoplasmic membrane and damage to the nuclear membrane, condensation of chromatin, fragmentation of DNA, and release of DNA fragments into cytoplasm, all signs of apoptosis. To measure the direct damage to hemocytes by GTX 2/3 epimers, we performed in vitro experiments and determined the induction caused by two known inducers of apoptosis: staurosporine, a non-selective protein kinase inhibitor obtained from the bacteria Streptomyces staurospores (Kabir et al., 2002) and camptothecin, a potent inhibitor of topoisomerase I extracted from the Chinese tree Camptotheca acuminata (Hsiang et al., 1989). The results suggest that GTX 2/3 epimers and the two other compounds cause apoptosis. This was corroborated with the observed morphological alterations, breakage of double-stranded DNA, translocation of phosphatidylserine to the extracellular leaflet of the plasma membrane, and activation of caspases. All of these activities are critical steps in the sequence of events in apoptosis (Cohen, 1993; Mourdjeva et al., 2005). Apoptosis is an important mechanism for preserving a healthy and balanced immune system in vertebrates, where most cells have the ability to self-destruct by activation of a cellular suicide program when they are no longer needed or have become seriously damaged; however little is known about how apoptotic processes regulate immune defenses in invertebrates. The apoptotic effects were more prominent and clear than other kinds of damage at 24 h; other forms of cell damage were observed such as rupture and/or necrosis. Hemocytes of many species of

mollusks show apoptosis or necrosis when exposed to toxic compounds; in some cases, it was possible to demonstrate that the type of cell death depends on the dose of the toxin (Sokolova, 2009; Kiss, 2010). The basic molecular components of apoptotic cells that are expressed in many mollusks indicate that the apoptotic machinery is structurally and functionally highly conserved, although there is evidence that some apoptotic mechanisms in mollusks may differ from model invertebrates, more closely resembling what is seen in vertebrates (Sokolova, 2009). Recently, Medhioub et al. (2013) studied the apoptotic process in hemocytes in the Pacific oyster, Crassostrea gigas, when fed Alexandrium catenella, a PSP producer. In this case, oysters exposed for 48 h to the toxic dinoflagellate had a significant increase of the number of hemocytes in apoptosis after 29 h; two pro-apoptotic genes (Bax and Bax-like), implicated in the mitochondrial pathway were significantly up-regulated at 21 h, followed by overexpression of two effector caspase genes (caspase 3 and caspase 7) at 29 h. Here, the GTX 2/3 epimers of G. catenatum diminished cell viability and induced apoptosis in the hemocytes of N. subnodosus at 24 h. We suggest, based on our work and Medhioub et al. (2013), that GTX 2/3 epimers trigger intracellular responses that activate the intrinsic pathway after DNA damage, followed by induction of apoptotic events, such as caspase activation cascades, which leads to the characteristic morphological and biochemical features of apoptosis. It should be kept in mind that hundreds of proteins are part of an extremely fine-tuned regulatory network consisting of pro- and anti-apoptotic factors in hemocytes of N. subnodosus. Apoptosis can be induced in response to signals from inside (intrinsic) and outside (extrinsic) the cell. Apoptotic signals coming from the inside of the cell frequently have their origin within the nucleus, a consequence of DNA damage induced by irradiation, drugs, or other forms of stress. Activation of caspase is the main executor of the apoptotic process, which uses different stimuli in mollusks. GTX 2/3 epimers, camptothecin, and staurosporine

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Fig. 6. In vitro caspase 3 and caspase 8 activation in hemocytes exposed to GTX 2/3 epimers and bona fide inductors of apoptosis at 24 h. Caspase 3 activity (A) and caspase 8 activity (B) were determined with hemocytes using a fluorimetric assay and results were expressed as arbitrary units (AU). Dashed line represents levels of apoptosis in controls (untreated) hemocytes, and shaded bars represent hemocytes exposed to different compounds without caspase inhibitor. GTX 2/3 epimers-, camptothecin- or staurosporineinduced caspase activation in hemocytes can be blocked by pan caspase inhibitor Z-VAD-FMK. Results are expressed as the means ± SD of three independent experiments. Different letters denote statistically significant difference (P < 0.05) among compounds tested to induce caspase activation. Asterisk denotes significant differences (P < 0.05) with respect to hemocytes exposed to different inductors without caspase inhibitor (shaded bars) compared with hemocytes exposed to caspase inhibitors.

induced activation of caspase 3 and caspase 8 in hemocytes of N. subnodosus. Caspase 3 is an essential effector protein in the intrinsic pathway, whereas caspase 8 is an initiator protein of the extrinsic pathway; however, a “cross-talk” between the death-receptor (extrinsic) pathway and the mitochondrial (intrinsic) pathway commonly occur. The high level of caspase 3 activities, compared with caspase 8, could explain the relation with intrinsic pathway of GTX 2/3 epimers at 24 h. Caspases are activated by a variety of signals, including damage to DNA and stress (Sokolova, 2009; Romero et al., 2011; Lee et al. 2011). Caspase activity induced by GTX 2/3 epimers is blocked by the pan-caspase inhibitor Z-VAD-FMK, as occurs in other molluskan cell types (Lacoste et al., 2002). This suggests that hemocyte death was induced by a caspase-dependent apoptotic pathway. In summary, practically nothing is known about hemocytes and their capacity to internalize, inactivate, and eliminate GTX 2/3 toxins, nor the effects or targets of these toxins and other biotoxins in bivalve hemocytes at the molecular level. There is a complex relationship between damaged DNA and apoptosis, and we suggest that apoptotic signals result from damaged DNA

induced by toxins or stress, since nuclear alterations were the most conspicuous feature we observed. In turn, activation of caspases eventually leads to the characteristic morphological and biochemical features of apoptosis. Experiments indicate that GTX 2/3 epimers may tip the balance of cellular homeostasis towards increased mortality of hemocytes, and the reduced or abolished hemocyte function results in increased susceptibility to pathogens and other xenobiotics. The widespread distribution of paralyzing toxins along coasts and their increasing occurrence in edible bivalves dramatizes the need for toxicological studies to measure the potential risk to mollusks and their predators. Understanding the relationship between immune function of bivalves and toxic microalgae can contribute to more informed interpretation and management of consequences from biotoxin outbreaks in shellfish populations. Acknowledgments We thank Carmen Rodríguez and Eulalia Meza Chávez of the Laboratorio de Histología e Histoquímica of Centro de

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´ Investigaciones Biologicas del Noroeste, S.C. (CIBNOR) for technical assistance and Ira Fogel of CIBNOR for valuable editorial services. Financial support was provided by CIBNOR grant AC 3.0. N.A.E. is a recipient of a Consejo Nacional de Ciencia y Tecnología of Mexico fellowship (CONACYT 172583). References AOAC, 1990. Paralytic Shellfish Poison, Biological method, Final action, Sec. 959.08. In: Hellrich, K. (Ed.), Official Methods of Analysis. 15th Edition, AOAC, International, Gaithersburg, Maryland, pp. 881–882. Blackburn, S.I., Hallegraeff, G.M., Bolch, C.J.S., 1989. Vegetative reproduction and sexual life cycle of the toxic dinoflagellate Gymnodinium catenatum from Tasmania, Australia. J. Phycol. 25, 577–590. Boe, R., Gjersten, B.T., Vintermyr, O.K., Houge, G., Lanotte, M., Doskeland, S.O., 1991. The protein phosphatase inhibitor okadaic acid induces morphological changes typical of apoptosis in mammalian cells. Exp. Cell. Res. 195, 237–246. Bricelj, M.V., Shumway, E.S., 1998. 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